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Temporal and Locational Variations of a Phytophthora spp. Community in an Urban Forested Water Drainage and Stream-runoff System
Devin S. Bily, Susan V. Diehl, Madeline Cook, Lisa E. Wallace, Laura L. Sims, Clarence Watson, and Richard E. Baird

Southeastern Naturalist, Volume 17, Issue 1 (2018): 176–201

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Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 176 2018 SOUTHEASTERN NATURALIST 17(1):176–201 Temporal and Locational Variations of a Phytophthora spp. Community in an Urban Forested Water Drainage and Stream-runoff System Devin S. Bily1, Susan V. Diehl1, Madeline Cook2, Lisa E. Wallace3, Laura L. Sims4, Clarence Watson5, and Richard E. Baird6,* Abstract - Species richness and diversity of Phytophthora spp. (water molds) in urban riparian-forest ecosystems, which serve as primary drainage passageways for surface-water runoff, may be attributed to surrounding landscape management, associated vegetation, and environmental conditions. These riparian areas, although generally small, are always flooded during wet seasons and almost completely dry during the hottest parts of each year when there is limited precipitation. Little is known about Phytophthora spp. diversity within these heavily impacted sites. We sampled water, soil, and vegetation (phenology dependent) across 14 dates, over ~2 y at a site containing a drainage ditch that enters Hog Creek, in Rankin County, MS. We cultured all Phytophthora spp. using 4 published protocols to ensure maximum isolation potential. Across all sampling dates, 65 isolations were positive for Phytophthora spp., 12 of which were recovered from vegetation. We employed morphological and internal transcribed sequence (ITS) data to confirm taxa. We determined a total of 11 taxa on the basis of their phylogenetic clustering with known species of Phytophthora in a bayesian analysis. The most common taxa were P. chlamydospora, P. mississippiae, and P. cinnamomi at frequencies of 12.5%, 11.0%, and 10%, respectively. We verified morphologically and by sequence similarity an undescribed species, Phytophthora oaksoil taxon, which has been reported previously in the Western US, as well as other countries, such as Australia. Overall, the bottle-of-bait (BOB) intact-leaf and water-filtration methods had numerically greater frequencies (P ≤ 0.05) than BOB leaf disks, soil-baiting leaf disks, or vegetationsampling protocols. Overall frequency (14%) of Phytophthora spp. was significantly greater (P ≤ 0.05) for the 17 December 2014 sampling date. Even though several taxa identified in this study are reported to be pathogenic to riparian forest trees and vegetation at the Hog Creek site, symptoms on surrounding trees and vegetation was generally limited to foliar lesions, and we observed no visible damage or decline during the study period. It would be judicious to visit different, similar urban habitats to determine if common Phytophthora in this study are present in other central and southern Mississippi riparian habitats. 1Department of Sustainable Bioproducts, Mississippi State University, Mississippi State, MS 39762. 2Department of Biological Sciences, Mississippi State University Mississippi State, MS 39762. 3Department of Biological Sciences, Old Dominion University, Norfolk, VA 23529. 4Department of Environmental Science, Policy, and Management, University of California, Berkeley, CA 94720. 5University of Arkansas Division of Agriculture, Fayetteville, AR 72701. 6Department of Biochemistry, Molecular Biology, Entomology, and Plant Pathology, Mississippi State University, Mississippi State, MS 39762. *Corresponding author - reb58@msstate.edu. Manuscript Editor: Kirsten Work Southeastern Naturalist 177 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 Introduction Phytophthora spp. (water molds) are Oomycete organisms grouped within the phylum Stramenopila, with a diploid life cycle and cellulosic cell walls (Hardham 2007). Distributed worldwide, Phytophthora spp. are significant primary invading pathogens of angiosperms and gymnosperms, and are some of the most destructive diseases of forested ecosystems (Hansen et al. 2000, Hardham et al. 2005, Jung et al. 2002, Rizzo et al. 2007). Phytophthora spp. release motile, biflagellate zoospores from asexual sporangia that act as effective dissemination propagules, which are both chemotactically and electrotactically attracted to their host plants (Hardham 2007). Molecular methods are crucial in determining the phylogenetic relationships of Oomycetes, and the internal transcribed spacer (ITS) regions of rDNA are especially valuable in distinguishing Phytophthora spp. from closely related genera such as Pythium, Peronospora, and Halophytophthora (Cooke et al. 2000). According to the Phytophthora database (phytophthoradb.org), there are 123 formally described species in 10 different clades. Common Phytophthora spp. that are pathogenic on trees and shrubs include P. cinnamomi Rands, P. cactorum (Leb. and Cohn) Schröeter, P. cambivora (Petri) Buisman, P. citricola Sawada, P. cryptogea Pethybridge and Lafferty, and P. palmivora Butler (Jeffers 2005). Many of these organisms occur in the US as previously introduced species that have naturalized, and which can routinely be recovered from areas with a favorable climate and accessible host species (Erwin and Ribeiro 1996). Recent surveys have resulted in the detection of many Phytophthora spp. within riparian and aquatic environments (Hulvey et al. 2010, Hwang et al. 2009, Olson et al. 2013, Shrestha et al. 2013, Warfield et al. 2008). The initiation of these surveys, along with advances in molecular technology, has provided clarity in understanding the prevalence, habitat niches, and distribution of Phytophthora populations in the US (Martin et al. 2014). In 2008, a survey of 8 watersheds in eastern Tennessee resulted in the isolation of P. citricola, P. citrophthora (R.E. Smith) Leonian, P. irrigata Hong and Gallegly, and P. hydropathica Hong and Gallegly (Hulvey et al. 2010). A 2010–2011 survey of 16 streams in middle and eastern Tennessee recovered P. cryptogea, P. hydropathica, P. irrigata, P. gonapodyides (Petersen) Buisman, P. lacustris Brasier, and P. polonica Belbahri (Shrestha et al. 2013). In the Appalachian Mountains of North Carolina, P. cinnamomi, P. citricola, P. citrophthora, P. heveae Thomps., and P. pseudosyringae Jung and Delatour were recovered from streams during a 2007 survey (Hwang et al. 2009). In 2003–2004, a soil survey from Quercus (oak)-dominated forests in 9 eastern and north-central states resulted in the new description of P. quercetorum Balci, as well as the recovery of P. cinnamomi, P. citricola, P. europaea Hansen and Jung, and P. cambivora, with P. cinnamomi recovered from 69.4% of the sites (Balci et al. 2007). Species of Phytophthora in ITS Clade 6, including, P. gonapodyides and P. chlamydospora Hansen, appear to have a widespread distribution with common recovery from irrigation systems (Copes et al. 2015, Hansen et al. 2015, Yang et al. 2013). Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 178 We are unaware of any data on populations of Phytophthora in riparian urban ecosystems of Mississippi. Copes et al. (2015) published survey results of Phytophthora spp. that occurred in irrigation ponds located in Alabama and Mississippi. The results of Nechwatal et al. (2015) demonstrated clear differences in Oomycete populations when comparing flooded and dry riparian environments. These data, however, may not reflect population data for urban drainage environments. Urban drainage areas often flood due to surface run-off during wet seasons, and are almost completely dry during the hottest parts of each year. Within these unusual and often heavily impacted habitats, little is known about Phytophthora spp. diversity. Therefore, we conducted a study to determine temporal occurrences and survival potential of Phytophthora populations at a forested riparian survey site that included a connecting drainage ditch and stream within an urban environment. Field-site Description The study site included a 50-m segment of an intermittent wet–dry drainage ditch that connects with the continuously flowing Hog Creek in Rankin County, MS. Geographical coordinates for the confluence of the drainage ditch and Hog Creek are 32°20'18.4"N, 90°05'50.2"W. Hog Creek flows through a mesic oak–pine forest community for ~1.82 km before it converges with the Pearl River (Fig. 1). During periods of heavy rainfall, the drainage ditch and Hog Creek crest up to 3 m with intermittent flooding into the surrounding forest that contains trees and other woody vegetation. Methods To determine the presence and survival of Phytophthora spp. present at the Hog Creek site, we employed 5 methods from 3 US Department of Agriculture, Animal and Plant Inspection Service (APHIS) protocols, including soil baiting (USDA APHIS 2010), water-sampling protocols (USDA APHIS 2014a), vegetation-sampling inspection and sampling protocol for nurseries (USDA APHIS 2014b), and selective-media formulas PARPH-V8 and PAR-V8 (Jeffers et al. 1986). We conducted stream-baiting in accordance with the APHIS in vitro water sampling with host-material leaf baits–bottle-of-bait (BOB) technique, updated 22 March 2014. On each sampling date, we measured the water temperature at the sampling location. We added 800 ml of stream water in 100-ml aliquots to 1-L plastic bottles: 2 bottles at the mid-point of the drainage ditch, 1 bottle at 25 m upstream from the convergence of Hog Creek and 1 bottle directly at the convergence of the drainage ditch with Hog Creek (Fig. 1). We followed the same methods to collect 2 bottles of water from Hog Creek: 1 at the confluence with the drainage ditch and the other 25 m downstream from that point. We added 10 circular leaf-disks (0.63 cm in diameter) and 1 intact, 1-y-old, healthy Rhododendron x Cunningham’s White (a rhododendron cultivar) leaf to each of the bottles immediately after water sampling. We excised the rhododendron leaves from plant stock located on the Mississippi State University campus within 8 h of sampling. We incubated the water Southeastern Naturalist 179 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 samples with intact rhododendron leaves and leaf pieces for 72 h at 20 °C in the dark, then processed them according to the literature cited above. We initiated the water-filtering portion of the study in October 2014, following the APHIS water sampling for filtration protocol, updated 22 March 2014. We filled two 1-L plastic bottles with 800 ml of stream water in 100 ml aliquots, collected 25 m upstream from confluence of the drainage ditch with Hog Creek. We placed the water samples in a cooler and immediately filtered them upon return to the laboratory, about 4 h after collection. We employed a 47-mm, 250-ml Nalgene (Nalge, Rochester, NY) filter-funnel with clamp and a hand-operated vacuum pump to process the samples. We filtered ~100 ml of clear water per filter and 50–75 ml Figure 1. Soil- and vegetation-sampling sections of the drainage ditch. Sampling sections 1–4 are in the alluvial plain, sections 5 and 6 are located in the floodplain. Lines indicating the 4 stream-baiting sampling sites are labeled. Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 180 for more turbid samples. We filtered clean water through a 3-μm Nuclepore polycarbonate membrane (GE Healthcare Bio-Sciences, Pittsburgh, PA) and more turbid water through a 5-μm Durapore polyvinylidene fluoride membrane (MilliPore Sigma, Billerica, MA), inverted the filters on PARPH-V8 media, and incubated them at 20 °C in the dark for 72 h. We observed colony morphology under a dissecting microscope, and transferred the leading edge of mycelium onto PAR-V8 media for isolation. If sporangia formed in media, zoosporogenesis was observed, and if zoospores were ejected into a vesicle prior to a cleavage, then we did not consider that isolate to be a species of Phytophthora. After 10 d of incubation in the dark at 20 °C, we followed the procedure outlined by Kong et al. (2004) to swab, process for DNA extraction, and PCR the sample mycelium, as described below. We conducted soil baiting in accordance with the APHIS soil and container mix protocol, revised November 2010. We collected soil samples with a 2 cm x 10 cm Oakfield soil sampler (Oakfield Apparatus, Oakfield, WI) disinfected between each sample using full-strength Lysol spray (Reckitt Benckiser LLC, Parsippany, NJ). We divided the sampling area of the drainage ditch into 6 sampling sections: four 25 m x 5 m riparian sections and two 50 m x 5 m floodplain sections (Fig. 1). We measured soil temperature at 4 locations 30 cm apart in each sampling section, and averaged the readings. In each sampling section, we collected 15 soil cores in a staggered manner: 1 close to the waterline, one 50 cm downstream and 25 cm away from the waterline, one 50 cm further downstream and back at the waterline, and so on, until all sampled were obtained. We combined the 15 core samples from each sampling section in a 3.8-L Ziploc bag (Dow Chemical Company, Racine, WI) to create 1 composite sample, for a total of 90 core samples constituting about 6 L of soil per sampling trip. The six 1-L composite soil samples were thoroughly mixed, placed in a cooler, and transported to the laboratory for processing according to the literature mentioned above. Vegetation samples included symptomatic leaves, twig and branch cankers, and roots exhibiting necrosis or water-soaked lesions. We visually inspected every plant in the sampling area on every sampling trip. We consulted the USDA Natural Resources Conservation Service Plants Database (https://plants.usda. gov), and the Mississippi Forestry Commission’s publication Mississippi Trees (http://www.fwrc.msstate.edu/pubs/ms_trees.pdf) to identify the plant species present. Each plant specimen was identified to genus, placed in a 1.5-L Ziploc bag labeled with the soil-sampling section in which it was collected, and placed in a cooler to be brought back to the laboratory. We surface-sterilized all vegetation samples with 70% ethanol for 5 min before culturing on PARPH-V8 media and processing as described below. We cultured all rhododendron bait leaves and pieces, and all symptomatic vegetation on PARPH-V8 media at 20 C° in the dark for up to 7 d, then transferred the leading edge of colony mycelium onto PAR-V8 media for isolation. We followed the procedure of Kong et al. (2004) to swab the mycelium and then extracted the DNA according to the Nucleospin Plant II Kit (Machery Nagel, Duren, Germany) protocol. We amplified the DNA with Phytophthora-specific Southeastern Naturalist 181 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 A2-forward (ACTTTCCACGTGAACCGTTTCAA) and I2-reverse primers (GATATCAGGTCCAATTGAGATGC) from the 3' end of the 18S gene to the 5' end of the 28S gene, amplifying a size between 752 bp and 832 bp in length that included the ITS1-5.8S-ITS2 region of the rDNA (Drenth et al. 2006, White et al. 1990). The PCR amplicon from above was digested with 3 different restriction enzymes: Msp1, Rsa1, and Taq1 (Promega, Madison, WI). We performed digestion in 20-μl volumes, consisting of 10 μl PCR product, 2 μl 10x appropriate restriction buffer, 6.8 μl water, 0.2 μg bovine serum-albumin, and 1μl restriction enzyme. We incubated samples in a water bath for 2–3 hours at 37 °C for Msp1 and Rsa1, and 65 °C for Taq1. After digestion, we froze the samples at -20 °C until we ran them on a 2% agarose gel with a 100-bp ladder to obtain the digest fingerprints (Drenth et al. 2006). We compared study samples to digests of known species, including: P. mississippiae, isolate 57J3, obtained from W. Copes (USDA Agricultural Research Service, Poplarville, MS) and P. citricola, P. cryptogea, P. palmivora, P. citrophthora, P. nicotianae Breda de Haan, P. cinnamomi, and P. gonapodyides, obtained from S. Jeffers (Clemson University, Clemson SC). We prepared select PCR products for sequencing following the protocol of the Eurofin–Operon sequencing center (Louisville, KY, http://www.operon.com/ products/downloads/Sample_Submission_Guidelines.pdf). Following clean-up, we employed FinchTV v1.4.0 (Geospiza, Seattle, WA) to align sequence data to generate a consensus sequence, and used BLASTnt to compare our sequence with samples in the NCBI database. The sequences have been deposited in GenBank under the accession numbers shown in the species descriptions in Results. As additional confirmation of the taxon identifications, we conducted a bayesian phylogenetic analysis on a data set consisting of (1) sequences generated for this study, (2) sequences of Phytophthora taxa in GenBank that were identified as strong matches to the sequences collected for this study, and (3) sequences from GenBank of other Phytophthora taxa to account for wider variation within the genus that may not have been identified in BLAST searches. Species identifications for sequences selected from GenBank were confirmed by expert opinion, published papers where the sequence was referenced, or comparison to the Phytophthora Database (http:// www.phytophthoradb.org). We also selected 3 sequences of Peronospora to serve as outgroup taxa and to demonstrate that the sequences derived in this study are most closely related to Phytophthora. These outgroup taxa included: Peronospora Corda spp. (AY831719), Peronospora tabacina D.B. Adam (AY198289), and Peronospora trifoliorum de Bary (EF174967). We used the web version of MAFFT (Katoh et al. 2005), which is provided by the CBRC (http://mafft.cbrc. jp/alignment/server/), with the AUTO option to align all sequences. The resulting alignment, which contained 1944 characters, was subjected to an assessment by jModeltest2 (Daribba et al. 2012, Guindon et al. 2003) to determine the best-fitting model of molecular evolution under the BIC. We restricted the model selection to 3 categories to identify the best-fitting model that could be implemented in MrBayes. We conducted this analysis in the Cipres Science Gateway (Miller et al. 2010). After HKY+G was selected as the best-fitting model, we performed a Bayesian Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 182 phylogenetic analysis on the data set using MrBayes v. 3.2.3 (Ronquist et al. 2012) in the Cipres Science Gateway (Miller et al. 2010). Markov chain Monte Carlo was conducted for 5 million generations with a sampling frequency of 1000, after which the split standard deviation was less than 0.007. We set a burn-in of 1250 trees prior to determining the posterior probability of the trees with the highest likelihood. The consensus tree is reported with posterior probability indicated as support for clades. We computed relative frequencies of Phytophthora occurrence for each fieldsampling method, and subjected the data to analysis of variance (ANOVA) in the general linear models procedure (Proc GLM) of Statistical Analysis System (SAS) software. The experimental design was a split plot, with Phytophthora species as the main plot and sampling dates as repeated measures (relative frequency = number of Phytophthora recoveries/sampling date). We compared percent frequencies between sampling dates and sampling methods, the totals from 17 December 2014, and using BOB leaf-disk, BOB intact-leaf, and filtered-water isolations. We conducted Fisher’s protected least significant difference test (LSD; (P < 0.05) to separate means. Results We collected and identified a total of 11 Phytophthora spp. over 14 sampling dates and 5 field-sampling methods between March 2014 and February 2015 (Table 1). The phylogenetic analysis indicated a reciprocal monophyly of all identified species within Phytophthora and strong support for most species-level clades (Fig. 2), which suggests that our identification of taxa collected in this study on the basis of ITS data is accurate. Species descriptions for the 11 taxa are discussed below. Of those, the most common species was P. chlamydospora (present in 6.3% of all samples), whereas we detected P. mississippiae, P. ramorum, and P. cinnamomi in 5.0–5.8% of samples (Table 2). The other 7 taxa were similar in frequency (P ≥ 0.05), varying from 1.8% to 0.1% occurrence. Based on sampling methods, P. chlamydospora had significantly greater frequencies than other the taxa in the soil-baiting samples, and non-significantly but numerically greater frequencies in the BOB leaf-disk samples. Furthermore, P. mississippiae was isolated or identified at numerically greater levels from water-filtration samples, and we detected P. cinnamomi more frequently in the BOB intact-leaf sampling method. We did not analyze vegetation-sampling data because of irregular sampling based on phenology and low number of actual confirmations overall during the st udy. We compared the total number of isolates across species and sampling methods (Table 3). Overall, the BOB intact leaf and water-filter sampling methods had the largest percentages of Phytophthora confirmations; the highest occurred on 17 December 2014 at 36.4% and 27.3%, respectively. On this date, the water Figure 2 (see page 10). Phylogenetic relationships among samples collected in this study. Sequences taken from GenBank are shown in bold. Posterior probability values resulting from a bayesian phylogenetic analysis are shown on branches as * = 0.90–0.94; ** = 0.95–0.98; *** = 0.99–1.0. Southeastern Naturalist 183 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 Table 1. All strains of Phytophthora recovered during 2014–2015 surveys from a urban forested site in Rankin County, MS. [Continued on next page.] Date Temperature Genbank Species Strain recovered Recovery method Geographical coordinates (°C) accession # P. cambivora T4 Vaccinium 5/6/2014 Vaccinium arboreum leaf 32°20'18.8"N, 90°05'48.9"W 21.9 air KY053259 P. cinnamomi T11 B.8 12/17/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.7 water KY696607 P. cinnamomi T11 B.15 12/17/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.7 water KY696595 P. cinnamomi T11 A.3 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696598 P. cinnamomi T11 A.10 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696599 P. cinnamomi T11 A.16 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696596 P. cinnamomi T11 A.18 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696602 P. cinnamomi T11 A.24 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696601 P. cinnamomi T11 A.31 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696608 P. cinnamomi T11 A.32 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696603 P. cinnamomi T11 A.34 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696600 P. cinnamomi T11 I.L. A.6 12/17/2014 Drainage ditch BOB intact leaf 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696597 P. cinnamomi T13 Willow 1 1/27/2014 Salix nigra leaf 32°20'18.5"N, 90°05'50.0"W 20.5 air KY696604 P. cinnamomi T13 Willow 2 1/27/2014 Salix nigra root 32°20'18.5"N, 90°05'50.0"W 9.6 soil KY696605 P. cinnamomi T6 Swamp Chestnut 2 6/3/2014 Quercus michauxii petiole 32°20'19.0"N, 90°05'49.3"W 31.6 air KY696606 P. chlamydospora T5 Ditch Azalea 2 5/20/2014 Rhododendron canescens leaf 32°20'19.0"N, 90°05'49.7"W 31.1 air KY696584 P. chlamydospora T6 Quercus sapling 6/3/2014 Quercus nigra leaf 32°20'18.8"N, 90°05'48.9"W 31.6 air KY696593 P. chlamydospora T8 BOB A.3 10/31/2014 Drainage ditch BOB leaf pieces 32°20'18.4"N, 90°05'50.2"W 6.8 water KY696588 P. chlamydospora T9 Soil 1 11/14/2014 Soil 32°20'18.8"N, 90°05'48.9"W 16.1 soil KY696583 P. chlamydospora T5 Soil 2.1 5/20/2014 Soil 32°20'18.8"N, 90°05'48.9"W 22.9 soil KY696592 P. chlamydospora T10 Soil 2 12/5/2014 Soil 32°20'18.8"N, 90°05'48.9"W 9.4 soil KY696586 P. chlamydospora T10 Soil 2.1 12/5/2014 Soil 32°20'18.8"N, 90°05'48.9"W 9.4 soil KY696591 P. chlamydospora T10 A.5 12/5/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.2 water KY696589 P. chlamydospora T10 A.6 12/5/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.2 water KY696578 P. chlamydospora T10 A.15 12/5/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.2 water KY696590 P. chlamydospora T10 A.29 12/5/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.2 water KY696582 P. chlamydospora T11 I.L. A.1 12/17/2014 Drainage ditch BOB intact leaf 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696579 P. chlamydospora T11 B.24 12/17/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.7 water KY696580 P. chlamydospora T11 BOB B.5 12/17/2014 Drainage ditch BOB leaf pieces 32°20'18.9"N, 90°05'49.7"W 6.7 water KY696581 P. chlamydospora T12 A.5 1/13/2015 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.2 water KY696578 P. chlamydospora T12 I.L. A.1 1/13/2015 Drainage ditch BOB intact leaf 32°20'18.4"N, 90°05'50.2"W 10.2 water KY696594 P. chlamydospora T12 Soil 6.2 1/13/2015 Soil 32°20'18.8"N, 90°05'48.9"W 9.2 soil KY696585 Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 184 Table 1, continued. Date Temperature Genbank Species Strain recovered Recovery method Geographical coordinates (°C) accession # P. citrophthora T4 H.C. Azalea 2 5/6/2014 Rhododendron canescens leaf 32°20'19.8"N, 90°05'51.6"W 28.3 air KY696609 P. citrophthora T6 I.L. C.2 6/3/2014 Drainage ditch BOB intact leaf 32°20'19.3"N, 90°05'51.1"W 26.1 water KY696610 P. cryptogea T3 Hickory 4/16/2014 Carya tomentosa leaf 32°20'18.7"N, 90°05'48.7"W 18.9 air KY696611 P. cryptogea T3 Sweet gum 4/16/2014 Liquidambar styraciflua leaf 32°20'18.9"N, 90°05'49.2"W 18.9 air KY696612 P. mississippiae T6 I.L. B.1 6/3/2014 Drainage ditch BOB intact leaf 32°20'18.9"N, 90°05'49.7"W 26.1 water KY696618 P. mississippiae T6 Willow necrosis 6/3/2014 Salix nigra leaf 32°20'18.5"N, 90°05'50.0"W 31.6 air KY696627 P. mississippiae T8 A.13 10/31/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.8 water KY696623 P. mississippiae T8 BOB B.7 10/31/2014 Drainage ditch BOB leaf pieces 32°20'18.9"N, 90°05'49.7"W 6.8 water KY696617 P. mississippiae T8 BOB B.8 10/31/2014 Drainage ditch BOB leaf pieces 32°20'18.9"N, 90°05'49.7"W 6.8 water KY696622 P. mississippiae T8 B.3 10/31/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.8 water KY696615 P. mississippiae T8 B.4 10/31/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.8 water KY696620 P. mississippiae T8 Soil 5.5 10/31/2014 Soil 32°20'18.8"N, 90°05'48.9"W 6.1 soil KY696619 P. mississippiae T9 A.13 11/14/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 17.1 water KY696621 P. mississippiae T10 A.22 12/5/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.2 water KY696616 P. mississippiae T11 BOB A 12/17/2014 Drainage ditch BOB leaf pieces 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696624 P. mississippiae T11 A.4 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696626 P. mississippiae T11 A.13 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696625 P. mississippiae T11 B.19 12/17/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.7 water KY696614 P. mississippiae T11 B.21 12/17/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.7 water KY696613 P. oaksoil taxon T7 Wetwood 6/17/2014 Quercus nigra trunk 32°20'19.8"N, 90°05'51.6"W 32.7 air KY696628 P. oaksoil taxon T13 BOB C.1 1/27/2015 Hog Creek BOB leaf pieces 32°20'19.3"N, 90°05'51.1"W 10.8 water KY696629 P. oaksoil taxon T13 BOB C.2 1/27/2015 Hog Creek BOB leaf pieces 32°20'19.3"N, 90°05'51.1"W 10.8 water KY696630 P. ramorum T3 Vaccinium 4/16/2014 Vaccinium arboreum leaf 32°20'18.8"N, 90°05'48.9"W 18.9 air KY696631 P. ramorum T11 A.32 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696633 P. ramorum T11 A.34 12/17/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.7 water KY696632 P. ramorum T11 B.5 12/17/2014 Filtered water 32°20'18.9"N, 90°05'49.7"W 6.7 water KY696634 P. riparia T8 A.3 10/31/2014 Filtered water 32°20'18.4"N, 90°05'50.2"W 6.8 water KY696635 P. riparia T8 BOB B.1 10/31/2014 Drainage ditch BOB leaf pieces 32°20'18.9"N, 90°05'49.7"W 6.8 water KY696636 P. rosacearum T6 Soil 2.2 6/3/2014 Soil 32°20'18.8"N, 90°05'48.9"W 24.6 soil KY696637 P. syringae T13 A.5 1/27/2015 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.8 water KY696639 P. syringae T13 A.7 1/27/2015 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.8 water KY696638 P. syringae T13 A.25 1/27/2015 Filtered water 32°20'18.4"N, 90°05'50.2"W 10.8 water KY696640 P. syringae T13 B.8 1/27/2015 Filtered water 32°20'18.9"N, 90°05'49.7"W 10.8 water KY696641 Southeastern Naturalist 185 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 Figure 2. Phylogenetic relationships. [See page 7 for full caption.] Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 186 Table 3. Percent isolation frequencies of 11 Phytophthora spp. across 14 sampling dates and by sampling method, including vegetation from an urban forested site in Rankin County, MS. Percent frequencies are based on total number of identifications (hits) across all sampling methods including vegetation (not shown). BOB = bottle-of-bait sampling method from USDA APHIS water-sampling protocol, 22 March 2014. Means within a column that do not share a common superscripted letter are significantly different according to Fisher’s Protected LSD (P ≤ 0.05). N/A = water filtration was not performed on these dates. Sampling Total % frequency Field sampling methods % frequencies date across taxa BOB leaf disks BOB intact leaf Soil leaf disks Water filter 3/18/14 less than 1.0B less than 1.0 less than 1.0B less than 1.0B N/A 4/2/14 less than 1.0B less than 1.0 less than 1.0B less than 1.0B N/A 4/16/14 less than 1.0B less than 1.0 less than 1.0B less than 1.0B N/A 5/6/14 less than 1.0B less than 1.0 less than 1.0B less than 1.0B N/A 5/20/14 less than 1.0B less than 1.0 less than 1.0B less than 1.0B N/A 6/3/14 1.7B less than 1.0 2.5B 1.5AB N/A 6/17/14 less than 1.0B less than 1.0 less than 1.0B less than 1.0B N/A 10/31/14 4.5B 6.8 4.5B 1.5AB 9.1B 11/14/14 1.1B less than 1.0 2.3B 1.5AB less than 1.0B 12/5/14 4.0B less than less than 1.0 9.1B 4.5A less than 1.0B 12/17/14 14.7A 9.0 36.4A less than 1.0B 27.3A 1/13/15 2.3B less than 1.0 4.5B 1.5AB less than 1.0B 1/27/15 3.4B 4.5 6.8B less than 1.0B 4.6B 2/12/15 2.3B 2.3 2.3B less than 1.0B 9.1B LSD 5.5 6.3 19.3 4.1 14.4 Table 2. Comparison of total percent isolations of 11 Phytophthora spp. across and between sampling methods isolated over 12 months from an urban forested site in Rankin County, MS. Percent frequencies are based on total number of identifications (hits) across all sampling methods including vegetation. BOB = bottle-of-bait sampling method from USDA APHIS water-sampling protocol, 22 March 2014. Means within a column that do not share a common superscripted letter are significantly different according to Fisher’s Protected LSD (P ≤ 0.05). Phytophthora Total % frequency Field sampling methods % frequencies taxa across taxa BOB leaf disks BOB intact leaf Soil leaf disks Water filter P. cambivora 0.1D less than 1.0 less than 1.0 less than 1.0 less than 1.0 P. chlamydospora 6.3A 5.4 8.9 5.9A 3.6 P. cinnamomi 5.0ABC 1.8 14.0 less than 1.0B 7.1 P. citrophthora 0.5CD less than 1.0 1.8 less than 1.0B less than 1.0 P. cryptogea 0.3D less than 1.0 less than 1.0 less than 1.0B less than 1.0 P. mississippiae 5.8A 3.6 12.5 1.2B 14.3 P. ramorum 5.4AB 3.6 11.0 less than 1.0B 11.0 P. riparia 0.9BCD 1.8 1.8 less than 1.0B less than 1.0 P. rosacearum 0.5CD less than 1.0 less than 1.0 1.2B less than 1.0 P. oaksoil taxon 0.9BCD 3.6 less than 1.0 less than 1.0B less than 1.0 P. syringae 1.8BCD less than 1.0 5.4 less than 1.0B 3.6 LSD 4.6 6.5 20.3 3.3 18.7 Southeastern Naturalist 187 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 temperature was 6.7 °C, and we recovered P. chlamydospora (3x), P. cinnamomi (11x), P. mississippiae (5x), and P. ramorum (8x, including morphological confirmations) from every sampling method except soil. The sampling date with the 2nd-highest prevalence was 31 October 2014, when water and soil temperatures were 6.8 °C and 6.1 °C, respectively, frequencies ranged from 4.5% to 9.1% across all sampling methods, and we recovered P. mississippiae (6x), P. chlamydospora (1x), and P. riparia (2x). Soil sampling had the highest percentage frequency on 5 December 2014 (4.5%), when the soil temperature was 9.4 °C; this date also had the 3rd-highest frequency across taxa at 4.0–9.1% for all sampling methods, and we recovered P. chlamydospora (6x), and P. mississippiae (1x). Species Descriptions Phytophthora cambivora Species-isolation method/confirmations. We retrieved this species from necrotic lesions on a leaf tip of a Vaccinium arboreum Marshall (Farkleberry) located in zone 1 of the drainage ditch on 18 March 2014 (Table 1). Morphological data. We observed no reproductive structures; thus we could not verify the identification of this species via morphology, but identification was confirmed through ITS sequence analysis. Molecular data. The sequence from strain T4 Vaccinium (KY696577) was 99% similar (100% coverage) to P. cambivora (KU053259). These sequences cluster together with 100% posterior probability (PP) and are sister to the clade containing P. cinnamomi (Fig. 2). Comments. Phytophthora cambivora is a heterothallic species in Clade 7 of Martin et al. (2014). It is soil-borne and has been documented in low frequency from oak forests in the eastern and central US, as well as from ornamental nursery plants in Maryland (Balci et al. 2007, Bienapfl et al. 2014). Phytophthora cinnamomi Species-isolation method/confirmations. We isolated and identified this species from ITS sequence analysis 14 times throughout the study. We isolated P. cinnamomi from samples at a wide range of temperatures—from 6.7 °C ,when recovered from stream water on 7 December 2014, to 26.1 °C, when retrieved from a Q. michauxii Nutt. (Swamp Chestnut Oak) petiole on 3 June 2014 (Table 1). Morphological data. The mycelium has distinctive coralloid hyphae and abundant knobby to globular, clustered hyphal swellings. Prolific globose, thin-walled chlamydospores were formed, ranging from 35 μm to 45μm in size (Fig. 3). Non-papillate sporangia were ovoid, obpyriform, or ellipsoid with a slight apical thickening. Sporangia were not deciduous and observed to be only borne singularly and terminally. We noted aerial mycelium in PAR-V8 agar. Molecular data. Consensus data from strains T11 B.8 (KY696607), T11 B.15 (KY696595), T11 A.3 (KY696598), T11 A.10 (KY696599), T11 A.16 (KY696596), T11 A.18 (KY696602), T11 A.24 (KY696601), T11 A.31 (KY696608), T11 A.32 (KY696603), T11 A.34 (KY696600), T11 I.L. A.6 (KY696597), T13 Willow 1 Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 188 (KY696604), T13 Willow 2 (KY696605), and T6 Swamp Chestnut 2 (KY696606) were 100% similar (100% coverage) to strain ATCC MYA-4058 (FJ746646). All of these sequences are clustered in a clade with 100% PP (Fig. 2). Comments. Phytophthora cinnamomi is a heterothallic species in Clade 7, with its closest relative P. parvispora Scanu and Denman (Martin et al. 2014). In Mississippi, P. cinnamomi is a devastating root-rot pathogen of Vaccinium corymbosum L. (Highbush Blueberry) (Smith 2012). The pathogen has also been associated with root and crown rot of Prunus persica (L.) Batsch (Peach) trees and Pinus spp. Figure 3. Morphological data. (A) Terminal chlamydospore of P. chlamydospora, bar = 40 μm; (B) Chlamydospores of P. ramorum, bar = 20 μm (C) Chlamydospores of P. cinnamomi, bar = 20 μm; (D) Nonpapillate sporangia of P. oaksoil taxon, bar = 20 μm; (E) Semi-papillate sporangia of P. ramorum, bar = 20 μm; (F) Nonpapillate sporangia of P. mississippiae, bar = 10 μm; (G) Internal proliferating sporangiophore of P. cryptogea, bar = 10 μm; (H) Papillate sporangia of P. citrophthora, bar = 10 μm . Southeastern Naturalist 189 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 (yellow pines) in Mississippi (Haygood et al. 1986, Mistretta 1984). P. cinnamomi has also been isolated from eastern oak species in both Florida and the central eastern US, contributing to the decline of Quercus phellos L. (Willow Oak), Quercus rubra L. (Northern Red Oak), and Quercus alba L. (White Oak) (Barnard 2006, Eggers et al. 2012, Robin et al. 2012). Phytophthora chlamydospora Species-isolation method/confirmations. This was the most common species recovered in our study, with 17 isolations using ITS sequence data throughout the study. We retrieved all of the isolates from water and soil when the temperatures were between 6.7 °C and 22.9 °C (Table 1). According to Hansen et al. (2015), optimum-growth temperature is 25–28 °C, with maximum growth at 36–37 °C. Morphological data. No sexual gametes were formed in a single culture. Sporangia were simple, mostly ovoid but occasionally obpyriform, non-papillate with a slight apical thickening, and non-caducous. We observed globose to subglobose hyphal swellings, along with a number of terminal and intercalary chlamydospores, a few of which were measured at 38 μm x 38 μm (Fig. 3). We noted coiled mycelium in PAR-V8 media. Molecular data. When entered into the NCBI database, the consensus ITS sequence of strains T5 Ditch Azalea 2 (KY696584), T6 Quercus Sapling (KY696593), T8 BOB A.3 (KY696588), T9 Soil 1 (KY696583), T5 Soil 2.1 (KY696592), T10 Soil 2 (KY696586), T10 Soil 2.1 (KY696591), T10 A.5 (KY696589), T10 A.6 (KY696578), T10 A.15 (KY696590), T10 A.29 (KY696582), T11 I.L. A.1 (KY696579), T11 B.24 (KY696580), T11 BOB B.5 (KY696581), T12 A.5 (KY696578), T12 I.L. A.1 (KY696594), and T12 Soil 6.1 (KY696585) showed 100% coverage and 100% similarity with 1 of the strains used to define this species, annotated as KC734370. All of these sequences are clustered in a clade with 98% PP (Fig. 2). Comments. Phytophthora chlamydospora, formally known as P. taxon pgchlamydo, is found worldwide in streams and forest soils and may be the 2nd most abundant Phytophthora species in the world; it is a weak pathogen of woody plants (Hansen et al. 2007). Phytophthora chlamydospora is heterothallic within Clade 6, with its closest neighbor P. pinifolia Durán, Gryzenh, and Wingf (Hansen et al. 2007). Its slow growth, aquatic habitat, and ability to tolerate high temperatures associates it with most other Clade 6 species, although it is distinguished from the rest of the clade by the production of asexual chlamydospores (Hansen et al. 2015). Phytophthora citrophthora Species-isolation method/confirmations. We isolated this species from necrotic leaf-lesions collected from a Rhododendron canescens (Michx.) Sweet (Mountain Azalea) along Hog Creek on 6 May 2014, and from a BOB intact leaf from Hog Creek on 3 June 2014 when the water temperature was 26.1 °C (Table 1). According to Matheron and Matejika (1992), P. citrophthora has an optimum growth temperature of 25 °C, with abundant sporangia production between 20 °C and 30 °C. Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 190 Morphological data. No sexual gametangia were produced in culture, and we observed only a few chlamydospores, which were both terminal and intercalary, and about 38 μm x 35μm in size. Papillae on sporangia were conspicuous, but sporangia lacked caducity, with variable shapes ranging from spherical, ovoid, to ellipsoid, and were borne singularly and sympodially (Fig. 3). Discharge-pore width was small, measuring 8 μm in 1 sporangiophore. The culture formed sparse aerial mycelium after 10 d at 20 °C on PAR-V8 medium. Molecular data. The consensus ITS sequence from strains T4 H.C. Azalea 2 (KY696609) and T6 I.L. C.2 (KY696610) showed 100% coverage and 99% similarity with P. citrophthora CBS #111726 culture collection (HQ643206). All sequences identified as P. citrophthora cluster in a strongly supported clade (100% PP) with other P. citrophthora and P. himasilva (Fig. 2). Comments. Phytophthora citrophthora is a heterothallic species in Clade 2, with its closest relative P. botryosa Chee (Martin et al. 2014). Uddin et al. (2007) analyzed 19 isolates of P. citrophthora from different geographical areas, hosts, and sampling periods, with genetic relationships among the isolates forming 1 clade distinct from other species of Phytophthora, and with no variations in the ITS region. It is commonly recovered from nursery-irrigation water in ornamental plant nurseries as well as forest streams in the eastern US due to its association with diverse plant hosts (Bush et al. 2006, Donahoo and Lamour 2008, Ferguson and Jeffers 1999, Hulvey et al. 2010). Phytophthora cryptogea Species-isolation method/confirmations. We isolated this species from Carya tomentosa (Lam.) Nutt. (Mockernut Hickory) and Liquidambar styraciflua L. (Sweetgum) leaves on 16 April 2014; both were located in the drainage ditch (Table 1). The Mockernut Hickory leaf had dark leaf spots (3–4mm), while the Sweetgum leaf had necrotic leaf edges with a halo of chlorotic tissue. Morphological data. Phytophthora cryptogea produces abundant obpyriform to ovoid, non-papillate, non-caducous sporangia, many deriving from internal proliferation of the sporangiophore (Fig. 3). Chlamydospores and hyphal swellings were absent. We noted the presence of aerial mycelium on PAR-V8 agar. Molecular data. When entered into the NCBI database, the consensus ITS sequence from strains T3 Hickory (KY696611) and T3 Sweet gum (KY696612) showed 100% coverage and 99% similarity with P. cryptogea (HQ455574). These sequences are clustered in a clade with 100% PP (Fig. 2). Comments. Phytophthora cryptogea is a heterothallic species in Clade 8, with its closest relative P. erythroseptica Tucker (Martin et al. 2014). Martin et al. (2014) and Safaiefarahani et al. (2015) suggested that the molecular deviation in P. cryptogea lineages validate the designation of the P. cryptogea clade into distinct species. Phytophthora cryptogea has been isolated from over 100 species of woody plants in 23 different families, and is a significant pathogen in domestic floral and ornamental greenhouses (Bush et al. 2006, Erwin and Ribeiro 1996, Ferguson et al. 1999, Hwang and Benson 2005, MacDonald et al. 1994, Olson et al. 2011). Southeastern Naturalist 191 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 Phytophthora mississippiae Species-isolation method/confirmations. We identified this species 15 times from ITS gene-region sequencing during the study, with 10 sequences derived from isolated cultures (Table 1). Morphological data. We observed no sexual gametes. Sporangia formed after submerging 7-d–old V8 agar plugs in a non-sterile soil extract solution (NSSES, 15 g soil/1 L distilled water) under fluorescent lighting at room temperature for 8 h. We also inoculated rhododendron disks and floated them on both NSSES and distilled water with moderate sporangia development. Sporangia were obpyriform and ovoid, borne singularly, non-papillate with a slight apical thickening, and not deciduous (Fig. 3). We observed successful zoosporogenesis from the sporangium with no vesicle formation prior to zoospore cleavage. No chlamydospores formed. Mycelium on PAR-V8 media after 10 d at 20 °C was appressed. Molecular data. When entered into the NCBI database, the consensus ITS sequence from strains T6 I.L. B.1 (KY696618), T6 willow necrosis (KY696627), T8 A.13 (KY696623), T8 BOB B.7 (KY696617), T8 BOB B.8 (KY696622), T8 B.3 (KY696615), T8 B.4 (KY696620), T8 Soil 5.5 (KY696619), T9 A.13 (KY696621), T10 A.22 (KY696616), T11 BOB A (KY696624), T11 A.4 (KY696626), T11 A.13 (KY696625), T11 B.19 (KY696614), and T11 B.21 (KY696613) showed 100% coverage and 99% similarity with KP780453, which is a strain from the American Type-Culture Collection MYA-4946. These sequences form a clade, but it is not strongly supported (50% PP; Fig. 2). Although this clade lacks strong support, we believe the identification is accurate due to similar morphological characteristics and identical banding patterns to a known sample of P. mississippiae in the enzyme digest. The enzyme digest comparison used a known isolate (exo-type: 57J3) reported in the type description of P. mississippiae from W. Copes ( USDA/ARS, Poplarville, MS). Comments. Phytophthora mississippiae is a recently described species that was isolated from irrigation water at an ornamental plant nursery in Mississippi in 2012. Phytophthora mississippiae is heterothallic in Clade 6 with P. thermophila Jung, Stukely, and Burgess as a closest relative (Yang et al. 2013). Yang et al. (2014) recognized P. xstagnum as a hybrid species of P. chlamydospora and a species genetically close to P. mississippiae, with similar morphological characteristics, and a comparable mitochondrial cox spacer sequence, ITS region, and beta-tubulin sequence analysis. Phytophthora oaksoil taxon Species-isolation method/confirmations. Phytophthora oaksoil taxon was originally observed and isolated by Braiser et al. (2003) from soil in France. We isolated P. oaksoil taxon on 27 January 2015 from a BOB leaf disk collected from Hog Creek downstream from the confluence with the drainage ditch, and from discolored lesions on a trunk of a Quercus nigra L. (Water Oak) tree on 17 June 2014, located adjacent to Hog Creek and periodically subjected to flooding (Fig. 4). When we cut away at the wound with a knife at the point of callus, there were no signs Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 192 of cambium necrosis or disease progression into the vascular tissue. Although we made numerous attempts to culture samples from the oak tree, this was the only Phytophthora sp. recovered. We recovered this species from Hog Creek when the water temperature was 10.8 °C (Table 1). Braiser et al. (2003) described moderate growth on carrot agar at 25 °C, with the upper limit for growth at 33 °C. Morphological data. We observed no sexual gametes. Sporangia were ovoid, non-papillate, non-caducous, and varied in size, with the majority borne terminally, Figure 4. Field Sampling Site. (A) Alluvial plain and sampling area of the drainage ditch during February. (B) Arrow points at the convergence of the drainage ditch and Hog Creek. (C) Arrow points at a wound on a Quercus nigra tree located along the banks of Hog Creek. Southeastern Naturalist 193 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 and a few from internally proliferated sporangiophores (Fig. 3). We measured a couple of exit pores at 11 μm and 15 μm. No chlamydospores formed in PAR-V8 media. Mycelium on PAR-V8 media after 10 d at 20 °C was appressed. Molecular data. The consensus ITS sequence from strains T7 Wetwood (KY696628), T13 BOB C.1 (KY696629), and T13 BOB C.2 (KY696630) from this investigation showed 100% coverage and 100% similarity with KJ666740, which is also identified as a P. oaksoil taxon. These sequences cluster within a clade strongly supported by 100% PP (Fig. 2). Comments. In addition to the original observation in France, P. oaksoil taxon has been recovered in low frequency from streams in Oregon dominated by Alnus sp. (alder) and Salix sp. (willow) (Reeser et al. 2011). Sims (2014) described the ubiquitous P. obrutafolium from Oregon streams as genetically similar to the original P. oaksoil taxon isolate recovered from France, but it differed in its optimal growth temperature and mitochondrial cox region, therefore, distinguishing it as a different haplotype from its closest relatives. We did not analyze the cox spacer region during this study; thus, it is unknown if these isolates are one of the neighboring Clade 6 relatives, including P. bilorbang Aghighi and Burgess, which was originally described from Western Australia (Aghighi et al. 2012). Phytophthora ramorum Species-isolation method/confirmations. We isolated P. ramorum once from Vaccinium arboreum foliage on 16 April 2014, and 3 times from filtered water on 17 December 2014 when the water temperature was 6.7 °C (Table 1), which lies within its vegetative cardinal growth temperatures of 2–30 °C (Werres et al. 2001). The leaf spots on the foliage were small (2–3 mm) and surrounded by a halo of brown, with necrotic tissue in the center. Culture morphology. Sporangia were borne singularly but mostly sympodially, semi-papillate, caducous with a short pedicel, ellipsoid, and embedded in the agar and on top of the surface of PAR-V8 medium, forming a botryose-like clump (Fig. 3). Large, globose, thin-walled chlamydospores formed intercalary and terminally, and in abundance. Hyphae exhibited a unique, knobby, coralloid growth form and grew relatively slowly in PAR-V8 medium incubated at 20 °C, which made zoospore-derived colonies from filtered water easy to distinguish under a dissection microscope. Molecular data. The consensus ITS sequence from strains T3 Vaccinium (KY696631), T11 A.32 (KY696633), T11 A.34 (KY696632), and T11 B.5 (KY696634) from this investigation showed 100% coverage and 100% similarity with HM004221, which is identified as P. ramorum. These sequences occur in a strongly supported clade (100% PP; Fig. 2). Comments. Phytophthora ramorum is heterothallic and placed in Clade 8, with its closest relative P. lateralis Tucker and Milbrath, which is distinguished from P. ramorum in the ITS-1 and ITS-2 regions by 3 and 8 nucleotides, respectively (Werres et al. 2001). We identified isolate numbers T11 (I.L. A.6, A.3, A.10, A.21, A.18) and T14 (I.L. A.6, B.12, A.12) morphologically as P. ramorum by the ellipsoid, semi-papillate, deciduous sporangia borne in sympodial clusters on the Southeastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 194 surface of the agar, although the cultures also had P. cinnamomi present. Therefore, many of these cultures within the same plates were sequenced and identified as P. cinnamomi, but we later pipetted P. ramorum sporangia from the surface of the agar and attempted a new culture. There are 7 P. ramorum-regulated or -associated host genera within the study area, including Acer (maple), Fagus (beech), Lonicera (honeysuckle), Quercus (oaks), Rhododendron (azalea), Salix (willows), and Vaccinium (blueberries) (USDA APHIS 2013). Phytophthora riparia Species-isolation method/confirmations. We isolated this species twice: once from a BOB leaf disk and once from filtered water, both collected from the drainage ditch on 31 October 2014 when the water temperature was 6.8 °C (Table 1). Hansen et al. (2012) described the organism’s optimum growth on carrot agar media at 25 °C, with slow growth at 35 °C, and no growth at 40 °C. Morphological data. No sexual gametes were produced in culture. Sporangia were ovoid to obpyriform, ellipsoid, non-caducous, and non-papillate with a slight apical thickening. Sporangia were mostly borne terminally and singularly, some from an internally proliferated sporangiophore. We measured 2 exit-pore openings at 12 μm and 14 μm. No chlamydospores were produced in culture. We observed coiled hyphae and elongated hyphal swellings in PAR-V8 media. Molecular data. The consensus ITS sequence from strains T8 A.3 (KY696635) and T8 BOB B.1 (KY696636) from this investigation showed 100% coverage and 99% similarity to HM004225, which has been identified as a strain of P. riparia. These sequences cluster within a clade with 100% PP (Fig. 2). Comments. Phytophthora riparia is a heterothallic species in Clade 6 with its closest relative P. taxon salix soil, differentiated in the ITS region by 6 nucleotide positions, plus 1 indel position (Hansen et al. 2012). This relatively benign species of Phytophthora was recently described from forest streams in Oregon, California, and Alaska, and has not been associated with any disease. Stamler et al. (2016) identified it as one of the most prevalent Phytophthora species recovered from major waterways across the southwest. To our knowledge, our identification of P. riparia from Mississippi during this investigation is the first report of this taxon from the eastern US. Phytophthora rosacearum Species-isolation method/confirmations. We isolated this species once from soil in section 2 (Fig. 1), located in the drainage ditch on 3 June 2014 when the soil temperature was 24.6 °C (Table 1). Hansen et al. (2009) described its optimum growth on carrot agar at 30 °C, and a maximum growth temperature at 36 °C. Morphological data. Phytophthora rosacearum did not produce any reproductive structures and had no distinguishable hyphal characteristics or other unique characteristics. Therefore, ITS sequence data was the only method available to confirm the identification. Molecular data. The consensus ITS sequence from strain T6 Soil 2.2 (KY696637) from this investigation showed 100% coverage and 99% similarity to Southeastern Naturalist 195 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 P. aff. rosacearum (KU211497). In addition to KU211497, several other accessions had Phytophthora aff. rosacearum, indicating that it resembled P. rosacearum. These 2 sequences are supported in a unique clade with 100% PP (Fig. 2). Comments. Phytophthora rosacearum is a Clade 6 species segregated from the P. megasperma Drechsler complex, and originally described as a pathogen of rosaceous fruit trees. Two plant genera, Rubus sp. (blackberry) and Crataegus sp. (hawthorn), are in the Rosaceae and were located within the study area. We sampled both plant species with null results. Phytophthora syringae Species-isolation method/confirmations. We isolated this species from filtered drainage-ditch water 4 times on 27 January 2015, when the water temperature was 10.8 °C (Table 1). Doster and Bostock (1988) stated that the optimum vegetative growth temperature for this taxon is 21 °C; oospore production was halted at temperatures higher than 12 °C. Morphological data. Even though the species is sexually fertile, we documented no sexual gametes. This finding matches the results of Doster and Bostock (1988), who suggested the use of vegetable oils to stimulate the production of oospores in culture. Sporangia were semi-papillate, unbranched and borne terminally, ranging from ovoid, obpyriform, globose, to ellipsoid, and non-caducous. We observed prolific catenulate hyphal swellings in PAR-V8 media. Molecular data. The consensus ITS sequence from strains T13 A.5 (KY696639), T13 A.7 (KY696638), T13 A.25 (KY696640), and T13 B.8 (KY696641) from this investigation showed 100% coverage and 99% similarity to P. syringae (HM004229). These sequences cluster in a strongly supported clade with 100% PP (Fig. 2). Comments. Phytophthora syringae is a homothallic species in Clade 8; its closest relative is P. austrocedrae Gresl. and Hansen (Martin et al. 2014). This pathogen has been reported on nursery plants, Malus pumila Miller (Paradise Apple), and Pyrus sp. (pear) trees (Laywisadkul et al. 2010). Discussion We identified a total of 11 species of Phytophthora through use of morphological and molecular ITS-sequence data, with bayesian phylogenetic analyses that indicated monophyletic species (Fig. 2). These results differed from those of Copes et al. (2015), who recovered 8 species and 1 taxon from irrigation ponds in Alabama and Mississippi; P. mississippiae was the only overlapping species. Species richness in this urban riparian-forest ecosystem may be attributed to anthropogenic involvement of surrounding landscapes, native and exotic plant host-species within the riparian-forest habitat, and regional climate. High summer temperatures may have restricted their occurrences; between 1950 and 2017, the June–August mean temperature in Jackson, MS, was 27.2 °C, and monthly mean precipitation was 38.1cm (NOAA 2017). The highest inoculum loads released into the environment may not only be dependent on seasonal temperatures, but also the amount of precipitation, UVSoutheastern Naturalist D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 196 radiation exposure, and relative humidity (Englander et al. 2006, Tooley et al. 2009). The sampling trip resulting in the most isolations (17 December 2014) coincided with the highest recorded water level—water levels in the drainage ditch were 3 m higher than average and inundated every plant in the study area. We isolated P. ramorum 3 times from filtered water on this date, demonstrating that the climate was conducive for zoosporogenesis. This speculation is supported by Davidson et al. (2011), who found that increased transmission rates of P. ramorum in multiple forest types were associated with high-rainfall years in California. The increase of inoculum in waterways during heavy rain events suggests (1) the possible inundation of infected vegetation or the aquatic deposition of residual inoculum from runoff, or (2) the increased production of sporangia and germination of chlamydospores from infected vegetation, which naturally disseminates into streams (Crone et al. 2013). In addition to climate, baiting and sampling methods may be selective for species depending on their mode of dissemination into the environment and their ecological niche. We confirmed all 11 taxa across all dates from at least 1 of the baiting methods used, but frequencies varied (Table 3). It is not surprising that species in Clade 6, which are ubiquitous in aquatic and forested environments and can be saprophytic (Kroon et. al 2012), were most commonly recovered from baited water. This finding is supported by the fact that P. mississippiae was the most prevalent species isolated from filtered stream water and the BOB intact-leaf samples, at frequencies of 14.3% and 12.5%, respectively (Table 2). We isolated P. cryptogea, P. riparia, and P. syringae numerous times, but from only 1 or 2 sampling trips, suggesting that their presence in the environment is irregular. We recovered P. chlamydospora from soil, water, and vegetation from 7 sampling trips over a period of 6 months, indicating a perpetual presence in the local environment, possibly because of the organism’s ability to tolerate high temperatures, its presence in aquatic habitats, and its broad distribution worldwide (Hansen et. al 2015). We recovered some species when temperatures were far from ideal, which provoked us to speculate on how and where these organisms complete their life cycle. We isolated P. riparia from water at 6.8 °C even though its optimal temperature in vitro is 25 °C (Hansen et al. 2012). We most often isolated P. mississippiae at a water temperature of 6.8 °C, but Yang et al. (2013) suggested an optimum in vitro temperature of 30 °C for this species. Copes et al. (2015) discussed these temperature associations after they recovered the high-temperature–tolerant P. irrigata during the month of December. Detection of these taxa under suboptimal growing conditions could stem from their morphogenesis in plant tissue. Survival or overwintering by P. cinnamomi was reported to occur biotrophically from oospores, thick-walled chlamydospores, and mass hyphae in stromata in herbaceous perennials (Crone et al. 2013, McCarren et al. 2005). Multihyphal survival structures of P. ramorum in vegetation, including dense clusters of chlamydospores and sporangia, could later infect hosts during active periods (Moralejo et al. 2006). It was further shown that these structures would enable survival across high summer temperatures. Tooley et al. (2009) reported that Southeastern Naturalist 197 D.S. Bily, S.V. Diehl, M. Cook, L.E. Wallace, L.L. Sims, C. Watson, and R.E. Baird 2018 Vol. 17, No. 1 P. ramorum infected rhododendron leaves at a notably higher range of temperatures than other Phytophthora spp. Thus, persistence of these reproductive structures within plant tissue would enable sampling for molecular detection of Phytophthora spp. during unfavorable environmental conditions. The natural infection of riparian host plants by P. cinnamomi and P. ramorum, resulting in a biotrophic disease scenario, could sustain a reservoir of inoculum that is deposited into streams during rain events without causing obvious plant mortality. Our sampling area contained numerous plant host-species and associated plant host-species for the 2 species of Phytophthora. Phytophthora cinnamomi was isolated from a Swamp Chestnut Oak petiole and a Salix nigra Marsh. (Black Willow) seedling that was located within the drainage ditch. We isolated P. ramorum once from a surfaced-sterilized Farklebery leaf that was submerged during periods of flooding. This finding was previously supported when P. ramorum was isolated from a riparian willow in 2010 at this same site (S. Jeffers, pers. comm.). Even though we isolated many Phytophthora spp. from host plants, vegetative damage never appeared to be severe during this investigation or prior samplings (S. Jeffers, pers. comm.). Phytophthora ramorum has prevailed at this site for at least 10 years, but there has never been any evidence of substantial vegetative dieback or tree decline. This apparent lack of pathogenic effect may be attributed to (1) the relatively small climatic window in Mississippi that is conducive for the pathogen to sporulate, (2) the precipitation frequency, and (3) the correlation of plant dormancy with the months of the year that are favorable for disease progression. The role of Phytophthora spp. in this urban ecosystem appears to be non-threating and similar to other species in the genus known to occur in this riparian habitat. In conclusion, our study demonstrates that a fairly broad range of Phytophthora species are present in this urban riparian-forest area of south-central Mississippi. It is critical to use the 5 detection methods developed by USDA APHIS across multiple sampling dates to ensure full sampling coverage of taxa present at any site being studied. Multiple dates of sampling for species of Phytophthora are of particular importance at sites where temperatures go above 29 °C or below 4.5 °C, which can limit detection. Furthermore, the use of molecular-sequence information supplemented with morphological data, when available, was key to ensuring accuracy in taxa verifications. Acknowledgments Special thanks to Dr. Steven Jeffers and Dr. Jaesoon Hwang at Clemson University for their technical training and expertise. Additional thanks to Dr. Warren Copes at the USDA Agricultural Research Service, Poplarville Mississippi for technical assistance and isolate of P. mississippiae. Devin S. 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