In general, fully senesced leaves have as much as 75% reduced protein content compared with green leaves, primarily from the dismantling of chloroplasts, and though yellow, senescing leaves still have live cells with active mitochondria, leaves that have turned brown as a result of drying no longer contain biologically active cells . Therefore, green, senescing and fully senesced bay leaves are substrates that likely vary in their suitability for colonization by P. ramorum and stream-resident clade 6 Phytophthora species, taxa that commonly occur at high inoculum levels in northern California coastal forest streams. We have shown that there is a difference in trophic specialization between the saprotrophically competent, clade 6 Phytophthora species, such as P. gonapodyides, and P. ramorum, an aggressive pathogen on many plant species. In that study, green California bay leaves were rapidly colonized by P. ramorum in streams but were succeeded nearly completely within three weeks by clade 6 Phytophthora species. It remains uncertain, however, whether P. ramorum was displaced by more competent saprotrophs or receded from an inability to persist in tissues that it had colonized as they progressively decomposed. Additionally, as most leaf litter consists of senesced leaves, it is important to know how these differently adapted taxa can compete for and persist on biologically inactive leaf tissue. Finally, though stream resident Phytophthora species are assumed to contribute to leaf decay given their regular recovery from streams and frequent association with decomposing vegetation, experimental evidence for the kind and extent of this contribution is lacking.

Moreover, it is unknown how the introduction of an exotic and plant pathogenic species, like P. ramorum, strawberrry gutter system into a stream ecosystem might affect the decomposition of leaf litter by other organisms, such as resident Phytophthora species. Therefore we undertook a laboratory study to determine: How well P. ramorum and P. gonapodyides could use senesced leaves as a substrate in comparison to green, live leaves, whether colonization by and persistence of P. ramorum on leaves was affected by competition with P. gonapodyides, and how much each of these Phytophthora species contribute to the decay of each leaf type. To test the capacity of P. ramorum and P. gonapodyides to colonize green and senesced bay leaves, we conducted controlled environment experiments exposing leaves to an inoculum of each species alone and in combination in microcosms designed to simulate an aquatic environment. The experiment consisted of a randomized complete block design with treatments representing a complete factorial of bay leaf type , stream water addition , and Phytophthora inoculation. These 16 treatment combinations were replicated in five blocks arranged in three growth chambers. The experimental unit was a mesh packet of five leaves which were sampled at intervals over 16 weeks from microcosms. One treatment packet per sampling served for decomposition as percent biomass loss and another for colonization based on isolations on a selective medium. We repeated the experiment once, with leaf types maintained in the same microcosm in the first and in separate microcosms in the second experiment.

We conducted a separate experiment with yellow, senescing leaves collected while still attached to trees and with the cuticle intact, with P. ramorum-only and combined P. ramorum/ P. gonapodyides treatments as well as non-inoculated controls, in a completely randomized design with four reps in a single growth chamber. We collected leaves from two sites where our previous field experiments were conducted. One was a canyon through which Graham creek runs at Jack London State Park which consists of redwood forest with California bay as a dominant riparian tree, along with redwood , Douglas fir , tanoak , bigleaf maple , and less frequently, madrone. The second included canyons around Copeland Creek at Sonoma State University’s Fairfield Osborn Preserve which is characterized by mixed evergreen forest with a prevalence of California bay, white alder , big leaf maple, and occasionally, tanoak, madrone and coast live oak. At each site, we collected green, symptom-free bay leaves with a mature cuticle from trees and brown, recently shed bay leaves from beneath trees in the manner of Wood et al.. Brown leaves were collected from both sites in September 2014, allowed to air dry in the laboratory, and then stored in sealed plastic bags at room temperature until used in experiments. Green leaves were collected on 12 December 2014 from the redwood forest site and on 5 August 2015 from the mixed evergreen forest site. Yellow leaves were collected directly from trees at the mixed evergreen forest site on 7 September 2015. Green leaves were stored at 4 ◦C for up to three weeks prior to use in experiments and yellow leaves were likewise stored but deployed in experiments within one week of collection. We collected leaves primarily from riparian areas around the described creeks, though, at the mixed evergreen forest site,grow strawberry in containers we had to seek symptomless leaves to some extent from plateaus above the canyons.

Leaf treatments were in the same microcosm for the first experiment using leaves from the redwood forest and separate in the second experiment with leaves from the mixed evergreen forest. Yellow leaves were collected from the mixed evergreen forest site only. We tested a sub-sample of 50 of each leaf type for both sites—through isolations attempted on a selective medium as described below—to verify that there were no pre-existing Phytophthora infections. Brown leaves were soaked in sterile deionized water at 4 ◦C for two days prior to these test isolations. Leaf packets were prepared for each leaf type by packing five leaves into a flat envelope of 1 mm plastic mesh approximately 20 × 20 cm so that the leaf surfaces were in minimal contact with one another and each packet was sealed by folding over the open lip and securing it with two common metal staples. We assembled microcosms simulating an aquatic decomposition environment similar to the approach described by Medeiros et al.. White plastic buckets were used in the first and yellow leaf experiments and opaque plastic containers in the second experiment. Each container was aerated through a tube terminating in an aeration stone fed by an air pump that was turned on for 30 minutes twice daily using an electric timer. Aeration intensity was moderated with the addition of adjustable valves inserted in the tubing. A dilute nutrient solution was used as the base for the water mixtures in microcosms in order to avoid osmotic stress on spores. This was achieved by adding Hoagland’s #2 salts to autoclaved Millipore®filtered water for a final concentration of 0.01× the standard concentration. To test for any effect of natural stream microbiota on Phytophthora colonization or leaf decomposition, we included an addition of auto-claved or non-sterilized stream water as a treatment factor. The final composition of water in microcosms consisted of 4 L nutrient solution and 2 L stream water in the first experiment, and 4 L nutrient solution and 1 L stream water in the second. We collected water from streams in a bucket, pouring it through several layers of cotton mesh into 4 L plastic bladders that we consolidated into larger plastic containers or used directly to transport water out of the field. Once brought to the laboratory, stream water was stored in plastic containers in a growth chamber at 12 ◦C and 12 h photoperiod for 20 and 23 days prior to deployment in the first and second experiments, respectively. After storing the water for seven days, we submerged symptomless California bay leaves collected at each site as baits in each container for two days to confirm that Phytophthora zoospores were not present.

We tested baits for infection using the isolation technique described below. No Phytophthora infections were detected from baits at this point. In the experiment with yellow leaves, we used only 4 L of a nutrient solution without stream water addition. We measured stream pH, electrical conductivity and temperature on site at the time of stream water collection and subsequently in each microcosm throughout the experiments with a portable sensor. Stream pH, EC, and water temperature were 8.55, 208 µS/cm, and 13.5 ◦C, respectively, for the redwood forest stream on the 9 December 2014 collection date, and 8.08, 363 µS/cm, and 17.7 ◦C, respectively, for the mixed evergreen forest stream on the 5 Aug 2015 collection date. To approximate natural stream pH in microcosms, we amended the mix of dilute nutrient solution and stream water in each microcosm with potassium carbonate buffer at approximately 10 mg/L and adjusted it with KOH and HCl for a target of pH 8.3. The average pH measured in microcosms periodically over the course of experiments was 7.99 , 8.33 and 8.24 in the first, second and yellow leaf experiments, respectively. The average EC was 208 , 141 , and 98 in the first, second, and yellow leaf experiments, respectivelyTo determine the rate of decomposition measured as leaf biomass loss, we weighed leaves to the hundredth decimal of a gram with an analytical balance prior to packing and we labeled the packets with aluminum tree tags secured with a plastic tie for future identification. We estimated the original dry mass of both leaf types from the average dry weight of a sub-sample of 50 fresh or air dried leaves. The average percent dry weight for green and brown leaves, respectively, was 40.9 and 94.3 for the first experiment and 53.5 and 92.6 for the second experiment. The average percent dry weight for yellow leaves was 55.0. The average estimated weight in grams for five green leaves was 0.84 and 1.13 , and that for five brown leaves, 0.91 and 0.85 for the first and second experiments, respectively. For five yellow leaves, the estimated average weight in grams was 0.98. At each sampling, leaves were retrieved from tagged leaf packets, rinsed gently with deionized tap water to remove adhering debris, oven-dried in a paper envelope or an open aluminum foil envelope at 55–60 ◦C for 48 h, and weighed as described above. The fraction of original biomass was calculated for all leaves in a packet by dividing the weight at the time of sampling by the estimated original dry biomass. To determine the level of Phytophthora colonization of leaves, at each sampling we collected a packet for each leaf type from each container to evaluate by culturing on Phytophthora-selective PARP-H medium 50 ppm and hymexazol 25 ppm. Upon retrieval, leaves were submerged and gently rubbed free of biofilm in 1% household bleach solution , surface sterilized in fresh bleach solution for three to seven minutes, rinsed with deionized tap water, and then laid out on paper towels and the excess water allowed to evaporate. Finally, leaves were wrapped in a paper towel and stored at 4 ◦C until isolations by culturing could be performed. Isolations were attempted from all leaves belonging to treatment using a ‘mosaic’ sampling approach whereby the leaf discs are removed from the petiole, midrib and flanking lobes of the leaf at approximately 1 cm distance from one another in order to collect a representative sample from the entire leaf. For experiments with green and brown leaves, isolations were initiated immediately after collection, with most samples processed within 29 days. All isolations were completed by 46 days after collection. Storage period did not alter results when included as a covariate in models for these experiments and was excluded from the final analyses. Isolations from leaves of the yellow leaf experiment were completed within nine days after collection, and all isolations from a single collection week were completed in one day. The presence of P. ramorum and P. gonapodyides was determined by microscopic examination of isolate morphologies directly from the isolation plates after four to five days and checked again periodically for three weeks. To test for active sporulation from colonized leaves in the microcosms, periodically a California bay leaf disc was floated as bait—either naked or in a roughly 35 mm2 mesh envelope—on the surface of the water in each microcosm for three to seven days, after which it was surface sterilized and isolations attempted from it on selective PARP-H medium. We conducted these tests of sporulation four times during the first and yellow leaf experiments, and three times during the second experiment. Additionally, we tested for sporulation periodically for up to eight weeks after all leaves had been removed from microcosms to determine if Phytophthora spores could persist in the absence of a substrate.