RESEARCHFree Access icon

Phytophthora spp. Associated with Appalachian Oak Forests and Waterways in Pennsylvania, with P. abietivora as a Pathogen of Five Native Woody Plant Species

    Affiliations
    Authors and Affiliations
    • Devin Bily1
    • Ekaterina Nikolaeva1
    • Tracey Olson1
    • Seogchan Kang2
    1. 1Bureau of Plant Industry, Pennsylvania Department of Agriculture, Harrisburg, PA 17110
    2. 2Department of Plant Pathology & Environmental Microbiology, Pennsylvania State University, University Park, PA 16802

    Published Online:https://doi.org/10.1094/PDIS-05-21-0976-RE

    Abstract

    To document the distribution of potentially harmful Phytophthora spp. within Pennsylvania, the Pennsylvania Department of Agriculture collected 89 plant, 137 soil, and 48 water samples from 64 forested sites during 2018 to 2020. In total, 231 Phytophthora strains were isolated using baiting assays and identified based on morphological characteristics and sequences of nuclear and mitochondrial loci. Twenty-one Phytophthora spp. in nine clades and one unidentified species were present. Phytophthora abietivora, a recently described clade 7a species, was recovered from diseased tissue of 10 native broadleaved plants and twice from soil from 12 locations. P. abietivora is most likely endemic to Pennsylvania based on pathogenicity tests on six native plant species, intraspecific genetic diversity, wide distribution, and recoveries from Abies Mill. and Tsuga Carrière plantations dating back to 1989. Cardinal temperatures and morphological traits are provided for this species. Other taxa, in decreasing order of frequency, include P. chlamydospora, P. plurivora, P. pini, P. cinnamomi, P. xcambivora, P. irrigata, P. gonapodyides, P. cactorum, P. pseudosyringae, P. hydropathica, P. stricta, P. xstagnum, P. caryae, P. intercalaris, P. ‘bitahaiensis’, P. heveae, P. citrophthora, P. macilentosa, P. cryptogea, and P. riparia. Twelve species were associated with diseased plant tissues. This survey documented 53 new plant-Phytophthora associations and expanded the known distribution of some species.

    The genus Phytophthora (Pythiaceae, Peronosporales, Oomycota, Chromista), including >150 described species and 140 to 540 estimated unknown species, threatens crops and ecologically significant plants across the globe (Brasier 2009; Jung et al. 2016b; Yang et al. 2017). In the past 2 decades, a surge of new taxa has been documented from natural ecosystems and tree plantations, including Phytophthora abietivora in 2019 as a root rot pathogen of Abies fraseri (Pursh) Poir. in Connecticut, U.S.A. (Jung et al. 2002, 2017a, b, 2020; Li et al. 2019; Mirabolfathy et al. 2001; Scanu et al. 2014). However, determining whether some species are endemic or introduced, as well as their full impact on nursery stock and plant ecosystems, can be difficult. In the United States, ∼69% of nonnative forest pests have been attributed to the live plant trade, exemplified by the widespread P. plurivora and P. cinnamomi (Beaulieu et al. 2017; Liebhold et al. 2012; Schoebel et al. 2014).

    Previous soil surveys in the United States of oak-forest ecosystems in eastern mid-Atlantic and central states detected P. cinnamomi, P. plurivora, P. xcambivora, P. pini, P. quercetorum, P. europaea, P. cactorum, P. cryptogea, and P. quercina, with P. cinnamomi as the most abundant species up to 40° latitude (Balci et al. 2007, 2010; McConnell and Balci 2014; Reed et al. 2019; Schwingle et al. 2007). Strains identified from natural waterways in eastern Atlantic and central states are dominated by species in clades 6, 9, and 10, including P. hydropathica, P. chlamydospora, P. irrigata, P. intercalaris, and P. lacustris (Brazee et al. 2016a; Hong et al. 2008, 2011, 2012; Hulvey et al. 2010; Hwang et al. 2009; Yang et al. 2014, 2016). The U.S. Department of Agriculture plant hardiness zones 6 and 7 harbor a diverse assemblage of Phytophthora, but because these taxa were recovered using traditional baiting methods, it is unclear which species cause disease on forest plants.

    The ecological role of Phytophthora in the health and fitness of broadleaved flora in natural environments of the eastern United States is poorly understood (Balci and Bienapfl 2013; Hansen 2015). P. cinnamomi is suspected of contributing to the decline of American chestnut before the arrival of chestnut blight, and is associated with declining Quercus alba L. in several mid-Atlantic states (McConnell and Balci 2014; Nagle et al. 2010; Westbrook et al. 2019). Hwang et al. (2009) isolated P. heveae and P. citricola sensu lato from symptomatic riparian Rhododendron L. and Kalmia latifolia L. in Appalachian forests of western North Carolina. Phytophthoras effect on plants largely depends on other abiotic and biotic factors including soil texture, climate, and antagonistic microorganisms (Corcobado et al. 2020; Kabrick et al. 2008; Moreira and Martins 2005; Roubtsova and Bostock 2009; Ruiz Gómez et al. 2019). Pathogenic species may lurk in soil with little detriment until their hosts are stressed or wounded, or when site conditions are conducive for disease progression, where they can cause aerial disease or fine root necrosis on forest plants (Balci and Bienapfl 2013; Davidson et al. 2005; Fichtner et al. 2009; Jung et al. 2018; McConnell and Balci 2014; Nechwatal et al. 2011).

    Pennsylvania’s forested land area totals 16.8 million acres (58% of the state’s total land area), with the oak/hickory forest type accounting for 53% of timberland (Albright 2017). Since 1975, the Pennsylvania Department of Agriculture has identified 36 Phytophthora spp. from nursery plants (Molnar et al. 2020). However, the distribution and impact of Phytophthora in native forests within Pennsylvania is not well known. To better understand the ecology of Phytophthora in forested environments and to confirm that regulated species including P. ramorum and P. kernoviae are not present, the Pennsylvania Department of Agriculture (PDA) conducted a 3-year survey sampling plants, soil, and water. Main findings from the survey are described here.

    Materials and Methods

    Field sampling.

    Field sampling took place during the growing season from May 2018 to October 2020 within Pennsylvania state parks, game lands, and national forests, at an elevation ranging between 65 and 650 m. The sampling sites were mostly dry-mesic Quercus L. and Carya Nutt. forest, but also included northern hardwood and mesophytic-type, with Acer L., Betula L., Prunus L., Fagus L., Liriodendron L., Tsuga, Pinus L., and Rhododendron. Plants were visually inspected for any symptoms of Phytophthora infection, and symptomatic tissue was placed in a 1-liter polyethylene bag kept inside a cooler until processed at PDA. Seedlings were defined as plants with a stem diameter <2.54 cm, saplings with a stem diameter between 2.54 and 12.7 cm, and mature plants with a stem diameter >12.7 cm. Stem diameters were measured using a standard diameter caliper (Forestry Suppliers Inc., Jackson, MS). Rhizosphere soil samples were collected under mature trees, and crown health was visually rated using the crown status criteria from Balci et al. (2007). Trees displaying a crown rating of 1 and 2 were considered healthy, while trees with a rating of 3 to 5 were considered in decline. After removing organic debris from around the tree, soil was sampled at cardinal directions to a depth of ∼30 cm in four subsamples until 4 liters of soil was collected as one sample, labeled, and placed in a cooler. Water was sampled from streams and lakes with forested watersheds and baited using the bait bag or bottle-of-bait (BOB) methods from the USDA APHIS PPQ (2014) water sampling protocol. Second-year, nonwounded, asymptomatic Rhododendron maximum leaves were used as bait.

    Sample processing.

    Water and soil were processed using modified isolation methods described in USDA APHIS PPQ (2014). After thoroughly mixing a soil sample, the pH was measured using an Accumet Research AR20 pH/conductivity meter (Thermo Fisher Scientific, Waltham, MA). Each soil sample was distributed into three 1,780 × 510 × 510 mm high-density polyethylene containers, where 1.3 liters of soil were deposited at a depth of 2 cm in each container and flooded with ∼4 liters of dH2O to a depth of 2.5 cm above the soil surface. After the soil settled, four first-year, nonwounded, asymptomatic R. maximum leaves were floated on the surface of the water. Baited leaves from water and soil samples were incubated at 18 to 25°C for 72 h, washed with dH2O, wrapped in paper towels moistened with dH2O, and incubated at 20°C for up to 10 days. Necrotic or water-soaked lesions were then punched out using a 6-mm hole puncher (Fiskars, Helsinki, Finland), surface-sterilized with 70% ethanol, then placed on CMA-PARP (17 g of corn meal agar in 1,000 ml of dH2O, amended with 0.4 ml of pimaricin [2.5% stock solution], 250 mg of ampicillin, 10 mg of rifampicin, and 5 ml of pentachloronitrobenzene [75% stock solution]; Jeffers and Martin 1986) and incubated for up to 5 days at 20°C. Symptomatic plant tissue was surface-sterilized with 70% ethanol, and sections between healthy and necrotic tissue were cut out with a sterile scalpel and placed on CMA-PARP. Single hyphal-tip cultures were derived from the edge of fresh cultures and transferred to V8 juice agar (Campbell’s Soup Co., Camden, NJ; V8a; 17 g of bacto agar, 200 ml of V8 juice, 800 ml of dH2O, and 2 g of CaCO3). After 5 days at 20°C, three 5-mm plugs from the edge of a colony were cut out, flooded with a 3% Non-Sterile Rhododendron Root Extract Solution (15 g of fine roots/500 ml of dH2O) modified from Jeffers and Aldwinckle (1987), and placed under fluorescent lighting for 24 to 48 h to induce the formation of sporangia.

    Morphological characteristics were observed to confirm that the strain was Phytophthora before DNA extraction. Strains identified as Pythium, Saprolegnia, or Mortierella were excluded. Morphological traits were photographed using a Nikon Eclipse Ti microscope and NIS Elements F software (v.4.60). Sporangia and gametangia were measured manually according to Dick (1990) at 1,000×. All strains are maintained in hemp seed water vials (three hemp seeds/10 ml sterile water).

    Colony growth patterns of P. abietivora were recorded from 15-day-old cultures (n = 12) grown at 20°C in the dark on malt extract agar (MEA), potato dextrose agar, corn meal agar (CMA), carrot agar (CA; 17 g of bacto agar, 200 ml of carrot juice, 800 of ml dH2O), hemp seed agar (HSA; 17 g of bacto agar, 50 g of hemp seeds, 1,000 ml of dH2O), and V8a. The temperature-growth relationship was determined using the four representative strains employed for phylogenetic analysis (three replicates each). A 5-mm plug from the edge of a 7-day-old culture was transferred onto the center of a 90-mm clarified V8a Petri plate for 24 h at 20°C to stimulate the onset of growth, a starting point was marked, and then each strain was transferred to 5, 10, 15, 20, 25, 30, and 35°C incubators. After 7 days, radial growth was measured along four perpendicular lines that intersected the center of the inoculum plug to calculate the mean growth. Additionally, three replicates of each strain were kept at 2 and 37°C for 7 days and returned to 20°C to check viability.

    Strains of P. cinnamomi and P. xcambivora were paired with compatibility-type testers in 90-mm, clarified V8a Petri plates. P. xcambivora strains were paired with A1 mating type P. cryptogea (ATCC 15402) and P. nicotianae (ATCC 15409), and A2 mating type P. cryptogea (ATCC 15403) and P. nicotianae (ATCC 15407). Strains of P. cinnamomi were paired with verified A1 (M106) and A2 (M23A) mating type P. xcambivora from previous Malus surveys in Pennsylvania. Strains were incubated at 20°C in the dark for 14 days, where oospore characteristics of each species were noted.

    Molecular identification and phylogenetic analysis.

    DNA was extracted from 235 strains from 1-week–old cultures on V8a. Mycelia were harvested with a sterile pin and transferred to a 1.5-ml tube containing an extraction buffer (100 mM of Tris-HCL, 10 mM of EDTA, and 1 M of KCl at pH 8), and macerated using a plastic mini-pestle. The supernatant was transferred to a new 1.5-ml tube after centrifugation. DNA was precipitated using isopropanol and washed with 70% ethanol and reconstituted in 50 μl of PCR-grade water. Targeted regions were amplified using the following primer sets; internal transcribed spacer (ITS; ITS4/ITS5; White et al. 1990), NADH dehydrogenase subunit 9 (NAD9; NAD9-F/NAD9-R; Martin et al. 2014), NADH dehydrogenase subunit 1 (NADH1; NADHF1/NADHR1; Kroon et al. 2004), heat shock protein 90 (HSP90; HSP90_F1/HSP90_R1; Blair et al. 2008), β-tubulin (β-TUB; BTUB_F1/BTUB_R1; Blair et al. 2008), and the cytochrome c oxidase subunit 1 (COX1; COXF4N/COXR4N; Kroon et al. 2004). After treating PCR products with ExoSap-IT (Thermo Fisher Scientific), respective sequencing primers were added at 1-μM concentration. Sequencing was performed at the Penn State Genomics Core Facility.

    Sequence reads were assembled with the program Geneious v.11.1.5 (Auckland, New Zealand), then ends of each read were trimmed, errors were manually corrected, and heterozygous sites were noted. NCBI’s GenBank was queried with assembled consensus sequences using the NCBI tool BLASTn. Consensus sequences were aligned with ex-type strains or verified reference sequences from Abad et al. (2019) to analyze genetic variation.

    Phylogenetic analysis of P. abietivora was performed by choosing ex-type and well-authenticated representatives from clade 7a with P. cinnamomi and P. niederhauserii as outgroups (Supplementary Table S1). Concatenated sequences of ITS, COX1, β-TUB, HSP90, and NADH1 from four representative strains PDA1791, PDA1792, PDA1795, and PDA1807 were aligned with 32 concatenated sequences and all sites were treated with equal weight. Assembled reads and selected reference sequences were aligned using the program MUSCLE v.3.8.425 (https://kbase.us/applist/apps/kb_muscle/MUSCLE_nuc/release) before being trimmed and corrected (Edgar 2004). A maximum-likelihood (ML) analysis was performed using the PhyML 3.3.20180621 plugin (Geneious) with the TN93 substitution model and was bootstrapped 1,000 times (Guindon et al. 2010). Gaps were treated as missing data. Representative sequences derived from strains in this survey were deposited to GenBank (Table 1).

    Table 1. Representative strains and their GenBank accession numbersz

    Koch’s postulates for P. abietivora.

    Koch’s postulates were fulfilled by performing a nonwounded, detached-leaf and a wounded-leaf, live-plant inoculation assay following modified methods of Blomquist et al. (2005). Six plant species that were repeatedly observed with foliar infections in the forest were tested; pathogenicity tests were not performed on four other plant species found infected. A stem inoculation assay following modified methods of Brazee (2016b) was performed on Q. rubra L. and B. lenta L. because stem cankers were found on these species. Additionally, an infested soil assay following modified methods of Holmes and Benson (1994) was performed on Q. rubra seedlings to determine if P. abietivora causes root disease on this host.

    Each plant species was inoculated with the strains originally isolated from the infected plant parts. Plants for stem and foliar inoculation were sourced from the Pennsylvania Department of Conservation and Natural Resources Forestry Bureau nursery, transferred to 2-liter pots in sterilized Lambert LM-8 bark mix (at pH 7.1; Lambert Peat Moss Company, Québec, Canada), then grown in a greenhouse for 1 month to maintain new growth and continuous vigor and confirmed to be free of Phytophthora. The average seedling height was 39 cm, with an average stem diameter of 12 mm at 5 cm from the soil line. For the infested soil assay, Q. rubra acorns were germinated in 2-liter pots in sterilized Lambert LM-8 bark mix and raised in the greenhouse for 3 months before inoculation. The average height of each seedling was 21 cm with a stem diameter of 5 mm at 5 cm from the soil line before inoculation.

    For the nonwounded detached-leaf assay, healthy leaves with petiole still attached were used. Six 3-month-old leaves and six 1-month-old leaves from each species were surface-sterilized with 70% ethanol and rinsed with dH2O. Two 5-mm V8a plugs were transferred from the leading edge of a 7-day-old colony and placed directly on top of each leaf mycelial-side down, comprising 24 plugs per host and 144 total inoculations. Three leaves from each host were also inoculated with two sterile V8a plugs as a control. Leaves were placed flat in an incubation chamber at 20°C and maintained at 95% relative humidity under 8 h of fluorescent lighting per day. After 10 days, the plugs were removed and lesions were measured. The leaves were surface-sterilized with 70% ethanol and rinsed with dH2O. Symptomatic tissue was placed on CMA-PARP for reisolation and confirmation of P. abietivora.

    For the wounded attached-leaf assay, three 2-year-old seedlings of each respective species were inoculated. Each leaf was surface-sterilized with 70% ethanol and rinsed with dH2O before inoculation. One 5-mm V8a plug was taken from the edge of a 7-day-old colony and pinned onto each leaf with a sterile pin, with four leaves per plant and 72 total inoculations. One plant of each species was also pinned with four sterile V8a plugs as a control. Plants were then kept at 20°C and misted in an incubation chamber with dH2O every 30 min for 6 s for 10 days, when lesion measurements were taken, but disease progression proceeded for 20 days. Symptomatic tissue was surface-sterilized with 70% ethanol and rinsed with dH2O before being placed on CMA-PARP for reisolation and confirmation of P. abietivora.

    For the stem inoculation assay, four 2-year-old plants of Q. rubra and B. lenta were inoculated (12 plants total), with two plants as controls for each host. Each plant’s stem was surface-sterilized with 70% ethanol, and a 1.5 × 0.5 cm section of the bark was removed at 5 cm above the soil line using a sterile scalpel. A 9 mm, V8a plug taken from the edge of a 7-day-old culture was placed, Parafilmed (Bemis Company Inc., Neenah, WI) with the mycelial-side down, and sealed with tape. For the control, two plants were inoculated using the same procedure but with a sterile V8a plug. The plants were incubated at 18 to 25°C in a greenhouse for 8 weeks and watered once a week with dH2O. After 8 weeks, the tape and Parafilm was removed, the bark was removed up to healthy vascular tissue, and the leading edge of the necrotic lesion was measured using a standard ruler from the edge of the original inoculation point. The wounds were then surface-sterilized with 70% ethanol and rinsed with dH2O. Symptomatic tissue was placed on CMA-PARP for reisolation and confirmation of P. abietivora.

    For the soil infestation assay, two 1-liter flasks with 150 g of rice and 220 ml of dH2O each were autoclaved and allowed to cool. Three 5-mm-diameter plugs from 7-day-old cultures of P. abietivora and P. cinnamomi that were recovered from soil were added to each flask and allowed to grow at 20°C for 20 days, with shaking of the flask every 48 h. After the rice was fully colonized, it was pulverized and 14 g of rice was mixed with 350 g of sterilized soil per pot, or a 4% inoculum mixture. In total, 20 Q. rubra seedlings were inoculated: 10 with P. abietivora, five with P. cinnamomi, and five as a negative control with sterile rice added. The plants were kept in the greenhouse at 18 to 25°C and watered heavily with dH2O, allowing the soil to completely dry before watering again. After 90 days, seedlings were removed from their pots, rinsed thoroughly with tap water, and analyzed for cankers or fine root necrosis under a dissection microscope. Lesions were measured using a standard ruler, then roots were surface-sterilized in 70% ethanol, rinsed again in dH2O, and allowed to dry before placing on CMA-PARP to confirm P. abietivora.

    Data analysis.

    To fulfill Koch’s postulates, significance values (P ≤ 0.05) were determined by conducting a single-factor analysis of variance between inoculated plants and their controls. A Tukey multiple comparison test (P ≤ 0.01) was used to separate the means and rank inoculation methods with host susceptibility toward P. abietivora. Relationships between the presence of Phytophthora and crown status was determined using logistic regression analyses from a contingency table and evaluated using a Fisher’s exact test (P ≤ 0.05), while significance among pH values, soil texture, temperature value, and Phytophthora spp. was analyzed using a two-tailed Student’s t test (P ≤ 0.05). Comparison of mean radial growth rates among P. abietivora strains was analyzed using analysis of variance and a Tukey test. Data were analyzed using the program Microsoft Excel 2016 v.16.0 (Redmond, Washington) and the R v.3.6.3 statistical software (R Core Team 2020). The geographical map was developed using the program ArcGIS Pro v.2.4.3 (Esri Inc., Redlands, California). Soil texture was determined by entering sample site coordinates into the California Soil Resource Lab mapping tool, SoilWeb (https://casoilresource.lawr.ucdavis.edu/gmap/).

    Results

    In total, 64 sampling sites yielded 235 Phytophthora strains representing 21 species in nine clades and one unidentified species (Table 2; Fig. 1). Four strains tentatively identified as P. cactorum (n = 1) and P. citricola sensu lato (n = 3) based on morphological traits, were not included in the data because of poor sequencing results. All strains displayed morphology consistent with the species unless otherwise noted. P. abietivora was the most frequently recovered species (n = 40; 17.3%), which was likely because of intensive surveys at four sites where this species was found over a period of 3 years. When strains from the same sampling dates and sites are omitted, P. abietivora represents 9.5% (n = 22) of the total isolates. According to the U.S. Department of Agriculture Fungal Database (Farr and Rossman 2021), this survey resulted in 30 new Phytophthora associations from diseased tissue, and 28 from soil around hosts (five associated with soil and plant tissue; Table 3).

    Table 2. Abundance of Phytophthora spp. from soil, plants, and waterz

    Fig. 1.

    Fig. 1. Approximate locations of Phytophthora spp. recovered. Repetitive recoveries from survey sites are omitted.

    Download as PowerPoint

    Table 3. Forest plants sampled and their association with Phytophthora spp.

    Isolations from symptomatic plants.

    In total, 89 strains were recovered from symptomatic tissues of 18 plant species in 12 families from 27 sites (Table 2; Fig. 1). Not all plant species were equally sampled because sampling was based on visual symptoms. Thirteen Phytophthora spp. were identified, with species from clades 2 and 7 representing 86% of the total infections. Five symptomatic plants were coinfected by two Phytophthora species. Although the survey was conducted from early May until mid-September, 74% of isolations were from samples collected in May and June. Out of the 89 strains, 13.4% were recovered from riparian plants previously inundated with water, and 56% were derived from succulent growth. B. lenta was observed infected the most (n = 25), followed by Fagus grandifolia Ehrh. (n = 19), R. maximum (n = 14), and Q. rubra (n = 8). Q. alba was associated with the most Phytophthora taxa (n = 11), followed by B. lenta (n = 10), Q. rubra (n = 9), and F. grandifolia (n = 8; Table 3). Observed infected plants included saplings (57.3%), mature plants (21.3%), seedlings (14.6%), and basal sprouts (5.6%). The most common symptoms observed on saplings, mature plants, seedlings, and basal sprouts were shepherd’s crook (45.1%), shoot blight (57.9%), shepherd’s crook (46.2%), and necrosis (100%), respectively. Bleeding cankers on the bole of mature F. grandifolia trees were sampled from five survey sites, which yielded no Phytophthora but Fusarium spp.

    Isolations from stream baiting.

    A total of 70 strains representing 12 Phytophthora taxa from four clades (50% from clade 6) were isolated from 48 water samples comprising 163 baited-Rhododendron leaves (four leaves per bait bag, one per BOB) at 34 survey sites (Table 2; Fig. 1). Three BOB water samples did not yield Phytophthora. Most of the water samples were collected in autumn, with only 20% deployed in the spring. Overall, 75.7% of strains were derived from streams, and 24.2% were from lakes (55.7 and 44.3% recovered from BOB and bait bags, respectively, although both methods were not equally deployed). From 48 water samples, 17 yielded more than one species, and two samples resulted in four species each. P. irrigata was coinhabiting water samples the most, with P. macilentosa, P. intercalaris, P. xstagnum, P. chlamydospora, P. stricta, and P. ‘bitahaiensis’. Phytophthora was recovered from a water pH ranging from 6.7 to 8.3, with 77% of samples in the range of 7.0 to 7.9. Water temperatures varied from 9.8 to 22°C, with seven taxa and 25% of the total samples collected between 18 and 19°C. P. chlamydospora displayed the widest water-temperature plasticity with sampling temperatures at 9.8 and 22°C.

    Isolations from soil baiting.

    From 137 soil samples, 72 Phytophthora strains representing 12 taxa in seven clades were obtained from 58 samples (42% recovery rate; Table 2; Fig. 1). Phytophthora was isolated most under Q. alba (n = 19), followed by Q. rubra (n = 17), B. lenta (n = 10), Q. velutina Lam. (n = 9), Q. montana Willd. (n = 6), and F. grandifolia (n = 5), although not all species were sampled equally (Table 3). Generally, one species was recovered per sample, but 10 and two samples yielded two and three species, respectively. P. abietivora was coinhabiting with P. cinnamomi and P. pini, and P. plurivora was coinhabiting with P. cinnamomi, P. xcambivora, P. cactorum, and P. chlamydospora. There was no significant association between crown status and the presence of Phytophthora spp. The pH varied from 3.3 to 8.3, with 39% of positive samples having a pH between 6.0 and 6.9. There was a significant difference (P = 0.004) in pH values between positive and negative samples, with Phytophthora residing in samples with a pH > 5.5 compared with the negative samples (pH ≤ 5.5). Phytophthora-positive samples had a soil texture of sandy loam (51.4%), followed by silt loam (45.8%), and clay loam (2.8%), compared with negative samples with a texture of silt loam (66.2%) and sandy loam (33.8%).

    Characterization of recovered Phytophthora spp.

    Clade 1.

    P. cactorum was isolated from stem cankers, necrotic basal shoots, and foliar necrosis of F. grandifolia and B. lenta seedlings and saplings, and once from soil (pH 6.6) under a healthy Q. alba tree. At one mesophytic-forest site, P. cactorum was isolated from sunken cankers on the root collar of four deceased B. lenta saplings. When aligned with P0714 WPC (MG783385), the ITS region of eight strains differed by one nucleotide. The COX1 region of PDA1788 was three-nucleotides different from P0714 WPC (MH136858).

    Clade 2.

    P. citrophthora was isolated from a single necrotic Sassafras albidum (Nutt.) Nees basal shoot 20 cm from the forest floor of a mature oak/hickory forest in June 2019. Strain PDA1843 is one base-pair (bp) different in the ITS region of P0479 WPC (MG865476). The COX1 region was five-nucleotides different from P0479 WPC (MH136872) and identical to NJJB2013-HR-27 (KJ631594).

    P. plurivora is a common soil inhabitant (pH range 3.9 to 7.3) associated with Quercus in Pennsylvania, and an aerial pathogen of R. maximum, B. lenta, and F. grandifolia, causing shoot blight and shepherd’s crook. The ITS region of 18 strains was identical to ex-type CBS 124093 (FJ665225), while 14 strains differed by one nucleotide. Alignment of the COX1 region showed that nine strains differed from the ex-type (FJ665236) by one nucleotide. The β-TUB region of two strains was identical to the ex-type (FJ665247).

    P. pini was recovered from soil under declining Q. alba (pH range 4.4 to 7.6), and from shepherd’s crook, stem cankers, and shoot blights of six plant species. The ITS region of 19 strains differed from ex-type ATCC 64532 (FJ392322) by one nucleotide. The COX1 region of seven strains differed from the ex-type (GQ247650) by one nucleotide. The β-TUB region of two strains was identical to the ex-type (GQ247656).

    P. caryae was isolated twice from soil (pH 6 and 7), from shoot blight of K. latifolia, and from necrotic stem tissue of a riparian B. lenta sapling. The ITS region of four strains was identical to ex-type NJB2013-AF-08 (KJ631538). The COX1 region of two strains was one-nucleotide different from the ex-type (KJ631586).

    Clade 3.

    P. pseudosyringae was recovered from five soil samples (pH range 5.1 to 6.4) and two streams with a pH of 7.5 and 7.3 and a temperature of 14 and 14.2°C. Sequences of the ITS region of all seven strains were identical to ex-type CBS 111772 (AY230190). However, PDA2082, PDA2043, and PDA2335 differed from the ex-type (LC595938) in the COX1 region by 19, 11, and 22 nucleotides, respectively. Although P. pseudosyringae was described as homothallic, no oospores formed in any strains after 14 days when grown on CMA-PARP, V8a, and HSA (Jung et al. 2003). When paired with A1 and A2 mating types, no oospores developed after 30 days, designating all strains as sterile. Apart from the lack of sexual gametangia, morphological traits were consistent with the species, with semipapillate, globose, ovoid-to-ellipsoid sporangia, borne terminally and sympodial, caducous with a short pedicel, and measuring 46.1 − 51.1 × 29.8 − 37.3 μm; n = 40 (Jung et al. 2003). Intercalary hyphal-swellings were common in flooded V8a plugs, but also abundant in V8a and HSA. Sporangia formed on V8a.

    Clade 5.

    P. heveae was recovered from four soil samples (pH 6.5 to 7.1) in a rich cove-hardwood forest, where it appears to be locally abundant. The four strains differed in the ITS region of P3428 WPC (MG865505) by two nucleotides. The β-TUB region of two strains was two-nucleotides different from P3428 WPC (MH493947).

    Clade 6.

    P. riparia was isolated once from a BOB water sample (pH 6.9, temperature 15.2°C) collected at a reservoir in the Allegheny National Forest (Marienville, PA). The ITS region was identical to ex-type VI_3-100B9F (HM004225).

    P. xstagnum was recovered from five streams with a pH range of 7 to 7.9 and a temperature range of 17 to 19.1°C. The ITS region of five strains was identical to ex-type 43 F3 (KJ705084). The COX1 region of two strains was identical to the ex-type (KC631619).

    P. gonapodyides was recovered from one lake and three streams (pH range of 6.9 to 7.4 and temperature range of 9.8 to 19.9°C), four soil samples (pH range 5.3 to 6.5), and once from necrotic stem tissue of a riparian B. lenta sapling. The ITS region of nine strains differed from CPHST BLD 172 (MG865502) by one nucleotide. The COX1 region of five strains differed from CPHST BLD 172 (MH136897) by two nucleotides.

    P. chlamydospora was recovered from 19 streams and five lakes (pH range of 6.7 to 8 and temperature range of 9.8 to 22°C), eight soil samples (pH range 3.4 to 6.2), and leaf necrosis from two riparian saplings. The ITS region of 28 out of 32 strains was identical to ex-type P236 (AF541900). Four strains differed by one, one, three, and four nucleotides. The COX1 region of four strains was identical to CPHST BL 156 (MH136867).

    P. ‘bitahaiensis’ was recovered from three streams and one lake with a pH range of 7.2 to 7.8 and a temperature range of 16.2 to 18.9°C. The ITS region of four strains differed from BTW2 (KT183447) at two sites. The COX1 region of two strains differed from DQ10-26 (KT899396) at two sites.

    Clade 7.

    P. abietivora was isolated from 10 plant hosts (Fig. 2) and twice from soil (pH of 4.5 and 6.8), including from a diseased Q. rubra sapling (PDA2194) and Carya sp. seeding (PDA2197) in Bath County, Virginia that was not associated with this survey. The ITS region of 32 out of 40 strains differed by one nucleotide from ex-type UAMH12075 (MK163944). The β-TUB region of 22 out of 25 strains was identical to the ex-type (MK164274). The strains from Virginia share a polymorphic site in the β-TUB region that no other strains share. The HSP90 region of 11 strains was identical to the ex-type (MK164275). The COX1 region of 32 strains displayed genetic variance with four distinct groupings (Supplementary Table S2). The NADH1 region also displayed variability with six out of seven strains differing from the ex-type (MK164269) at sites 463 (A instead of G) and 532 (A instead of C). Strain PDA1795 differed at sites 413 (A instead of G) and 472 (C instead of T).

    Fig. 2.

    Fig. 2. Symptoms of Phytophthora abietivora infection observed in the forest. A, Betula lenta sapling stem canker. B, Quercus rubra seedling foliar necrosis. C, Hamamelis virginiana sapling shepherd’s crook. D, Acer rubra shepherd’s crook and foliar necrosis. E, B. lenta sapling terminal dieback. F, Mature Fagus grandifolia foliar necrosis. G, Q. montana seedling shepherd’s crook and foliar necrosis. H, Cornus florida sapling foliar necrosis. I, Ilex montana seedling terminal dieback. J, Q. montana seedling terminal dieback.

    Download as PowerPoint

    P. ‘sp. 1’ was isolated once from a F. grandifolia sapling with shepherd’s crook in a previously unlogged, mesophytic-forest remnant along the Susquehanna River in south-eastern Pennsylvania. Ornamented and smooth-walled oospores with amphigynous antheridia formed in single culture on V8a. When flooded in Non-Sterile Rhododendron Root Extract Solution, nonpapillate sporangia formed abundantly. The ITS region had only 2-bp differences from P. xcambivora neo-type CBS 141218 (KU899179). In contrast, the COX1 region displayed 11-bp differences, and the β-TUB displayed 17-bp differences. Compared with GenBank strains, the COX1 region shared a 98.96% identity with P. xincrassata ex-type CBS 141209 (KU517150) with 10-bp differences, and a 98.85% identity with P. xheterohybrida ex-type CBS 14107 (KU517145), also with 10-bp differences. The β-TUB region was 99.28% identical to P. xalni ex-type P772 (KU899238) with 3-bp differences, and 99.54% identical to P. uniformis ex-type P16206 (MH493905), with 4-bp differences. The NAD9 region was 98.96% identical to P. xalni P10564 ATP (JF771616) with 8-bp differences. The NADH1 region was 99.36% identical to P. ‘xmultiformis-like’ 4971496 (KU899488) with 6-bp differences.

    P. xcambivora was isolated from six soil samples (pH range of 3.9 to 6), and from stem cankers, shepherd’s crook, and foliar necrosis of nine plants (two riparian). The ITS region of 12 out of 15 strains was identical to neo-type CBS 141218 (KU899179), while three strains differed by one nucleotide. The COX1 region of 10 out of 13 strains was identical to the neo-type (KU899334), while three strains differed by three sites. The β-TUB region of one strain differed from the neo-type (LC595893) by four sites. All strains are the A1 mating type, forming oospores characteristic of P. xcambivora when paired with A2 mating type P. nicotianae (Erwin and Ribeiro 1996).

    P. cinnamomi was recovered from 15 soil samples, twice from necrotic roots of wilting R. maximum, and once from symptomatic R. maximum foliage hanging in a stream. The six soil samples collected under healthy trees were significantly more basic (P = 0.034; x¯ = 6.2) than the nine samples collected from declining trees (x¯ = 4.9). The ITS region of 13 out of 19 strains was identical to ex-type CBS 144.22 (KU899160), while six strains differed by one nucleotide. The COX1 region of three strains differed from the ex-type (KU899315) by one nucleotide. All strains are the A2 mating type, forming oospores characteristic of P. cinnamomi when paired with A1 mating type P. xcambivora (Erwin and Ribeiro 1996).

    Clade 8.

    P. cryptogea was isolated once from silt loam soil (pH 6.7) under a recently dead, mature Q. alba tree. The ITS region differed from ex-type CBS 113.19 (MG865483) by seven nucleotides, and from W253 (KF271791) by one nucleotide. The COX1 region differed from the ex-type (MW927571) by three nucleotides.

    P. stricta (no ITS clade, designated in clade 8 according to Yang et al. 2017) was recovered from four streams and one lake (pH and temperature range of 6.7 to 7.9 and 15.7 to 19.9°C, respectively), and twice from necrotic, riparian R. maximum leaves and stems. The ITS region of six out of seven strains differed from ex-type 58A1 (KF192694) by one nucleotide. The COX1 region of three strains was identical to the ex-type (KF192702). The β-TUB region of one strain differed from the ex-type (MH494015) by four nucleotides.

    Clade 9.

    P. macilentosa was isolated once from a lake in east-central Pennsylvania with a pH of 7.7 and a temperature of 20.5°C. The ITS region differed from ex-type 58A7 (KF192700) by one nucleotide. The β-TUB region was identical to 58A5 (KX252330).

    P. irrigata was recovered from five streams and five lakes with a pH and temperature range of 6.9 to 7.7 and 14.6 to 20.5°C, respectively. The ITS region of four out of 10 strains was identical to ex-type 23J7 (EU334634), with four and two strains differing by one and two nucleotides, respectively.

    P. hydropathica was recovered from two lakes and five streams with a pH and temperature range of 7.1 to 8.3 and 14.7 to 19.3°C, respectively. The ITS region of three strains was identical to ex-type 5D1 (EU583793), while four strains differed by four nucleotides. The B-TUB of three strains was identical to the ex-type (GQ260069).

    Clade 10.

    P. intercalaris was recovered from three streams with a pH and temperature range of 7 to 7.9 and 15.7 to 18.9°C, respectively. The ITS region of the three strains was identical to ex-type 45B7 (KT163268). The β-TUB region of one strain was identical to the ex-type (KT163336).

    Koch’s postulates for P. abietivora.

    In the nonwounded leaf assay, immature leaves were significantly more susceptible to infection than mature leaves (P = 0.01). Only immature leaves of Q. rubra, Q. montana, F. grandifolia, and B. lenta were significantly different from the controls (Table 4). Nonwounded immature leaves resulted in an overall infection rate of 41.7% for all plant hosts. The symptoms on immature leaves included brown, water-soaked lesions, with no significant differences in lesion size between hosts. Light-brown lesions formed on the controls but no Phytophthora was recovered. P. abietivora could not be recovered from mature tissue of any hosts without wounding, and lesions were not significantly different from the controls (Table 4).

    Table 4. Average necrotic leaf area (mm ± standard deviation) from three foliar inoculation assays 10 days after inoculation

    Wounding immature foliage once with a pin significantly increased the rate of infection compared with nonwounded immature leaves (P = 0.002). When lesions formed, they were larger than those on the nonwounded leaves or the controls (Table 4). Lesions were water-soaked, eventually turning necrotic, and systemically progressed through veins in the leaf and into the petiole (Fig. 3). P. abietivora was recovered from symptomatic tissue from 79.2% of the inoculation points, although the severity of infection differed between hosts. Except for H. virginiana, the averaged lesion size for all plants treated after 10 days was significantly different from the controls. Light-brown lesions formed on the controls but no Phytophthora was recovered.

    Fig. 3.

    Fig. 3. Symptoms on seedlings from the Phytophthora abietivora wounded-leaf assay 10 days after inoculation. A, Foliar necrosis on Quercus rubra. B, Terminal shoot dieback on Q. rubra. C, Foliar necrosis on Betula lenta. D, Foliar necrosis on Fagus grandifolia. E, Foliar necrosis on Acer rubrum. F, Necrotic leaf spots on Hamamelis virginiana. G, Foliar necrosis on Q. montana. H, Dieback on Q. montana seedlings 15 days after inoculation.

    Download as PowerPoint

    P. abietivora was reisolated from necrotic vascular tissue in 50% of Q. rubra and 100% of B. lenta seedlings 56 days after stem inoculation. The necrotic tissue from B. lenta was significantly larger than Q. rubra (P = 0.009), but because two Q. rubra seedlings died during incubation, the extent of infection could not be determined. Excluding the two dead specimens, there was no significant difference in lesion size from the controls; however, with B. lenta there was (P = 0.003; Table 5). With both species, infection progressed lengthwise, causing stem necrosis (B. lenta seedlings developed sunken cankers) covering ∼50% of the circumference of the stem in B. lenta and 30% in Q. rubra (Fig. 4).

    Table 5. Average necrotic vascular tissue (mm ± standard deviation) 56 days after inoculation (n = 12)

    Fig. 4.

    Fig. 4. Symptoms from the stem inoculation assay on Betula lenta (A to E) and Quercus rubra (F to J) seedlings 56 days from inoculation with Phytophthora abietivora. E, B. lenta control. J, Q. rubra control. Bar = 1 cm.

    Download as PowerPoint

    After 90 days post soil inoculation, P. abietivora was isolated from stem cankers and necrotic fine roots from four out of 10 Q. rubra seedlings, with lesions significantly larger than the controls (P = 0.027; Table 6; Fig. 5). In comparison, P. cinnamomi was recovered from all the seedlings inoculated, causing extensive root cankers and fine root necrosis, with a significant difference in lesion size from the controls (P = 0.007). P. cinnamomi caused significantly more and larger lesions than P. abietivora (P = 0.002). All the controls exhibited fine root necrosis but no Phytophthora was recovered. After a 90-day incubation period, seedlings inoculated with P. cinnamomi prematurely dropped some of their leaves while seedlings inoculated with P. abietivora did not.

    Table 6. Average necrotic root area (mm ± standard deviation) on Quercus seedlings 90 days after inoculation (n = 20)

    Fig. 5.

    Fig. 5. Symptoms from the infested soil assay on Quercus rubra seedlings 90 days after inoculation with Phytophthora abietivora. A, B, and D, Root cankers. C, Fine root necrosis are shown. Ruler intervals = 1 mm.

    Download as PowerPoint

    Phylogenetic analysis of P. abietivora.

    Including outgroups, the aligned and trimmed sequences from five loci included 36 strains and a total of 4,245 characters. The nuclear ITS, β-TUB, and HSP90 loci and the mitochondrial COX1 and NADH1 loci resulted in a dataset of 832, 918, 837, 861, and 797 nucleotides, respectively. There were 1, 3, and 10 gaps in the COX1, HSP90, and ITS regions. Excluding outgroups and ambiguous sites, the aligned datasets of ITS, β-TUB, HSP90, COX1, and NADH1 contained 35 (4.2%), 43 (4.7%), 61 (7.3%), 95 (11%), and 54 (6.8%) polymorphic nucleotides, respectively, and in total 288 (6.8%) sites were variable. Most variations (91.6%) were single-bp polymorphisms, with 4, 1, 2, and 3 two-bp variations in ITS, HSP90, COX1, and NADH1 regions, respectively. The four representative strains and P. abietivora UAMH12075 differed by a total of 12 nucleotides in all five loci and contain two unique polymorphisms in the COX1 region that no other members of clade 7a share. The topologies from the ML phylogeny were comparable to Jung (2017a, b) and Li et al. (2019), revealing 17 lineages within clade 7a (Fig. 6). The four PDA strains were grouped with the P. abietivora holotype in the same subclade with P. flexuosa and P. europaea, although at different branch lengths and bootstrap values.

    Fig. 6.

    Fig. 6. A maximum-likelihood tree of clade 7a based on ITS, COX1, β-TUB, HSP90, and NADH1 sequences, with four representative strains of Phytophthora abietivora included. P. niederhauserii and P. cinnamomi are used as outgroups. Bootstrap (1,000 bootstraps) support values ≥ 50% are shown next to the nodes. T = ex-type strain.

    Download as PowerPoint

    Morphological and growth characteristics of four P. abietivora strains.

    Morphological traits are shown in Supplementary Fig. S1. The four representative strains produced gametangia abundantly and quickly on V8a, CMA, HSA, and CA, but none on PDA and MEA after 30 days. All four strains developed similar colonies after 10 days at 20°C (Supplementary Fig. S2). P. abietivora on V8a displayed stellate patterns with woolly aerial mycelium; on MEA, it displayed dense, tufted, undulate-to-lobate growth with little aerial mycelium; on CMA, it displayed rhizome-to-dendritic growth; on PDA, it displayed dense, fluffy-to-cottony aerial mycelium; on CA, it displayed filamentous, radiating growth with fleecy aerial mycelium; and on HSA, it displayed appressed mycelium with diffused, sectoring hyphae.

    The cardinal temperatures of the four strains were comparable with most other species in clade 7a (Jung et al. 2017a, b). There was no significant difference in mean growth rates between the four strains and seven temperatures tested (Supplementary Fig. S3). The combined temperature average daily radial growth after 7 days for PDA1792 was 2.93 ± 2.14 mm, PDA1791 was 3.64 ± 2.52 mm, PDA1795 was 3.63 ± 2.37 mm, and PDA1807 was 3.51 ± 2.35 mm. The minimum temperature for viability was 2°C and the maximum was 35°C. Strains kept at 2°C for 7 days survived while strains kept at 37°C did not. The optimum temperature tested was 25°C with an average radial growth rate of 6.46 ± 0.47 mm/day.

    Discussion

    The Appalachian forests and waterways of Pennsylvania harbor a diverse assemblage of Phytophthora spp. that can be recovered from symptomatic plants, soil, and water during favorable seasons. Taxa recovered in the natural environment but not detected in Pennsylvania nurseries and greenhouses are P. macilentosa, P. pseudosyringae, P. riparia, and P. stricta (Molnar et al. 2020). Excluding P. abietivora, P. ‘sp. 1’, and P. ‘bitahaiensis’, all species from this survey have been previously identified from natural forests and waterways of the eastern-central United States, demonstrating that these species have a broad, but perhaps regional geographical range (Balci et al. 2007, 2010; Brazee et al. 2016a; Cambell and Gallegly 1965; Hwang et al. 2009; Jung et al. 2018; Reed et al. 2019; Widmer et al. 2018). Taxa that were previously identified from eastern-central forest soils but not recovered in this survey include P. europaea, P. quercetorum, and P. quercina (Balci et al. 2007, 2008; Reed et al. 2019; Schwingle et al. 2007). Taxa previously identified from eastern-central waterways but not in this survey include P. lacustris, P. lacustris × riparia, P. hydrogena, P. polonica, P. caryae, P. pini, P. citrophthora, P. ‘pocumtuck’, P. ‘personii’, and P. ‘aquatilis’ (Brazee et al. 2016a, b; Hong et al. 2011, 2012; Hulvey et al. 2010; Hwang et al. 2009; Yang et al. 2014, 2016). The recovery of certain species over others may be dependent on local plant communities, sampling methodology, and regional climate (Jung et al. 2018).

    R. maximum leaves used as bait may have imparted a bias toward some species. Previous research comparing baiting types indicated a preference of certain Phytophthora taxa to specific plant material (Ferguson and Jeffers 1999; Jeffers and Aldwinckle 1987; Reeser et al. 2011; Rollins et al. 2016). Brazee et al. (2016a) found that Rhododendron leaves yielded the greatest number of isolates and taxa from Massachusetts waterways compared with pear and pepper baits. McConnell and Balci (2014) observed that English oak leaflets were more sensitive than Rhododendron leaves in baiting soilborne Phytophthora, doubling the isolation frequency. In this survey, R. maximum leaves were chosen because of the species natural distribution in the sampling area, access to fresh material, susceptibility to Phytophthora infection, uniform size, and ability to withstand degradation in both ex situ and in situ baiting assays. It is likely that using different types of baits, filtering water, and air-drying soil samples before baiting would affect the amount and diversity of Phytophthora recoveries.

    Most Phytophthora symptoms (74%) on plants were observed from May through June, when new growth on vegetation is expanding and plants are most vulnerable to infection (Balci et al. 2008). These months had the highest monthly rainfall for south-central Pennsylvania from 2018 to 2020, with a combined yearly average of 160, 117, and 196 mm for May, June, and July, respectively (NOAA 2021). The combined average temperatures for south-central Pennsylvania during May, June, and July were 19, 22, and 26°C, respectively (NOAA 2021), which are within the cardinal temperatures for many Phytophthora spp., and close to the optimum temperature for P. abietivora (Supplementary Fig. S3; Erwin and Riberio 1996). Weather patterns are a significant impetus for disease transmission of Phytophthora in the forest, with frequent precipitation and relative humidity >90% correlating with a larger production of zoospores (Davidson et al. 2005; Erwin and Riberio 1996; Kuske and Benson 1983; Redondo et al. 2018). Therefore, in most of Pennsylvania, the periods with the greatest Phytophthora activity coincide closely with the seasons that plants are most susceptible to infection.

    The assemblage, distribution, and prevalence of Phytophthora from soil was comparable to findings from previous surveys, with some differences. Results from this survey and previous surveys suggest that P. plurivora, P. chlamydospora, P. pini, P. cinnamomi, P. abietivora, and P. xcambivora are widely distributed in Pennsylvania (Balci et al. 2007; McConnell and Balci 2014). P. plurivora was the most prevalent species from soil, contrasting other surveys in eastern-central states where P. cinnamomi prevailed (Balci et al. 2007, 2010; McConnell and Balci 2014; Reed et al. 2019). This is most likely because Pennsylvania lies on the climatic threshold for P. cinnamomi while P. plurivora is more cold-tolerant (Beaulieu et al. 2017; Burgess et al. 2017). The three P. cinnamomi strains from soil under Q. rubra, Q. alba, and F. grandifolia at 41° latitude are the most northern recoveries from forested ecosystems in the eastern United States (McConnell and Balci 2014). The presence of P. chlamydospora and P. gonapodyides from soil confirms that these traditional opportunists are deposited along vernal gullies and poorly-drained depressions during seasonal flooding and can still be recovered during drier seasons. Nine species were recovered from plant hardiness zone 5b, including P. cinnamomi; P. pseudosyringae was only recovered from zone 5b. P. heveae, P. cryptogea, and P. citrophthora were encountered infrequently from zone 6b in south-eastern Pennsylvania, suggesting that these species may be cold-intolerant or uncommon.

    The three most prevalent aquatic species in this survey were P. chlamydospora, P. irrigata, and P. hydropathica. In general, the pH-neutral (x¯ = 7.4) waterways of Pennsylvania are conducive for zoospore survival and motility (Kong et al. 2009). One mountain stream in Michaux State Forest (Fayetteville, PA) recovered 11 taxa over 2 years from water baiting and symptomatic, riparian plants. Overall, the Phytophthora communities found in soil (n = 12) and water (n = 12) shared only three species: P. chlamydospora, P. gonapodyides, and P. plurivora. In this survey, P. plurivora had a significant presence in waterways, while other clade 2c members P. pini and P. caryae were restricted to plants and soil. P. cinnamomi, P. cactorum, and P. xcambivora were isolated from symptomatic, previously submerged riparian foliage, suggesting that inoculum from these species is deposited in streams. P. stricta was isolated twice from foliar and stem lesions of R. maximum, supporting evidence from Widmer et al. (2018) that this species causes disease on riparian hosts. It should be noted that P. ‘bitahaiensis’ has never been reported from the United States, although it has been recovered from both streams and nursery irrigation reservoirs in Pennsylvania (unpublished data).

    Several soil-residing species were found causing aerial infections in the forest. It is unclear whether these infections are a result of motile zoospores or from infested soil splashing onto aerial parts during heavy rains. Infections from this survey were found as high up as 1.5 m in the canopy from species with persistent sporangia, including P. abietivora, P. pini, P. xcambivora, P. plurivora, P. caryae, P. citrophthora, and Phytophthora ‘sp. 1’. This phenomenon was observed on Fagus sylvatica L. understory plants in Germany (Nechwatal et al. 2011). Infested soil may serve as the primary inoculum for a foliar infection, and resting spores that persist in plant tissue may be released back into the soil after senescence and decomposition by other saprobe fungi and bacteria, resulting in a multicyclic disease (Jung et al. 2018). The infections observed from P. abietivora on the same hosts over 3 years at four sites suggest that this mode of transmission occurs.

    Our results show that P. abietivora is a pathogen of five out of six hosts tested, causing foliar necrosis, terminal-shoot dieback, stem and root cankers, and fine root necrosis on seedlings under favorable artificial conditions. Overall, foliage age and wounding were important factors in lesion formation, and observations from the forest were most comparable to the wounded, immature-leaf assay (Figs. 2 and 3). However, disease progression appeared to be highly dependent on climate, with no infections observed after mid-July. Although P. abietivora has been shown to cause root rot of A. fraseri in plantations, it is unclear whether it causes perennial infections that may lead to mortality of native broadleaved hosts. Further pathogenicity tests and surveys are needed to fully understand its virulence on canopy trees and impact on understory forest succession.

    In conclusion, a diverse assemblage of Phytophthora taxa can be recovered from natural environments of Pennsylvania using fairly simple baiting methods. No exotic or regulated species of concern were detected during the survey. Further research is needed to determine the extent of P. abietivora’s host range and distribution in nurseries and natural environments.

    Acknowledgments

    The authors thank the Pennsylvania Department of Agriculture assistant Phoebe Nelson and the Pennsylvania Department of Conservation and Natural Resources Forestry Bureau staff Jill Rose, Tim Tomon, and Sarah Johnson for assistance in sample collection.

    The author(s) declare no conflict of interest.

    Literature Cited

    • Abad, Z., Burgess, T., Bienapfl, J., Redford, A. J., Coffey, M., and Knight, L. 2019. IDphy: Molecular and morphological identification of Phytophthora based on the types. U.S. Department of Agriculture’s Animal and Plant Health Inspection Service, Plant Protection and Quarantine S&T ITP, Beltsville, MD. http://idtools.org/id/phytophthora/lucid_key.php Google Scholar
    • Albright, T. 2017. Forests of Pennsylvania, 2016. Resource Update FS-132. U.S. Department of Agriculture, Forest Service Northern Research Station, Newtown Square, PA. CrossrefGoogle Scholar
    • Balci, Y., Balci, S., Blair, J. E., Park, S.-Y., Kang, S.-C., and MacDonald, W. L. 2008. Phytophthora quercetorum sp. nov., a novel species isolated from eastern and north-central USA oak forest soils. Mycol. Res. 112:906-916. https://doi.org/10.1016/j.mycres.2008.02.008 CrossrefGoogle Scholar
    • Balci, Y., Balci, S., Eggers, J., MacDonald, W. L., Juzwik, J., Long, R. P., and Gottschalk, K. W. 2007. Phytophthora spp. associated with forest soils in eastern and north-central U.S. oak ecosystems. Plant Dis. 91:705-710. https://doi.org/10.1094/PDIS-91-6-0705 Link, ISIGoogle Scholar
    • Balci, Y., and Bienapfl, J. 2013. Phytophthora in US forests. Pages 135-145 in: Phytophthora: A Global Perspective. Centre for Agriculture and Bioscience International, Wallingford, Oxfordshire, UK. https://doi.org/10.1079/9781780640938.0135 CrossrefGoogle Scholar
    • Balci, Y., Long, R., Mansfield, M., Balser, D., and MacDonald, W. L. 2010. Involvement of Phytophthora species in white oak (Quercus alba) decline in southern Ohio. For. Pathol. 40:430-442. https://doi.org/10.1111/j.1439-0329.2009.00617.x Crossref, ISIGoogle Scholar
    • Beaulieu, J., Ford, B., and Balci, Y. 2017. Genotypic diversity of Phytophthora cinnamomi and P. plurivora in Maryland’s nurseries and mid-Atlantic forests. Phytopathology 107:769-776. https://doi.org/10.1094/PHYTO-05-16-0215-R Link, ISIGoogle Scholar
    • Blair, J. E., Coffey, M. D., Park, S.-Y., Geiser, D. M., and Kang, S.-C. 2008. A multi-locus phylogeny for Phytophthora utilizing markers derived from complete genome sequences. Fungal Genet. Biol. 45:266-277. https://doi.org/10.1016/j.fgb.2007.10.010 Crossref, ISIGoogle Scholar
    • Blomquist, C., Irving, T., Osterbauer, N., and Reeser, P. 2005. Phytophthora hibernalis: A new pathogen on Rhododendron and evidence of cross amplification with two PCR detection assays for Phytophthora ramorum. Plant Health Prog. 6:27. https://doi.org/10.1094/PHP-2005-0728-01-HN LinkGoogle Scholar
    • Brasier, C. 2009. Phytophthora Biodiversity: How many Phytophthora species are there? Pages 101-115 in: Proceedings of the 4th Meeting of the International Union of Forest Research Organizations (IUFRO) Working Party S07.02.09: Phytophthoras in Forests and Natural Ecosystems. Gen. Tech. Rep. PSW-GTR-221. U.S. Department of Agriculture, Forest Service, Pacific Southwest Research Station, Albany, CA. Google Scholar
    • Brazee, N., Wick, R., and Hulvey, J. 2016a. Phytophthora species recovered from the Connecticut River Valley in Massachusetts, USA. Mycologia 108:6-19. https://doi.org/10.3852/15-038 Crossref, ISIGoogle Scholar
    • Brazee, N., Yang, X., and Hong, C. 2016b. Phytophthora caryae sp. nov., a new species recovered from streams and rivers in the eastern United States. Plant Pathol. 66:1-13. ISIGoogle Scholar
    • Burgess, T. I., Scott, J. K., McDougall, K. L., Stukely, M. J. C., Crane, C., Dunstan, W. A., Brigg, F., Andjic, V., White, D., Rudman, T., Arentz, F., Ota, N., and Hardy, G. E. S. J. 2017. Current and projected global distribution of Phytophthora cinnamomi, one of the world’s worst plant pathogens. Glob. Change Biol. 23:1661-1674. https://doi.org/10.1111/gcb.13492 Crossref, ISIGoogle Scholar
    • Cambell, W., and Gallegly, M. 1965. Phytophthora heveae from eastern Tennessee and western North Carolina. Plant Dis. Rep. 49:233-234. Google Scholar
    • Corcobado, T., Cech, T. L., Brandstetter, M., Daxer, A., Hüttler, C., Kulacek, T., Jung, M. H., and Jung, T. 2020. Declining of European beech in Austria: Involvement of Phytophthora spp. and contributing biotic and abiotic factors. Forests 11:895. https://doi.org/10.3390/f11080895 Crossref, ISIGoogle Scholar
    • Davidson, J. M., Wickland, A. C., Patterson, H. A., Falk, K. R., and Rizzo, D. M. 2005. Transmission of Phytophthora ramorum in mixed evergreen forest in California. Phytopathology 95:587-596. https://doi.org/10.1094/PHYTO-95-0587 Link, ISIGoogle Scholar
    • Dick, M. 1990. Keys to Pythium. University of Reading Press, Reading, UK. Google Scholar
    • Edgar, R. 2004. MUSCLE: Multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32:1792-1797. https://doi.org/10.1093/nar/gkh340 Crossref, ISIGoogle Scholar
    • Erwin, D., and Riberio, O. 1996. Phytophthora Diseases Worldwide. American Phytopathological Society Press, St. Paul, MN. Google Scholar
    • Farr, D., and Rossman, A. 2021. Fungal Databases. U.S. National Fungus Collections. Agricultural Research Service, U.S. Department of Agriculture, Washington, DC. Google Scholar
    • Ferguson, A., and Jeffers, S. 1999. Detecting multiple species of Phytophthora in container mixes from ornamental crop nurseries. Plant Dis. 83:1129-1136. https://doi.org/10.1094/PDIS.1999.83.12.1129 Link, ISIGoogle Scholar
    • Fichtner, E., Lynch, S., and Rizzo, D. 2009. Survival, dispersal, and soil-mediated suppression of Phytophthora ramorum in a California redwood-tanoak forest. Phytopathology 99:608-619. https://doi.org/10.1094/PHYTO-99-5-0608 Link, ISIGoogle Scholar
    • Guindon, S., Dufauard, J., Lefort, V., and Anisimova, M. 2010. New algorithms and methods to estimate maximum-likelihood phylogenies: Assessing the performance of PhyML 3.0. Syst. Biol. 59:307-321. https://doi.org/10.1093/sysbio/syq010 Crossref, ISIGoogle Scholar
    • Hansen, E. 2015. Phytophthora species emerging as pathogens of forest trees. Curr. For. Rep. 1:16-24. https://doi.org/10.1007/s40725-015-0007-7 ISIGoogle Scholar
    • Hansen, E., Reeser, P., Sutton, W., and Brasier, C. 2015. Redesignation of Phytophthora taxon Pgchlamydo as Phytophthora chlamydospora sp. nov. N. Am. Fungi 10:1-14. Google Scholar
    • Holmes, K., and Benson, D. 1994. Evaluation of Phytophthora parasitica var. nicotianae for biocontrol of Phytophthora parasitica on Catharanthus roseus. Plant Dis. 78:193-199. Crossref, ISIGoogle Scholar
    • Hong, C.-X., Gallegly, M. E., Richardson, P. A., Kong, P., and Moorman, G. W. 2008. Phytophthora irrigata, a new species isolated from irrigation reservoirs and rivers in Eastern United States of America. FEMS Microbiol. Lett. 285:203-211. https://doi.org/10.1111/j.1574-6968.2008.01226.x Crossref, ISIGoogle Scholar
    • Hong, C., Gallegly, M., Richardson, P., and Kong, P. 2011. Phytophthora pini Leonian resurrected to distinct species status. Mycologia 103:351-360. https://doi.org/10.3852/10-058 Crossref, ISIGoogle Scholar
    • Hong, C.-X., Richardson, P. A., Hao, W., Ghimire, S. R., Kong, P., Moorman, G. W., Lea-Cox, J. D., and Ross, D. S. 2012. Phytophthora aquimorbida sp. nov. and Phytophthora taxon ‘aquatilis’ recovered from irrigation reservoirs and a stream in Virginia, USA. Mycologia 104:1097-1108. https://doi.org/10.3852/11-055 Crossref, ISIGoogle Scholar
    • Hulvey, J., Gobena, D., Finley, L., and Lamour, K. 2010. Co-occurrence and genotypic distribution of Phytophthora species recovered from watersheds and plant nurseries of eastern Tennessee. Mycologia 102:1127-1133. https://doi.org/10.3852/09-221 Crossref, ISIGoogle Scholar
    • Hwang, J., Oak, S., and Jeffers, S. 2009. Monitoring occurrence and distribution of Phytophthora species in forest streams in North Carolina using bait and filtration methods. Gen. Tech. Rep. PSW-GTR-221. U.S. Department of Agriculture, Forest Service, Pacific Southwest Research Station 91-95, Albany, CA. Google Scholar
    • Jeffers, S., and Aldwinckle, H. 1987. Enhancing detection of Phytophthora cactorum in naturally infested soil. Phytopathology 77:1475-1482. https://doi.org/10.1094/Phyto-77-1475 Crossref, ISIGoogle Scholar
    • Jeffers, S., and Martin, S. 1986. Comparison of two media selective for Phytophthora and Pythium species. Plant Dis. 70:1038-1043. https://doi.org/10.1094/PD-70-1038 Crossref, ISIGoogle Scholar
    • Jung, T., Hansen, E. M., Winton, L., Oswald, W., and Delatour, C. 2002. Three new species of Phytophthora from European oak forests. Mycol. Res. 106:397-411. https://doi.org/10.1017/S0953756202005622 Crossref, ISIGoogle Scholar
    • Jung, T., Jung, M. H., Cacciola, S. O., Cech, T., Bakonyi, J., Seress, D., Mosca, S., Schena, L., Seddaiu, S., Pane, A., Magnano di San Lio, G., Maia, C., Cravador, A., Franceschini, A., and Scanu, B. 2017a. Multiple new cryptic pathogenic Phytophthora species from Fagaceae forests in Austria, Italy and Portugal. Int. Mycol. Assoc. Fungus 8:219-244. https://doi.org/10.5598/imafungus.2017.08.02.02 Google Scholar
    • Jung, T., Jung, M. H., Scanu, B., Seress, D., Kovács, G. M., Maia, C., Pérez-Sierra, A., Chang, T.-T., Chandelier, A., Heungens, K., Van Poucke, K., Abad-Campos, P., Leon, M., Cacciola, S. O., and Bakonyi, J. 2017b. Six new Phytophthora species from ITS Clade 7a including two sexually functional heterothallic hybrid species detected in natural ecosystems in Taiwan. Persoonia 38:100-135. https://doi.org/10.3767/003158517X693615 Crossref, ISIGoogle Scholar
    • Jung, T., Nechwatal, J., Cooke, D. E. L., Hartmann, G., Blaschke, M., Oßwald, W. F., Duncan, J. M., and Delatour, C. 2003. Phytophthora pseudosyringae sp. nov., a new species causing root and collar rot of deciduous tree species in Europe. Mycol. Res. 107:772-789. https://doi.org/10.1017/S0953756203008074 Crossref, ISIGoogle Scholar
    • Jung, T., Orlikowski, L., Henricot, B., Abad-Campos, P., Aday, A. G., Aguín Casal, O., Bakonyi, J., Cacciola, C. O., Cech, T., Chavarriaga, C., Corcobado, T., Cravador, A., Decourcelle, T., Denton, G., Diamandis, S., Doğmuş-Lehtijärvi, H. T., Franseschini, A., Ginetti, B., Green, S., Glavendekić, M., Hantula, J., Hartmann, G., Herrero, M., Ivic, D., Jung, M. H., Lilja, A., Keca, N., Kramarets, V., Lyubenova, A., Machado, H., Magnano di San Lio, G., Mansilla Vásquez, P. J., Marcais, B., Matsiakh, I., Milenkovic, I., Moricca, S., Nagy, Z. Á., Nechwatal, J., Olsson, C., Oszako, T., Pane, A., Paplomatas, E. J., Pintos Varela, C., Prospero, S., Rial Martínez, C., Rigling, D., Robin, C., Rytkönen, A., Sánchez, M. E., Sanz Ros, A. V., Scanu, B., Schlenzig, A., Schumacher, J., Slavov, S., Solla, A., Sousa, E., Stenlid, J., Talgø, V., Tomic, Z., Tsopelas, P., Vannini, A., Vettraino, A. M., Wenneker, M., Woodward, S., and Pérez-Sierra, A. 2016b. Widespread Phytophthora infestations in European nurseries put forest, semi-natural and horticultural ecosystems at high risk of Phytophthora diseases. For. Pathol. 46:134-163. https://doi.org/10.1111/efp.12239 Crossref, ISIGoogle Scholar
    • Jung, T., Pérez-Sierra, A., Durán, A., Jung, M. H., Balci, Y., and Scanu, B. 2018. Canker and decline diseases caused by soil- and airborne Phytophthora species in forests and woodlands. Persoonia 40:182-220. https://doi.org/10.3767/persoonia.2018.40.08 Crossref, ISIGoogle Scholar
    • Jung, T., Scanu, B., Brasier, C. M., Webber, J., Milenkoviç, I., Corcobado, T., Tomšovsky, M., Panek, M., Bakonyi, J., Maia, C., Bačová, A., Raco, M., Rees, H., Pérez-Sierra, A., and Jung, M. H. 2020. A survey in natural forest ecosystems of Vietnam reveals high diversity of both new and described Phytophthora taxa including P. ramorum. Forests 11:93. https://doi.org/10.3390/f11010093 Crossref, ISIGoogle Scholar
    • Kabrick, J., Dey, D., Jensen, R., and Wallendorf, M. 2008. The role of environmental factors in oak decline and mortality in the Ozark Highlands. For. Ecol. Manage. 255:1409-1417. https://doi.org/10.1016/j.foreco.2007.10.054 Crossref, ISIGoogle Scholar
    • Kong, P., Moorman, G. W., Lea-Cox, J. D., Ross, D. S., Richardson, P. A., and Hong, C.-X. 2009. Zoosporic tolerance to pH stress and its implications for Phytophthora species in aquatic ecosystems. Appl. Environ. Microbiol. 75:4307-4314. https://doi.org/10.1128/AEM.00119-09 Crossref, ISIGoogle Scholar
    • Kroon, L. P. N. M., Bakker, F. T., van den Bosch, G. B. M., Bonants, P. J. M., and Flier, W. G. 2004. Phylogenetic analysis of Phytophthora species based on mitochondrial and nuclear DNA sequences. Fungal Genet. Biol. 41:766-782. https://doi.org/10.1016/j.fgb.2004.03.007 Crossref, ISIGoogle Scholar
    • Kuske, C., and Benson, D. 1983. Survival and splash dispersal of Phytophthora parasitica, causing dieback of Rhododendron. Phytopathology 73:1188-1191. https://doi.org/10.1094/Phyto-73-1188 Crossref, ISIGoogle Scholar
    • Li, D., Schultes, N., LaMondia, J., and Crowles, R. 2019. Phytophthora abietivora, a new species isolated from diseased Christmas trees in Connecticut, U.S.A. Plant Dis. 103:3057. https://doi.org/10.1094/PDIS-03-19-0583-RE Link, ISIGoogle Scholar
    • Liebhold, A. M., Brockerhoff, E. G., Garrett, L. J., Parke, J. L., and Britton, K. O. 2012. Live plant imports: The major pathway for forest insect and pathogen invasions of the US. Front. Ecol. Environ. 10:135-143. https://doi.org/10.1890/110198 Crossref, ISIGoogle Scholar
    • Man in ’t Veld, W. A. 2007. Gene flow analysis demonstrates that Phytophthora fragariae var. rubi constitutes a distinct species, Phytophthora rubi comb. nov. Mycologia 99:222-226. https://doi.org/10.3852/mycologia.99.2.222 Crossref, ISIGoogle Scholar
    • Martin, F., Blair, J., and Coffey, M. 2014. A combined mitochondrial and nuclear multilocus phylogeny of the genus Phytophthora. Fungal Genet. Biol. 66:19-32. https://doi.org/10.1016/j.fgb.2014.02.006 Crossref, ISIGoogle Scholar
    • McConnell, M., and Balci, Y. 2014. Phytophthora cinnamomi as a contributor to white oak decline in mid-Atlantic United States forests. Plant Dis. 98:319-327. https://doi.org/10.1094/PDIS-06-13-0649-RE Link, ISIGoogle Scholar
    • Mirabolfathy, M., Cooke, D. E. L., Duncan, J. M., Williams, N. A., Ershad, D., and Alizadeh, A. 2001. Phytophthora pistaciae sp. nov. and P. melonis: The principal causes of pistachio gummosis in Iran. Mycol. Res. 105:1166-1175. https://doi.org/10.1016/S0953-7562(08)61987-5 Crossref, ISIGoogle Scholar
    • Molnar, C., Nikolaeva, E., Kim, S.-H., Olson, T., Bily, D., Kim, J.-E., and Kang, S.-C. 2020. Phytophthora diversity in Pennsylvania nurseries and greenhouses inferred from clinical samples collected over four decades. Microorganisms 8:1056. https://doi.org/10.3390/microorganisms8071056 Crossref, ISIGoogle Scholar
    • Moreira, A., and Martins, J. 2005. Influence of site factors on the impact of Phytophthora cinnamomi in cork oak stands in Portugal. For. Pathol. 35:145-162. https://doi.org/10.1111/j.1439-0329.2005.00397.x Crossref, ISIGoogle Scholar
    • Nagle, A., Long, R., Madden, L., and Bonello, P. 2010. Association of Phytophthora cinnamomi with white oak decline in Southern Ohio. Plant Dis. 94:1026-1034. https://doi.org/10.1094/PDIS-94-8-1026 Link, ISIGoogle Scholar
    • Nechwatal, J., Hahn, J., Schonborn, A., and Schmitz, G. 2011. A twig blight of understory European beech (Fagus sylvatica) caused by soilborne Phytophthora spp. For. Pathol. 41:493-500. https://doi.org/10.1111/j.1439-0329.2011.00711.x Crossref, ISIGoogle Scholar
    • NOAA. 2021. NOWData—NOAA online weather data. National Oceanic and Atmospheric Administration, Washington, DC. https://www.weather.gov/wrh/Climate?wfo=okx Google Scholar
    • R Core Team. 2020. R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing, Vienna, Austria. Google Scholar
    • Redondo, M., Boberg, J., Stenlid, J., and Oliva, J. 2018. Contrasting distribution patterns between aquatic and terrestrial Phytophthora species along a climatic gradient are linked to functional traits. Multidiscip. J. Microb. Ecol. 12:2967-2980. https://doi.org/10.1038/s41396-018-0229-3 Google Scholar
    • Reed, S., English, J., and Muzika, R. 2019. Phytophthora species detected in two Ozark forests with unusual patterns of white oak mortality. Plant Dis. 103:102-109. https://doi.org/10.1094/PDIS-02-18-0253-RE Link, ISIGoogle Scholar
    • Reeser, P. W., Sutton, W., Hansen, E. M., Remigi, P., and Adams, G. C. 2011. Phytophthora species in forest streams in Oregon and Alaska. Mycologica 103:22-35. https://doi.org/10.3852/10-013 Crossref, ISIGoogle Scholar
    • Rollins, L., Coats, K., Elliott, M., and Chastagner, G. 2016. Comparison of five detection and quantification methods for Phytophthora ramorum in stream and irrigation water. Plant Dis. 100:1202-1211. https://doi.org/10.1094/PDIS-11-15-1380-RE Link, ISIGoogle Scholar
    • Roubtsova, T., and Bostock, R. 2009. Episodic abiotic stress as a potential contributing factor to onset and severity of disease caused by Phytophthora ramorum in Rhododendron and Viburnum. Plant Dis. 93:912-918. https://doi.org/10.1094/PDIS-93-9-0912 Link, ISIGoogle Scholar
    • Ruiz Gómez, F., Navarro-Cerrillo, R., Pérez-de-Luque, A., Oßwald, W., Vannini, A., and Morales-Rodríguez, C. 2019. Assessment of functional and structural changes of soil fungal and oomycete communities in holm oak declined dehesas through metabarcoding analysis. Sci. Rep. 9:5315. https://doi.org/10.1038/s41598-019-41804-y Crossref, ISIGoogle Scholar
    • Scanu, B., Hunter, G. C., Linaldeddu, B. T., Franceschini, A., Maddau, L., Jung, T., and Denman, S. 2014. A taxonomic re-evaluation reveals that Phytophthora cinnamomi and P. cinnamomi var. parvispora are separate species. For. Pathol. 44:1-20. https://doi.org/10.1111/efp.12064 Crossref, ISIGoogle Scholar
    • Schoebel, C. N., Stewart, J., Gruenwald, N. J., Rigling, D., and Prospero, S. 2014. Population history and pathways of spread of the plant pathogen Phytophthora plurivora. PLoS One 9:e85368. https://doi.org/10.1371/journal.pone.0085368 Crossref, ISIGoogle Scholar
    • Schwingle, B., Juzwik, J., Eggers, J., and Moltzan, B. 2007. Phytophthora species in soils associated with declining and non-declining oaks in Missouri forests. Plant Dis. 91:633. https://doi.org/10.1094/PDIS-91-5-0633A Link, ISIGoogle Scholar
    • USDA APHIS PPQ. 2014. Phytophthora ramorum Domestic Regulatory Program Manual. U.S. Department of Agriculture’s Animal and Plant Health Inspection Service, Plant Protection and Quarantine, Washington, DC. Google Scholar
    • Westbrook, J. W., James, J. B., Sisco, P. H., Frampton, J., Lucas, S., and Jeffers, S. N. 2019. Resistance to Phytophthora cinnamomi in American chestnut (Castanea dentata) backcross populations that descended from two Chinese chestnut (Castanea mollissima) sources of resistance. Plant Dis. 103:1631-1641. https://doi.org/10.1094/PDIS-11-18-1976-RE Link, ISIGoogle Scholar
    • White, T., Bruns, T., Lee, S., and Taylor, J. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. Pages 315-322 in: PCR Protocols: A Guide to Methods and Applications. Academic Press, San Diego, CA. CrossrefGoogle Scholar
    • Widmer, T., McMahon, M., and Frederick, R. 2018. Phytophthora stricta isolated from Rhododendron maximum in Pennsylvania. Plant Dis. 102:827. https://doi.org/10.1094/PDIS-09-17-1435-PDN Link, ISIGoogle Scholar
    • Yang, X., Balci, Y., Brazee, N., Loyd, A., and Hong, C. 2016. A unique species in Phytophthora clade 10, Phytophthora intercalaris sp. nov., recovered from stream and irrigation water in the eastern USA. Int. J. Syst. Evol. Biol. 66:845-855. https://doi.org/10.1099/ijsem.0.000800 Crossref, ISIGoogle Scholar
    • Yang, X., Copes, W., and Hong, C. 2014. Two novel species representing a new clade and cluster of Phytophthora. Fungal Biol. 118:72-82. https://doi.org/10.1016/j.funbio.2013.11.003 Crossref, ISIGoogle Scholar
    • Yang, X., Tyler, B., and Hong, C. 2017. An expanded phylogeny for the genus Phytophthora. IMA Fungus 8:355-384. https://doi.org/10.5598/imafungus.2017.08.02.09 Crossref, ISIGoogle Scholar

    Funding: This survey was supported by the U. S. Department of Agriculture’s McIntire-Stennis program under grant no. AES 4686, the U. S. Department of Agriculture’s Forest Service grant no. 18-DG-11420004-079, the U. S. Department of Agriculture’s National Institute of Food and Agriculture and Federal Appropriations under grant no. 1016291, and the U. S. Department of Agriculture’s Plant Protection Act Forest Pest Oak Commodity program under grant nos. AP19PPQFO000C298 and AP20PPQFO000C366.

    The author(s) declare no conflict of interest.