Cultural Controls for Suppressing Phytophthora Cinnamomi Root Rot of Blueberry

AN ABSTRACT OF THE DISSERTATION OF

John R. Yeo for the degree of Doctor of Philosophy in Soil Science presented on September 5, 2014. Title: Cultural Controls for Suppressing Phytophthora cinnamomi Root Rot of Blueberry

Abstract approved:

Dan M. Sullivan

Phytophthora cinnamomi is a soilborne pathogen that causes root rot disease of highbush blueberry (Vaccinium corymbosum L.). When new installations of susceptible blueberry cultivars are infected with P. cinnamomi, plants often fail to grow significant new tissue, greatly reducing yields over the life of the planting. Chemical fungicides are available for disease suppression including mefenoxam and phosphonate compounds. However these tools are not available for organically certified growers.

Initially, this research focused on non-chemical control strategies for blueberry root rot disease, including cultivar selection, soil amendments (gypsum and a variety of organic materials) via a series of trials in the greenhouse. Based on the findings of the greenhouse experiments, a field trial was conducted to evaluate a combination of cultural factors in a two year field experiment. Pre-plant gypsum incorporation into soil was evaluated as a control strategy in the field experiment in combination with other cultural practices that were hypothesized to affect disease including mulch type (geotextile weedmat or sawdust), and drip irrigation line placement.

Eighteen highbush blueberry cultivars and advanced breeding selections were evaluated for susceptibility or resistance in three greenhouse experiments. Cultivars varied widely in susceptibility, with ‘Duke,’ ‘Draper,’ ‘Bluetta,’ ‘Blue Ribbon,’ ‘Cargo,’ ‘Last Call,’

‘Top Shelf,’ and ‘Ventura’ exhibiting high levels of susceptibility to the disease. More

resistant cultivars included ‘Legacy,’ ‘Liberty,’ ‘Aurora,’ ‘Overtime,’ ‘Reka,’ and

‘Clockwork’. By selecting cultivars with superior resistance, growers may avoid yield

losses associated with the disease, enhancing production profitability. Although choosing

a disease resistant cultivar is likely the most effective control strategy for blueberry, other

cultural disease control practices are needed when susceptible cultivars are grown to fill

market demands. Fungicides can control disease in conventional plantings, but cannot be

applied within certified organic production systems.

Gypsum and organic amendments sometimes provide suppression of Phytophthora root

rot in crops other than blueberry. Three greenhouse trials were conducted to evaluate

organic soil amendments (peat, sawdust, dairy solids compost, yard debris compost, and

composted municipal biosolids blended with douglas-fir bark) incorporated at 20% v/v

into soil, and gypsum (CaSO4) incorporated at 5% (v/v) for disease suppression in factorial combination. Trials were conducted with a disease susceptible cultivar

('Draper'), and soil moisture was maintained near saturation, favoring disease development. Organic amendments did not suppress disease in any of the experiments.

Gypsum was effective in disease suppression in one of the three experiments.

Gypsum provides a source of soluble calcium, and previous reported studies have

demonstrated a mechanism for calcium-mediated suppression of P. cinnamomi. An

additional greenhouse study was conducted to determine the relationship between

gypsum application rate, soluble calcium in soil solution, and disease suppression.

Soluble Ca in soil solution reached a plateau concentration of ~454 mg Ca L-1 soil

solution at gypsum application rates equal to or above 16 meq gypsum 100g-1 soil.

Higher gypsum application rates did not increase soil solution Ca, or provide additional

disease suppression.

Rates of gypsum required for disease suppression increase soil electrical conductivity

(EC) beyond current recommended salinity guidelines for blueberries (> 2.0 mS/cm). The effects of salinity on plant growth were evaluated in a six month greenhouse experiment, using gypsum (CaSO4) or potassium sulfate (K2SO4) salts to increase EC. Treatment EC levels ranged from 0.3 to 2.6 mS/cm, in ten increments. The maximum EC value chosen for this experiment approximates the maximum value for gypsum solubility in soil solution. Aboveground plant biomass declined more rapidly when EC was supplied by potassium sulfate than it did with gypsum. Root biomass declined when EC was supplied by potassium sulfate, but it was not affected by gypsum rate. Leaf cation concentrations responded strongly to increasing rates of potassium sulfate application. Leaf K increased dramatically, accompanied by declines in leaf Ca and Mg concentration. In contrast, leaf cation concentrations were much more stable when EC was adjusted by gypsum addition.

Under the conditions of this greenhouse experiment, the increase in EC accompanying gypsum application had only minor effects on plant growth and nutrient uptake, suggesting that gypsum application is a viable option for trial in the field.

A two year field trial was conducted to evaluate cultural practices for efficacy in suppressing P. cinnamomi root rot disease. Pre-plant gypsum incorporation into soil was evaluated as a control strategy in combination with other cultural practices that were hypothesized to affect disease: mulch type (geotextile weed mat vs. sawdust), and drip irrigation line placement. Disease suppression over two growing seasons was evaluated with the highly susceptible cultivar ‘Draper’ grown on a site with clay loam soil. P. cinnamomi inoculum was mixed into soil before planting. Experimental design was a

2x2x2 factorial with 2 mulch types (geotextile weed mat or douglas-fir sawdust), 2 drip line placements (narrow or wide placement relative to plant row), and 2 gypsum rates (0 and + gypsum). Gypsum was incorporated into planting beds in a 30-cm band at an

application rate of 22,420 kg ha-1 in-band equivalent to 2,242 kg ha-1 on a whole-field basis. Drip irrigation treatments consisted of two drip lines placed either adjacent to the plant crown or 20-cm on either side of the crown (wide placement). A fungicide treatment was included to provide an assessment of the efficacy of cultural disease control vs. a chemical alternative. The fungicide treatment was mulched with sawdust and irrigated with drip lines adjacent to plants, and did not receive gypsum application.

Mulch type had no significant effect on plant biomass after two years, but plants grown under sawdust had slightly higher biomass and less root infection. Soluble Ca, as measured by mini-lysimeters placed in the rootzone, was increased by gypsum application, especially with wide placement of drip irrigation lines. The disease suppressive effect of gypsum amendment depended on irrigation line placement. Soluble

Ca movement in soil was associated with water movement away from drip emitters.

When drip irrigation lines were placed adjacent to the plant crown, soluble Ca was moved away from the plant with the wetting front. With wide drip line placement , soluble Ca was moved toward the roots. Plants grown with wide drip line placement and gypsum addition had the lowest root infection incidence and highest plant biomass, likely as the result of more soluble Ca in the rootzone. Despite significant increases in plant biomass with gypsum and widely-placed irrigation lines, plants treated with conventional fungicide had approximately twice as much biomass after two growing seasons.

An integrated control program is required for cultural suppression of blueberry root rot disease. Organic production using highly susceptible cultivars in the presence of P. cinnamomi is difficult, and may not produce equivalent yields to conventional production. Improved plant performance in the presence of P. cinnamomi was observed in these trials using cultivar resistance, gypsum, and widely-spaced drip irrigation lines.

Other cultural practices, such as careful irrigation scheduling, and appropriate rate and timing of N fertilizer application are also important for disease suppression. In the future,

greater P. cinnamomi disease suppression should be possible by using the disease suppressive cultural practices identified in this research in combination with a disease-

resistant cultivar.

Cultural Controls for Suppressing Phytophthora cinnamomi Root Rot of Blueberry

John R. Yeo

A DISSERTATION

submitted to

Oregon State University

in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

Presented September 5, 2014 Commencement June 2015

Doctor of Philosophy dissertation of John R. Yeo presented on September 5, 2014

APPROVED:

Major Professor, representing Soil Science

Head of the Department of Crop and Soil Science

Dean of the Graduate School

I understand that my dissertation will become part of the permanent collection of Oregon State University libraries. My signature below authorizes release of my dissertation to any reader upon request.

John R. Yeo, Author

CONTRIBUTION OF AUTHORS Dr. Jerry Weiland assisted with the experimental design, statistical interpretation, and writing of chapter two, three, and four. Dr. David R. Bryla assisted with the experimental design, statistical interpretation, and writing of chapter two, three, four, and five. Dr. Dan M. Sullivan assisted with the experimental design, statistical interpretation, and writing of chapter two, three, four, and five, and also contributed to the writing of the general abstract.

TABLE OF CONTENTS

Page 1. General Introduction……………………………………………………….…. 1 History of the global blueberry supply chain ………………………... 2 Origin, global distribution, and hosts of Phytophthora cinnamomi………………………………………………………...... 2 Oregon blueberry production practices and status of Phytophthora cinnamomi in the Oregon blueberry industry………………...…….... 3 Disease control……………………………………………………….. 5 Relationship between soil moisture, matric potential, and Phytophthora spp. root rot disease…………………………………… 7 Influence of water status on P. cinnamomi root rot of blueberry…….. 9 Efficacy of gypsum application for Phytophthora root rot control in non-blueberry crops…………………………………………….…….. 9 Response of Phytophthora cinnamomi to temperature……………...... 10 Works Cited…………………………………………………………. 11

2. Differences in highbush blueberry (Vaccinium corymbosum) cultivar susceptibility to Phytophthora cinnamomi Abstract ……………………………………………………………….… 21 Introduction…………………………….…………………….………..… 21 Methods………………………………….…………………....……….… 22 Results………………………….……………………………..…………. 27 Discussion…………………………………………………….…………. 31 Works cited…………………………………………………….………... 29 Figures and tables…………………………………………….…………. 31

3. Evaluation of gypsum and organic matter soil amendments for suppressing Phytophthora cinnamomi root rot disease of blueberry Abstract ………………………………….……………………………… 38 Introduction………………………………….…...... …………….……… 39

Page Methods……………………….………….……………………….……...... 40 Results …………………………………………………………….………...46 Discussion………………………………………………………….………..49 Works cited……………………………………………………….……...... 51 Tables and Figures………………………………………………………….54

4. Influence of gypsum, mulch, and drip irrigation line placement on Phytophthora cinnamomi root rot disease during field establishment of highbush blueberry Abstract……………………………………………………………………...69 Introduction………………………………………………………………....70 Methods…………………………………………………………………...... 71 Results……………….. ……………………….…………………………....76 Discussion……………………………………………………………..…….78 Works cited………………………………………………………………… 81 Tables and figures……………………………………………………..…… 83

5. Cation-specific salinity effects from calcium and potassium on highbush blueberry Abstract…………………………………………………………………….. 98 Introduction………………………………………………………………… 99 Methods……………………………………………………………………..99 Results……………………………………………………………….……...101 Discussion ……………………….……………………………….………...103 Works cited………………………………………………………….……...105 Tables and Figures…………………………………………………………..108

6. General Conclusion……………………………………………………………..114

Literature Cited…………………………………………………………………118

LIST OF FIGURES Figure Page 2.1 Correlation between root infection incidence and shoot mass of 10 blueberry cultivars infected with P. cinnamomi, relative to shoot mass of non-infected plants of the same cultivar…………………………………... 35 2.2 Correlation between root infection incidence and root mass of 10 blueberry cultivars infected with P. cinnamomi relative to root mass of non-infected plants of the same cultivar…………………………………... 36 3.1 Effect of increasing gypsum amendment rate on soluble Ca2+ in soil solution sampled with Rhizon tube mini-lysimeters……………………… 64 3.2. Effect of gypsum rate on P. cinnamomi root infection incidence………… 65

3.3. Effect of gypsum rate on shoot biomass of plants infected with P. cinnamomi………………………………………………………………… 66 3.4. Effect of gypsum rate on root biomass of plants infected with P. cinnamomi. ……………………………………………………………….. 67 4.1. Differences in root infection incidence between geotextile weed mat and sawdust mulch at five sampling dates over two growing seasons………… 90 4.2. Influence of drip irrigation line spacing on P. cinnamomi root infection incidence over two growing seasons……………………………………… 91 4.3. Influence of gypsum amendment on P. cinnamomi root infection incidence over two growing seasons……………………………………… 92 4.4. Root infection incidence and plant biomass after two growing seasons of cultural treatments with the consistently highest and lowest root infection incidence, and the fungicide control………………………………………. 93 4.5. Diurnal soil temperature fluctuation at 15-cm beneath either sawdust or woven black geotextile weed mat…………………………………………. 94 4.6. Mean main effects of (A) drip irrigation line placement, (B) mulch type, and (C) gypsum incorporation on volumetric soil moisture in the center of raised beds1-hr after a 15-min irrigation event on 12-Jul. and 8-Aug. 2013……………………………………………………………………….. 95 4.7. Calcium levels in soil solution at the plant crown 5 and 16 months after planting……………………………………………………………………. 96

5.1. Influence of increasing EC with either gypsum or K2SO4 on vegetative biomass of ‘Draper’ highbush blueberry………………………………….. 109 5.2 Ca2+ leaf concentration, soil solution concentration, and exchangeable 2+ Ca with increasing salinity added with either gypsum or K2SO4……….. 110

LIST OF FIGURES (Continued) Figure Page 5.3 K+ leaf concentration, soil solution concentration, and exchangeable K+ with increasing salinity added with either gypsum or K2SO4……………... 111 5.4. Mg2+ leaf concentration, soil solution concentration, and exchangeable 2+ Mg with increasing salinity added with either gypsum or K2SO4………. 112 2- 5.5. S leaf concentration, soil solution SO4 concentration, and Mehlich III 2- extractable S (as SO4 ) with increasing salinity added with either gypsum or K2SO4…………………………………………………………………… 113

LIST OF TABLES Table Page 2.1 Shoot mass of P. cinnamomi infected blueberry plants, relative to non- infected shoot growth of the same cultivar………………………………... 31 2.2. Root mass of P. cinnamomi infected blueberry plants, relative to root mass of non-infected plants of the same cultivar. ………………………... 32 2.3. Incidence of 1-cm root sections infected with P. cinnamomi…………….. 33 2.4. Root infection incidence, and relative shoot and root mass compared to non-infected plants of the same cultivar. Trial 3………………………….. 34 3.1. Main and interaction effects for aerial and root biomass, and all treatment mean separation. Spring-Summer 2011 greenhouse experiment…………. 54 3.2. Root infection incidence and P. cinnamomi propagule density in soil after 14 wks. Spring-Summer 2011 greenhouse experiment…………………… 55 3.3. Soil chemical properties of soil-organic matter-gypsum blends after 14- wk. Trial 1………………………………………………………………… 56 3.4. Total nutrient uptake of ‘Draper’ highbush blueberry grown in different gypsum and organic matter amendments, either infected or non-infected with P. cinnamomi root rot. Trial 1……………………………………….. 57 3.5. Main and interaction effects for aerial and root biomass, and all treatment mean separation. Fall-Winter 2011 greenhouse experiment……………… 58 3.6. Root infection incidence and P. cinnamomi propagule density in soil after 14 wks. Fall-Winter 2011 greenhouse experiment………………………... 59 3.7. Main and interaction effects for aerial and root biomass, and all treatment mean separation. Spring 2013 gypsum and organic matter greenhouse experiment…………………………………………………………………. 60 3.8. P. cinnamomi root infection incidence after 9 wks. Spring 2013 greenhouse experiment……………………………………………………. 61 3.9. Chemical analysis of saturated deionized water extract analysis of organic residues used in the spring and fall 2011 experiments evaluating organic amendments and gypsum for blueberry root rot disease suppression……………. 62 3.10. Total nutrient analysis of organic amendments used in the spring and fall 2011 experiments evaluating organic amendments and gypsum for blueberry root rot disease suppression. Nutrient concentrations are reported on a dry-weight basis…………………………………………….. 63 4.1. Biomass of ‘Draper’ blueberry grown in Phytophthora cinnamomi infested soil after one growing season…………………………………….. 83

LIST OF TABLES (Continued) Table Page 4.2. 2013 Main and interaction effects of mulch, drip irrigation spacing, and gypsum incorporation on plant biomass after two growing seasons……… 84 4.3. Root infection incidence at five sampling dates over two growing seasons…………………………………………………………………….. 85 4.4. Leaf tissue nutrient concentration Aug 2, 2012, 10-wks after planting (Year 1)……………………………………………………………………. 86 4.5. Leaf tissue nutrient concentration August 1, 2013, 14-months after planting (Year 2)………………………………………………………...... 87 4.6. Soil heat unit accumulation, and P. cinnamomi hyphal growth on sealed and buried PDA petri plates, beneath sawdust and black woven plastic geotextile weed mat mulch.……………………………………………….. 88 4.7 Soil pH and Mehlich III extractable cations from plots treated with all combinations of gypsum, drip irrigation line placement, and mulch type... 89 5.1. Relationships between electrical conductivity (EC) and plant biomass, nutrient status, and soil nutrient status vary between gypsum and K2SO4... 108

LIST OF APPENDIX TABLES

Table Page A.1. Nutrient concentration of ‘Draper’ highbush blueberry grown in different gypsum and organic matter amendments, either infected or non-infected with P. cinnamomi root rot………………………………………………... 128 A.2. Vegetative and root biomass of ‘Draper’ highbush blueberry grown in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m-1……………………………………………. 129 A.3. Total nutrient uptake of ‘Draper’ highbush blueberry grown in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m-1…………………………………………… 130 A.4. Leaf nutrient concentration of ‘Draper’ highbush blueberry grown in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m-1……………………………………………. 131 A.5. Soil solution nutrient concentration in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m-1…. 132 A.6. Mehlich III extractable nutrients in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m-1……. 133

CULTURAL CONTROLS FOR SUPRESSING Phytophthora cinnamomi ROOT ROT OF BLUEBERRY

Chapter One: General Introduction

John R. Yeo

History of the global blueberry supply chain Research into blueberry (Vaccinium spp.) cultivation began in the early 1900s, investigating mycorrhizal relationships, irrigation and drainage requirements, and, soil chemistry for blueberry mineral nutrition (Coville, 1910). High variability exists in fruit quality of wild‐type blueberries, and genetic improvement began in 1910 by Frederick Coville and Elizabeth White with interspecific pollen crosses of superior wild‐types selected for large fruit size (Coville, 1916). Prior to these improvement and cultivation efforts, blueberries were wild‐harvested throughout the native range, spanning from the North American boreal forests to the Florida Everglades, and were used as a food source for indigenous cultures and European settlements (Hummer, 2013). Significant genetic improvement was accomplished in the mid‐1900s by Arlen Draper, George Scott, and George Darrow, who exploited congruent ploidy levels amongst Vaccinium spp., creating many interspecific crosses (Finn et al., 2014). Growth of the blueberry cultivation industry proceeded relatively slowly throughout the 1900s due competing berry crops such as strawberry with more aggressive marketing campaigns (Martin and Garren, 1970). In the late 1900s and early 2000s, consumer demand for blueberries exponentially increased in response to marketing campaigns, and a global supply chain arose to satisfy consumer demand (Finn et al., 2014).

Global blueberry production increased from 64,769 metric tons in 1980 to 356,533 metric tons in 2011. Countries producing >1000 metric tons in 2011 included the United States, Canada, Poland, Mexico, Germany, Netherlands, Sweden, New Zealand, Lithuania, Russia, Romania, and France. States in the U.S. producing >1000 metric tons in 2011 included Michigan, Georgia, Washington, Oregon, New Jersey, California, Florida, and Mississippi (USDA ERS, 2013).

Origin, global distribution, and hosts of Phytophthora cinamomi Robert Delafield Rands (1890‐1970) first isolated Phytophthora cinamomi from stripe canker lesions on bark of plantation cinnamon trees (Cinnamomum burmannii Blume) in Sumatra while working in rubber industry development. Rand cultured and described the causal organism as a new species in the genus Phytophthora in 1921 (Zentmyer, 1980; Polhamus, 1971). P. cinamomi has the broadest host range of all known Phytophthora species with over 950 hosts, and has since been reported in >67 countries (Zentmyer, 1980). Plant hosts of P. cinamomi tend to be

woody and include economically important species in Cupressaceae, Ericaceae, Fagaceae, Lauraceae, and Fabaceae, and other taxonomic families (Zentmyer, 1980). Definitive information on the geographic origin and temporal global epidemiology of P. cinamomi is lacking, as inter‐continental dispersal likely occurred before phytopathological methods were developed to trace pathogen movements in international trade (Hodgson, 1943; Tryon, 1893).

Destruction of ecosystem functionality from P. cinamomi has occurred in the Jarrah forests in western Australia, where widespread forest dieback was first observed in 1922 (Shearer and Tippett, 1989), one year after the initial species description by Rands in 1921. However, P. cinamomi was not diagnosed as the cause of Jarrah forest dieback until 40 years later. Jarrah forests are mostly comprised of plant species in the families Proteaceae, Ericaceae, Dilleniaceae, and Myrtaceae, which are highly susceptible to infection by P. cinnamomi. Jarrah diebackcauses ecosystem collapse when net primary productivity by these species is eliminated, and no plant community shift to restore NPP in these ecosystems has occurred (Shearer and Tippett, 1989). The majority of the fundamental research investigation into P. cinamomi physiology has been in response to Jarrah dieback. However, no conclusive solutions for implementing corrective measures have been discovered (Jung et al., 2013).

Oregon blueberry production practices and status of Phytophthora cinamomi in the Oregon blueberry industry Root rot of blueberry caused by P. cinamomi has been reported as prevalent in Florida, Oregon, Georgia, Arkansas, New Jersey, and British Columbia production regions, and control accounts for the highest variable annual cost of blueberry production in Georgia (Clark et al., 1986; Fonsah et al., 2006; Strik and Yarborough, 2005). A survey of 55 commercial blueberry fields in Oregon detected Phytophthora cinamomi in 24% of fields sampled (Bryla et al., 2008). Two highly susceptible cultivars, ‘Duke’ and ‘Bluecrop’, comprised 42% of the fields surveyed in the study, and Phytophthora was recovered more frequently than expected from fields of these cultivars than from other cultivars sampled. Irrigation method was not correlated with Phytophthora incidence in this study, as 96% of the fields sampled were irrigated with overhead sprinklers.

Currently, the Oregon blueberry industry is adopting drip irrigation on new plantings for increased water use efficiency and fertigation capacity (Bryla et al., 2011). Phytophthora root rot disease severity increases with increasingly wet soils, and drip emitters dispense irrigation water over a much smaller area than sprinklers, creating a zone of higher water content conducive for the pathogen (Bryla and Linderman, 2007). In a replicated field experiment, increased Phytophthora severity under drip irrigation has been correlated with approximately 50% reduction in canopy cover, shoot weight, and root weight compared to sprinkler‐irrigated plants, after the first two years of establishment (Bryla and Linderman, 2007). As blueberries have a relatively long establishment period before growers begin to recoup planting costs, maximizing plant growth and health in the establishment period is essential for profitable production (Eleveld et al., 2005).

Traditional blueberry cropping systems such as high rates of organic amendment and raised beds likely provide a degree of Phytophthora suppression. However, root rot suppression by these methods has never been documented for blueberry, and root rot remains a significant problem in the industry.

Douglas fir (Pseudotsuga menziesii Franco) sawdust has been used historically in Oregon as both an amendment to increase soil organic matter and as a surface mulch for weed control primarily (White, 2006). Increasing cost of sawdust, along with the tendency to immobilize N fertilizer, is driving a trend in the industry of reducing sawdust application (Strik and Yarborough, 2005). In other areas in the United States, pine bark, hardwood bark, and peat are used as soil amendments for blueberry production, based on regional availability (Haynes and Swift, 1986; Miller‐Butler and Curry, 2009; Strik and Yarborough, 2005). Despite similarities between mulched raised beds used in blueberry production and the Ashburner system (see below), no correlation between sawdust mulch application and P. cinamomi incidence was detected in a field survey of Oregon blueberry fields (Bryla et al., 2008). While the suppressive capability of Douglas fir sawdust against P. cinamomi has never been measured directly in blueberry, potentially, cellulase enzymes produced by saprophytes to degrade the sawdust may inhibit P. cinamomi (Downer et al., 2002; Richter et al., 2011a).

Disease Control Fungicide subsection Phytophthora root rot of blueberries may be controlled by fungicide application in conventional farming systems. However, this option is not available to organic producers who must rely on cultural practices for Phytophthora suppression (Brannen et al., 2009). Mefenoxam, an enantiomer of metalaxyl, is soil applied and inhibits RNA polymerase activity. Since introduction in 1977, many isolates of P. cinamomi have evolved resistance to mefenoxam, limiting its utility for disease control (Hu et al., 2010). Multiple variations of phosphonate compounds, including phosphorous acid, phosphonic acid, alkali phosphite salts, and aluminum tris o‐ethyl phosphonate, are available for foliar application, are translocated systemically to roots, and provide plant protection against Phytophthora and Pythium pathogens (Landschoot and Cook, 2005). The specific mode of action of the phosphonate compounds is only partially understood, but it appears to inhibit fatty acid synthesis and pyrophosphate metabolism, and also disrupts many enzymatic pathyways of Phytophthora (Niere et al., 1994; Soulié et al., 1995, Stehman and Grant, 2000).

The Ashburner System of biological control The concept of creating a soil system suppressive to P. cinamomi dates to the late 1960s in Australia when it was observed that the pathogen would not penetrate rainforest soils adjacent to cleared land planted in Avocado and infested with P. cinamomi. Guy Ashburner, an Australian farmer, successfully suppressed the pathogen in his orchards by heavy application of mulch, manure, and calcium to mimic the rainforest soil conditions (Downer et al., 2002; Cook and Baker 1983). Variations of the Ashburner system have been adapted for suppression of P. cinamomi in Avocado production systems worldwide (Downer et al., 2002; Messenger et al., 2000a; Richter et al., 2011a).

Incorporation and mulch application of organic residues into soil can increase soil microflora populations, including organisms capable of suppressing Phytophthora spp. (Downer et al., 2001; Hoitink et al., 1977). In addition to stimulation of microbial antagonists, toxic compounds

in organic residues may provide direct pathogen suppression (Hoitink et al., 1977; Kayusa et al., 2006). Efficacy of organic residue incorporation for Phytophthora suppression varies, depending on the residue source and the level of decomposition. Composted hardwood bark amendment has been demonstrated as effective for suppression against P. cinamomi in pots, while pine bark was found to be less suppressive, and peat was not suppressive at all (Hoitink et al., 1977; Spencer and Benson, 1981). Eucalyptus mulch has also demonstrated efficacy for P. cinamomi suppression in field conditions (Downer et al., 2001). Pine sawdust mulch demonstrated no suppressive capacity in a potted study. However, a 24‐month field study reported suppression of P. cinamomi in Frazer fir mulched with both pine chips and pine bark (de Silva et al., 1999; Richter 2011b).

Microbial suppression of Phytophthora cinamomi in native soil ecosystems Microbial suppression of P. cinamomi has been widely documented and occurs through hyphal lysis of the pathogen by extracellular cellulase enzymes produced by soil micorflora (You and Sivasithamparam, 1994; Downer et al., 2001; Richter et al., 2011a). In response to hyphal lysis, P. cinamomi commonly forms sporangia for zoospore production as a survival mechanism, although sporangia abortion frequently occurs in suppressive soils (Nesbitt et al., 1979). Researchers studying the Ashburner soil and other suppressive and conducive soils in Australia attributed suppressive effects to higher populations of bacteria and actinomycetes in suppressive soils (Broadbent and Baker, 1974). Many subsequent investigations have correlated populations of various fungi, actinomycetes, pseudomonids, and total microbial activity with disease suppression. However, single biological agents have rarely been correlated with disease suppression in field settings (You and Sivasithamparam, 1994; You et al., 1996, Nesbitt et al., 1979, Termorshuizen et al., 2006; Aryantha et al., 2000, Stirling et al., 1992).

Biological control agents for biocontrol of Phytophthora cinamomi Dissertation research by A. de Silva (1999) reported reduction in root infection incidence by P. cinamomi on ‘Bluecrop’ blueberry in greenhouse studies. The plants were grown with pasteurized and non‐pasteurized Typic Hapludult field soil, containing 10% (v/v) rice bran P. cinamomi inoculum and an application of biocontrol agents, including Pseudomonas corrugate 114, Gliocladium virens GL‐21, Pseudomonas fluorescens PRA 25, and Bacillus pumilus T4.

Although P. cinamomi root colonization was significantly reduced compared to non‐treated controls, infection rates remained >38% of roots sampled, indicating none of the biocontrol agents provided curative effects, and only G. virens GL‐21 increased the stem diameter and only P. corrugate increased leaf number. In these studies, microbial agents, Trichoderma harzanum, Pseudomonas fluorescens 101, Pseudomonas fluorescens 101, and Pseudomonas fluorescens PF5, provided no suppression of % root infection by P. cinamomi. Strong antagonism against P. cinamomi by Myrothecium roridum has been reported on Persea indica seedlings. However, M. roridum is also an aggressive plant pathogen with many hosts, and commercialization as a biocontrol agent is unlikely (Gees and Coffey, 1989).

Calcium effects on P. cinamomi In addition to mulch application, heavy applications of Ca2+ are a component of the Ashburner system. High concentrations of Ca2+ have been shown to influence the asexual reproductive biology of various Phytophthora species (Hill et al., 1998; von Broembsen and Deacon 1997). Relatively few studies have thoroughly investigated fundamental Ca2+ influence on P. cinamomi specifically. However, Ca2+ appears to reduce sporangia production and decrease zoospore motility and viability (Irving and Grant, 1984; Messenger et al., 2000b).

The effects of Ca2+ on Phytophthora zoospores are not shared by other cations. Therefore, the effect of Ca2+ is not attributed to changes in osmotic pressure. Calcium ions cause rapid encystment of motile zoospores, often followed by immediate germination (Byrt et al., 1982, Irving and Grant, 1984). The ions increase adhesiveness of the zoospores during the encystment process, decreasing likelihood of transport with saturated flow through the soil (Gubler et al., 1988). Motile zoospores have thin cell walls and low energy reserves and, therefore, are less persistent in soil than hyphal structures. However, encysted zoospores are relatively persistent in soil and can act as latent infection agents, if soil conditions do not stimulate premature germination (Malajczuk et al., 1983). Premature stimulation of zoospore germination reduces pathogen persistence in soil, especially if microbial antagonists are present to lyse any newly‐ formed hyphae (Malajczuk et al., 1977).

Relationship between soil moisture matric potential and Phytophthora spp. root rot disease Soil moisture status governs rates of Phytophthora root rot disease progression by enabling repeating zoospore infection cycles near saturation. Soil moisture can be expressed as volumetric or gravimetric content, or as matric potential (Ψm) which is a measurement of energy required to overcome capillary forces and remove water from soil pores. Matric potential is a suction force, and is expressed as negative pressure in millibars (1000 mb = 1 bar) in this dissertation. For purposes of comparing high and low Ψm, this dissertation follows terminology guidelines set forth by the International Soil Science Society where more negative values farther from 0 are considered high Ψm, and values closer to 0 are considered low Ψm (Hillel, 1998). Matric potential influences many oomycete physiological processes including oospore and chlamydospore germination, sporangia formation, and zoospore discharge from sporangia (Reeves, 1974; MacDonald and Dunniway 1978a; 1978b). Sporangia are able to form under relatively high Ψm, but the repeating disease cycle is not triggered until Ψm thresholds approaching saturation allow release of motile zoospores from the sporangia (Kuan and Erwin, 1982; Hardy and Sivasithamparam, 1991).

Indirect sporangia germination is considered synonymous with zoospore discharge, however zoospore discharge is not equivalent to zoospore dispersal and infection. Matric potential near zero is commonly associated with zoospore infection (Zentmyer and Erwin, 1970), while indirect sporangia germination can occur at much higher Ψm. Bernhardt and Grogan (1982) reported P. parasitica and P. capsici zoospore discharge up to ‐300 mb based on observations of empty sporangia. Dunniway (1975) also reported P. drechsleri indirect sporangia germination at ‐300 mbar, however observed that zoospores recovered from soil samples at high Ψm were encysted and germinated very infrequently on selective media.

Dunniway (1976) again reported indirect sporangia germination by P. cryptogea up to ‐300 mbar, but placed seedling baits in the tension funnels at varying distance from the inoculum to determine infectivity and motility of zoospores presumably released at high Ψm. Zoospores were motile and infectious 5 mm from the sporangia up to ‐50 mb Ψm in UC‐type potting soil, and ‐10 mb Ψm in field soils. Differences in Ψm threasholds between soils were attributed to differences in pore size, with small pores likely inhibiting the helical patterns of zoospore motion. Despite

considerable reduction in motility at ‐50 mb Ψm, intermediate Ψm values between ‐300, ‐50, and ‐10 mb were not tested, therefore the true upper limits are not known.

MacDonald and Dunniway (1978b) investigated zoospore discharge in fine sandy loam using microscopic dilution techniques for P. megasperma and P. cryptogea at 0, ‐1, ‐5, ‐10, and ‐25 mb Ψm in fine sandy loam soil, sampled each hour for six hours. Zoospore density decreased dramatically at ‐10 mbar, and zoospores were not observed at ‐25 mb Ψm. Differences in upper

Ψm thresholds for zoospore discharge between MacDonald and Dunniway (1978b), and Dunniway (1976) are likely due to sampling method and sampling time. The baiting method with

UC‐type potting soil is more likely to detect zoospores and reveal the true upper Ψm threshold of Phytophthora species, as sampling time is longer, zoospore motility is not limited by pore size, and there are fewer potential problems related to sampling, handling, and counting zoospores.

Influence of water status on P. cinamomi root rot of blueberry Many soil amendments used in blueberry production are applied with the intention of creating increased drainage and porosity conditions favored by blueberry roots, and can significantly reduce the water holding capacity of soil (Spencer and Benson, 1981; White, 2006). Reducing the tendency for water to linger at low matric potential is imperative for prevention of P. cinamomi infection in blueberry production. Frequency of periodic saturation has been shown to increase severity of blueberry root rot (de Silva et al., 1999). Field irrigation systems are not designed typically to saturatethe soil. However, soil zones directly beneath drip emitters often exceed field capacity during irrigation events.

Efficacy of gypsum application for Phytohpthora root rot control in non‐blueberry crops Gypsum application is a common practice in for suppressing P. fragariae var. rubi in Red Raspberry. Field studies at Ithaca, NY with red raspberry cultivar ‘Heritage’ demonstrated a 48% increase in fruit yield on a site infested with P. fragariae var rubi, however the difference was non‐significant due to plot variation (Maloney et al., 2005). Trials in Clark County, WA investigating control of P. fragariae var rubi with gypsum failed to demonstrate gypsum main effects of root rot suppression in a gypsum x raised bed x solarization 2x2x2 factorial design. However, the gypsum + solarization + raised bed treatments increased primocane count and

fruit yield compared to non‐gypsum + solarization + raised bed treatments (Pinkerton et al., 2009).

Application of gypsum at 5% of total soil volume was effective for control of P. cinamomi on avocado in greenhouse trials, reducing infection of avocado by 71%. However only 1% v/v gypsum provided equivalent suppression when 10% v/v unspecified organic matter was included. When these treatments were applied to small plots in a mature avocado grove, there was no difference in root biomass after one year (Messenger et al., 2000a).

Response of Phytophthora cinamomi to temperature Temperature plays a critical role in the physiological development and pathogenicity of P. cinamomi. Shearer et al. (1987) reported different hyphal growth response rates to temperature for different isolates of P. cinamomi in vitro, however isolates tended to cause similar rates of lesion development across different temperatures in Eucalyptus marginata roots. Lesion development did not occur at 5 oC, proceeded very slowly at 15 oC, was greatest at 30 oC, and did not occur at 35 oC. Minimum temperature thresholds for hyphal growth in vitro vary between isolates of P. cinamomi, ranging from 9 oC to 12 oC. Maximum temperature for hyphal growth for most isolates ranges from 30 oC to 33 oC, however some isolates retain hyphal growth function up to 36 oC. Optimal temperature for in vitro hyphal growth of most isolates ranges from 21 oC to 30 oC (Zentmyer et al., 1976).

Mortality of Douglas‐Fir seedlings by P. cinamomi requires a minimum of 20 oC soil temperature and also sufficient moisture (Roth and Kuhlman, 1966). However some degree of infection likely occurs on Douglas‐Fir below 20 oC, as infection occurs on Frazer Fir at 14 oC, but mortality does not occur until 16 oC (Shew and Benson, 1983). Despite high virulence of P. cinamomi on Douglas‐Fir, deficient soil moisture when summer soil temperature rises above 20 oC is hypothesized to prevent widespread Douglas‐Fir dieback in the Pacific Northwest (Roth and Kuhlman, 1966).

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Chapter Two: Differences in highbush blueberry (Vaccinium corymbosum) cultivar susceptibility to Phytophthora cinnamomi

John R. Yeo, Jerry Weiland, Dan M. Sullivan, and David R. Bryla

Abstract Phytophthora cinnamomi causes root rot disease of blueberry, which impedes plant establishment and reduces annual fruit yields in many growing regions worldwide. Plant host tolerance is a promising tool to overcome economic losses while minimizing fungicide input to the environment. Most cultivars can be infected, but susceptibility differs among cultivars. To identify cultivars with superior tolerance to the disease, 18 cultivars and three advanced selections of highbush blueberry (Vaccinium spp.) were evaluated in three greenhouse experiments. Disease tolerance was based on relative shoot and root dry mass of infected plants compared to non‐infected controls of the same cultivar. Relative root infection incidence (infected vs. non infected plants of same cultivar) was also used to assess tolerance. Highly susceptible cultivars included ‘Duke’, ‘Draper’, ‘Bluetta’, ‘Blue Ribbon’, ‘Cargo’, ‘Last Call’, ‘Top Shelf’, and ‘Ventura’. More tolerant cultivars included ‘Legacy’, ‘Liberty’, ‘Aurora’, ‘Overtime’, ‘Reka’, ‘Clockwork’, ‘FC Selection 1’, and ‘FC Selection 3’. By choosing cultivars with more tolerance to P. cinnamomi, growers can reduce economic losses related to the disease. Although fungicides and cultural controls may provide some disease control, plant host tolerance is likely the most promising tool for avoiding disease when the pathogen is present. This research also suggests that including root rot tolerance as a selection criteria in a blueberry breeding program could result in cultivars with superior growth on infested sites. Introduction Highbush blueberry acreage is increasing globally in response to consumer demand, and plant breeding communities are responding by rapidly developing new cultivars for fruit quality, chilling hour requirements, and harvest timing for growing regions worldwide (Finn et al., 2014). The genetic background for most blueberry cultivars is broad, consisting of interspecific crosses of Vaccinium corymbosum, V. angustifolium, V. myrtilloides, V. darrowi, V. ashei, and V. constable (Hancock, 2006a; 2006b).

Disease tolerance is often overlooked in the selection process (Hancock, 2006b). However, it is of high importance for growers (Brannen et al., 2009). Phytophthora cinnamomi is a highly virulent root rot pathogen of highbush blueberry and is present in most growing regions worldwide (Strik and Yarborough, 2005). Production practices and fungicides provide mitigation for root rot disease, but genetic tolerance is an integral component of an integrated disease

22 management program (Bryla et al., 2011). Although P. cinnamomi is frequently found in blueberry production fields throughout the United States (Bryla et al., 2008), tolerance or susceptibility to the pathogen in many highbush blueberry cultivars is unclear. Most studies have evaluated southern highbush and rabbiteye blueberry cultivars but only a limited number of northern highbush cultivars have been evaluated (Bryla et al., 2008; Erb et al., 1987, Larach et al., 2009; Milholland and Galletta 1967, Smith 2002).

The objective of this study was to evaluate relative tolerance of 18 cultivars and advanced selections of highbush blueberry to P. cinnamomi in three greenhouse experiments.

Materials and Methods

Fall/Winter Cultivar Evaluation (Trial 1): Ten highbush blueberry cultivars (‘Aurora,’ ‘Bluecrop,’ ‘Bluegold,’ ‘Bluetta,’ ‘Draper,’ ‘Duke,’ ‘Elliott,’ ‘Legacy,’ ‘Liberty,’ and ‘Star’) were evaluated for tolerance to P. cinnamomi. Twelve plants of each cultivar were obtained as 5‐cm‐diameter tissue culture plugs (Fall Creek Nursery, Lowell, OR), transplanted into 2.6 L, 15‐cm‐diameter plastic pots containing Metro Mix 840PC potting media (Sun Gro Horticulture, Agawam, MA), and allowed to establish for 26 days before inoculation with P. cinnamomi. All plants were actively growing at the time of transplant (2 Sept. 2011) and were maintained in a greenhouse at constant 20 oC day/night air temperature. Photoperiod was extended to 16‐hr day length using two 1000 W high‐pressure sodium grow lights per greenhouse bench.

Inoculum was prepared using a single‐spore isolate of P. cinnamomi grown on 20 ml of potato dextrose agar (PDA) in 13, 84‐mm‐diameter petri plates. This isolate was originally obtained in 2010 from a naturally infected, field‐grown plant of ’Draper’ blueberry. The plates were fully colonized after 10 days, and the agar from each plate was excised and blended for 10 sec with 500 ml of distilled water at low speed (≈18,000 rpm) in a Waring blender (Conair Corporation, Stamford, CT). The blended slurry was then diluted to 6.5 L total volume, and a 0.5 ml aliquot was plated on PARP, a semi‐selective medium for Pythiaceous species (Kannwischer and Mitchell, 1978), to count colony forming units (cfu). Applied inoculum density was 76 cfu/ml.

Six plants of each cultivar were chosen at random for inoculation, with the remaining six serving as healthy, non‐inoculated controls. For each inoculated plant, 100 ml of the inoculum slurry was poured on the soil surface, followed by hand‐sprinkler irrigation. Inoculated and control plants were then placed in saucers on two greenhouse benches in a completely randomized design (CRD) with ≈25 cm between pots. Irrigation was supplied daily with an automated drip irrigation system, and supplemental hand‐watering was applied to maintain standing water in saucers for the duration of the experiment. Fertilization was applied weekly using diluted Miracle‐Gro azalea camellia rhododendron fertilizer (30‐10‐10; The Scotts Miracle‐Gro Company, Marysville, OH), supplying 26 mg N plant‐1 wk ‐1. Plants were destructively harvested 15 weeks after planting.

Spring/Summer Cultivar Evaluation (Trial 2): In order to determine whether there was an effect of season on disease development, the same cultivars evaluated in the Fall/Winter Cultivar Evaluation Experiment were evaluated again in the spring and summer of 2012. On 11 Apr. 2012, 12 actively‐growing tissue culture plants of each cultivar were obtained, transplanted, and allowed to establish, as described for trial 1. Temperature in the greenhouse was maintained at 25/20 oC (day/night). Supplemental lighting was not used, silver‐colored, reflective shade cloth (60% shade reduction) was placed over the top of the greenhouse to reduce heat load.

Six plants of each cultivar were inoculated 26 d after transplanting and arranged in a CRD according to the methods described above. The remaining six plants for each cultivar served as healthy, non‐inoculated controls. The final P. cinnamomi inoculum density for this experiment was 122 cfu/ml. All plants were maintained as described in trial 1, and were destructively harvested 20 weeks after planting.

Evaluation of New Cultivars and Advanced Breeding Selections (Trial 3): We evaluated several recently‐released blueberry cultivars and advanced selections from the Fall Creek Farm and Nursery private blueberry breeding program. ‘Draper’ and ‘Legacy’ were

24 also included as highly‐susceptible and somewhat‐tolerant standards, per results from Trial 1 and 2.

Fall Creek nursery provided twelve 5‐cm diameter plugs of cultivars Blue Ribbon, Cargo, Clockwork, Draper, Last Call, Legacy, Overtime, Reka, Top Shelf, Ventura, and advanced selections FC Selection 1, FC Selection 2, and FC Selection 3. Plants were transplanted on 16‐Sep into 15‐cm diameter pots with Sun Gro Metro Mix 840PC potting soil. Pots were arranged randomly on benches in a climate‐controlled greenhouse regulated near 20oC with supplemental lighting to maintain 16‐hr daylength. Each pot was placed in a separate saucer, which were maintained with standing water for the duration of the trial to promote disease development (de Silva et al., 1999). Fertilization was applied weekly using diluted Miracle‐Gro azalea camellia rhododendron fertilizer (30‐10‐10; The Scotts Miracle‐Gro Company, Marysville, OH), supplying 26 mg N plant‐1 wk ‐1. Plants were destructively harvested 16 weeks after planting.

First Inoculation (Trial 3): Six plants of each cultivar were inoculated with P. cinnamomi and the remaining six plants were not inoculated with P. cinnamomi. Inoculum was prepared by growing the pathogen on 17 petri plates containing potato dextrose agar (PDA). Plates were fully colonized after ten days, and the agar was excised and blended with 500 mL water at low speed in a Waring blender. The slurry was diluted with deionized water to 8.5 L inoculum volume. Inoculum (100 mL per plant) was applied to the soil surface of each inoculated pot seven days after planting. An inoculum sample was diluted plated on PARPH selective media to estimate pathogen propagule density. Propagules were most abundant yet individually distinct at the 10x dilution, and average propagule density was 656 propagules / mL.

Second Inoculation (Trial 3): Plants were inoculated with Phytophthora a second time, as it appeared root rot disease was not developing. Plants inoculated with Phytophthora had not developed foliar leaf symptoms, and several root samples from the ‘Draper’ susceptible controls did not grow Phytophthora when placed on PARPH selective media. Communication with the nursery indicated the plants

25 had been treated with mefenoxam which is a long‐residual soil active fungicide against Phytophthora, and likely inhibited disease development.

Plants were re‐inoculated on 9‐Nov‐2013 with V‐8 juice broth saturated vermiculite inoculum prepared on 15‐Apr‐2013 and stored in darkness at 20oC. Autoclave bags with Tyvek gas exchange filter patches were filled with 3 L of vermiculite, 150 mL filtered V‐8 juice, 1350 mL deionized water, and 3 g CaCO3. Bags were autoclaved, and one PDA petri plate fully colonized with P. cinnamomi was sectioned into >100 pieces, and added to each bag. Density of P. cinnamomi propagules was quantified between 4‐Nov and 9‐Nov, and sufficient vermiculite was added to 0.2% water agar to achieve the original inoculation density of 650 propagules /mL. One hundred mL of the suspension was added to the surface of each infected pot on 9‐Nov, followed by irrigation.

Destructive harvest and growth response evaluation: Plants in all three trials were destructively harvested and partitioned into shoot and root components. Stems and leaves were bagged together, dried, and weighed to determine shoot mass. Potting media was washed away from the root systems, and ten 1‐cm root sections were sampled from each plant and placed on PARPH selective media to estimate percent root infection (Tsao and Guy, 1977). Petri plates were monitored periodically for 10 days to count infected root sections. Remaining roots were oven‐dried at 100 oC and weighed to determine root mass.

Data analysis and statistics: Due to differences in absolute vigor between cultivars, susceptibility to P. cinnamomi infection was expressed as incidence of infected roots, and mass of infected plants relative to mass of non‐infected plants of the same cultivar (Allardyce et al., 2012). Data were analyzed using the general linear model procedure in Statgraphics Centurion XVI statistical software, and mean separation were revealed via Fisher’s LSD procedure. The first and second experiments were combined in one statistical model including terms cultivar, trial, and the trial x cultivar interaction as sources of variation. Mean separation was performed for each trial separately using Fisher’s LSD test because the trial x cultivar interactions were significant at α=0.05. The

26 third experiment included a different set of cultivars than the first and second, and only the cultivar term was included in the model. Mean separation was performed using Fisher’s LSD test. Residuals from the data had normal distribution and no data transformation was required before analysis.

Results: Many inoculated cultivars exhibited symptoms of P. cinnamomi infection including fine root necrosis, reduced root and shoot mass relative to the controls, red‐orange leaf discoloration, and in extreme cases, leaf necrosis and senescence. However, a wide variation in disease response existed between the cultivars.

Trial 1 and 2: ‘Bluetta’, ‘Draper’, and ‘Duke’ were the most susceptible cultivars we evaluated. Roots from these cultivars failed to grow from the nursery plugs into the surrounding infected soil, and root and shoot mass were dramatically reduced compared to the controls (Table 1; 2). ‘Aurora’, ‘Liberty,’ ‘Legacy’, and ‘Star’ were most tolerant to Phytophthora infection. ‘Legacy’ had the the highest relative root mass of all cultivars, but root mass of the infected plants was only 56.6% of the non‐infected controls, indicating tolerance to infection rather than absolute resistance (Table 2). Incidence of infected root sections was very high for all cultivars, ranging from 38% to 85% of 1‐cm roots positive for P. cinnamomi in trial 1. There was no significant difference in root infection incidence in trial 2, and incidence ranged from 45% to 81% of sampled root sections (Table 3).

Trial 3: ‘Overtime,’ ‘Clockwork,’ and ‘Reka’ and ‘Legacy’ had the highest level of tolerance to P. cinnamomi infection. Shoot mass of infected Overtime plants was not significantly different than non‐infected controls (Table 4). ‘Blue Ribbon,’ ‘Cargo,’ ‘Draper,’ ‘FC Selection 2,’ FC Selection 3,’ ‘Last Call,’ ‘Top Shelf,’ and ‘Ventura,’ and were the most susceptible cultivars with <30% relative root mass of the non‐infected controls. There was no significant difference in relative root growth of these cultivars (Table 4). Trial 3 was not repeated, however cultivars ‘Legacy’ and

‘Draper’ were included in the assay as controls, and performance of these controls were congruent with trial 1 and 2, indicating similar disease pressure.

Discussion: Trial 1 and 2: High reductions in root growth of all cultivars tested were likely exacerbated by excellent conditions for disease development including potting media near saturation and conducive temperature for pathogen growth (Zentmyer, 1980). In a field irrigation experiment with ‘Duke’ and ‘Elliott’ inadvertently infected by P. cinnamomi, ‘Duke’ failed to establish, however ‘Elliott’ outgrew the infection and established successfully (Bryla, personal communication). In our trials, ‘Elliott’ failed to demonstrate high levels of tolerance. Cultivars that exhibited more tolerance than ‘Elliott’ in our trials may demonstrate even better field performance than Elliott in field conditions with Phytophthora presence . High susceptibility of ‘Duke’ to P. cinnamomi in our trials is congruent with field observations, and also other greenhouse screening work (Bryla 2008; Larach et al., 2009).

The significant cultivar x trial interaction in shoot growth response reflects differences in ‘Aurora’ and ‘Legacy’ between the two trials. ‘Aurora’ had high shoot growth in trial 1, and poor shoot growth in trial 2, while ‘Legacy’ had higher shoot growth in trial 2 than trial 1 (Table 1). There was no significant cultivar x trial interaction for the root growth response, or the root infection incidence data (Table 2; Table 3). Relative root mass and infection incidence are perhaps more appropriate metrics for assessing tolerance to a root pathogen, and the interaction in shoot mass between the two trials may be a response to differences in temperature in the greenhouse or artificial season prolongation in trial 2.

Root infection incidence was significantly correlated with both decreased root and shoot mass (Figure 1; 2). Cultivars with higher root or shoot mass than predicted by the regression coefficients may prove valuable as parents in a plant breeding program selecting for root rot tolerance. Cultivars vary significantly in vigor regardless of infection, and more vigorous plants may accumulate more shoot mass than less vigorous plants at the same infection level. All plants in these experiments were grown in similar disease pressure conditions, and differences

28 in root infection incidence may be attributed to either plant host defenses against tissue colonization, or lack of zoospore attraction to roots (Erb et al., 1986; Milholland, 1975). Validation of trial 1 was performed by repeating the experiment again (trial 2), but validation of trial 3 was accomplished by including ‘Legacy’ as a tolerant standard, and ‘Draper’ as a highly susceptible standard. Root and shoot mass of infected ‘Legacy’ relative to mass of non‐infected controls was slightly lower in trial 3 than trial 1 and 2. In trial 3, ‘Legacy’ performed similarly to ‘Clockwork,’ and FC Selection 1, and both ‘Overtime’ and ‘Reka’ outperformed ‘Legacy’ indicating superior disease tolerance. Relative infected shoot mass of ‘Draper’ was higher in trial 3 than in trial 1 and 2, but relative infected root mass was equivalent. In trial 3, ‘Draper’ performed similarly to ‘Top Shelf,’ ‘Ventura,’ ‘Last Call,’ ‘Cargo,’ and ‘Blue Ribbon,’ indicating these cultivars are highly susceptible to root rot (Table 1; 2; 3). Trial 3 differed from Trial 1 and 2 by failed infection from the first inoculation, possibly due to mefenoxam residue in the nursery plug. The second inoculation was successful, thus a longer establishment period elapsed before infection than trial 1 and 2, and plants may have a larger before succumbing to infection.

The assay we used for screening cultivars for tolerance or susceptibility provided relatively consistent results for both ‘Draper’ and ‘Legacy’ across three trials. However, the assay was subject to space constraints and relatively long duration, which limited the number of cultivars that we evaluated. For breeding programs rapidly screening larger numbers of selections, the physical dimensions and trial duration of our assay is likely impractical. Similar inoculation methods could be used with smaller plants, smaller pots, possibly shorter duration, and a more efficient arrangement on greenhouse benches. In our assay, each plant was maintained separately in individual saucers to ensure independence of the experimental units. For more efficient screening of large numbers of selections, whole trays of plants could be either infected or non‐infected at the expense of replication and sampling unit independence.

Genetic tolerance or resistance to root rot disease is likely the most promising tool for minimizing the economic losses caused by the disease. On sites where root rot disease is present or suspected, growers should avoid planting susceptible cultivars. This study identified several highly susceptible cultivars including ‘Duke’, ‘Draper’, ‘Bluetta’, ‘Blue Ribbon’, ‘Cargo’, ‘Last Call’, ‘Top Shelf’, and ‘Ventura,’ but this list is not exhaustive. For breeders, including

29 parents with Phytophthora tolerance such as ‘Legacy’, ‘Liberty’, ‘Aurora’, ‘Overtime’, ‘Reka’, ‘Clockwork’, ‘FC Selection 1’, and ‘FC Selection 3’ and imparting Phytophthora selection pressure on the progeny could lead to further development of new cultivars with both desirable fruit quality and also disease tolerance.

Works Cited

Allardyce, J.A., J.E. Rookes, and D.M. Cahill. 2012. Defining plant resistance to Phytophthora cinnamomi: a standardized approach to assessment. J Phytopathology 160:269‐276.

Bryla D.R., R.G. Linderman, and W. Q. Wang. 2008. Incidence of Phytophthora and Pythium infection and the relation to cultural conditions in commercial blueberry fields. Hortscience 43:260‐263.

Bryla D.R., J.L. Gartug, and B.C. Strik. 2011. Evaluation of irrigation methods for highbush blueberry – I. Growth and water requirements of young plants. HortScience 46:195‐101.

Brannen P.M., P. Harmon, and D.S. NeSmith. 2009. Utility of phosphonate fungicides for management of Phytophthora root rot of blueberry. Proc IX IS on Vaccinium. Eds: K.E. Hummer et al. Acta Hort 810:331‐340. de Silva A., K. Patterson, C. Rothrock, and R. Mc New. 1999. Phytophthora root rot of blueberry increases with frequency of flooding. HortScience 34:693‐695.

Erb W.A., J.N. Moore, and R.E. Sterne. 1986. Attraction of Phytophthora cinnamomi zoospores to blueberry roots. HortScience 21:1361‐1363.

Hancock J.F. 2006a. Highbush blueberry breeders. HortScience 41:20‐21.

Hancock J.F. 2006b. Northern highbush blueberry breeding. Proc. VIII IS on Vaccinium Culture. Acta Hort. 715

Kannwischer, M.E. and Mitchell, D.J. 1978. The influence of a fungicide on the epidemiology of black shank of tobacco. Phytopathology 68: 1760‐1765.

Larach A., X. Besoain, and E. Salgado. 2009. Crown and root rot of highbush blueberry caused by Phytophthora cinnamomi and P. citrophthora and cultivar susceptibility. Sciencia e Investigación Agraria 36:433‐442.

Milholland R.D., and G.J. Galletta. 1967. Relative susceptibility of blueberry cultivars to Phytophthora cinnamomi. Plant Disease Reporter 51:999‐1001.

Milholland R.D. 1975. Pathogenicity and histopathology of Phytophthora cinnamomi on highbush and rabbiteye blueberry. Phytopathology 65:789‐793.

Smith B.J. 2002. Susceptibility of southern highbush blueberry cultivars to Phytophthora root rot. Proc. 7th IS on Vaccinium. Acta Hort 574:75‐79.

Strik B. C., and D. Yarborough. 2005. Blueberry production trends in North America 1992 to 2003, and predictions for growth. HortTechnology 15:391‐398.

Tsao, P.H. and Guy, S.O. 1977. Inhibition of Mortierella and Pythium in a Phytopthora isolation medium containing hymexazol. Phytopathology 67: 796‐801.

Tables and Figures

Table 1. Shoot mass of P. cinnamomi infected blueberry plants, relative to non‐infected shoot growth of the same cultivar. Cultivar Spring/Summer Cultivar Evaluation Fall/Winter Cultivar Evaluation Rank Relative % Rank Relative % Aurora 1 99.0 ± 15.1 d 3 57.2 ± 10.2 b Bluecrop 6 53.2 ± 3.1 ab 4 56.2 ± 5.3 b Bluegold 7 52.6 ± 4.0 ab 7 52.0 ± 9.4 b Bluetta 10 37.1 ± 6.2 a 10 19.4 ± 4.3 a Draper 5 53.3 ± 2.6 ab 8 46.1 ± 6.1 ab Duke 9 49.7 ± 4.9 a 9 42.4 ± 3.0 ab Elliott 8 49.9 ± 3.9 a 5 56.2 ± 4.4 b Legacy 2 74.3 ± 6.0 c 1 112.7 ± 20.4 c Liberty 3 69.5 ± 10.2 bc 2 102.9 ± 14.6 c Star 4 69.4 ± 2.5 bc 6 52.5 ± 4.3 b

p‐value <0.0001 <0.0001 Combined Model p‐value Cultivar <0.0001 Trial 0.7825 Trial*Cultivar 0.0002

Table 2. Root mass of P. cinnamomi infected blueberry plants, relative to root mass of non‐ infected plants of the same cultivar. Cultivar Trial 1 Trial 2 Rank Relative % mass Rank Relative % mass Aurora 3 49.4 ± 2.5 e 2 38.9 ± 5.2 bc Bluecrop 5 36.7 ± 4.5 cd 7 23.3 ± 2.5 ab Bluegold 6 30.4 ± 1.5 bc 6 26.0 ± 3.4 ab Bluetta 8 25.3 ± 2.7 abc 10 17.7 ± 3.3 a Draper 10 17.4 ± 2.3 a 9 21.3 ± 2.3 ab Duke 9 19.7 ± 1.7 ab 8 21.7 ± 1.8 ab Elliott 7 29.3 ± 4.6 abc 4 29.5 ± 2.9 ab Legacy 4 45.7 ± 6.5 de 1 56.6 ± 16.9 c Liberty 1 52.1 ± 8.0 e 3 31.2 ± 9.1 ab Star 2 50.2 ± 4.0 e 5 27.8 ± 2.6 ab

p‐value <0.0001 0.0088 Combined Model p‐value Cultivar <0.0001 Trial 0.0154 Trial*cultivar 0.0692

Table 3. Incidence of 1‐cm root sections infected with P. cinnamomi. Cultivar Trial 1 Trial 2 Rank % Root Infection Rank % Root Infection Aurora 2 41.7 ± 8.7 ab 4 55.0 ± 13.8 ns Bluecrop 5 61.7 ± 7.9 bcd 2 51.7 ± 10.8 ns Bluegold 6 61.7 ± 6.5 bcd 10 81.7 ± 4.8 ns Bluetta 10 85.0 ± 5.0 e 9 73.3 ± 15.8 ns Draper 7 63.3 ± 7.6 bcde 3 53.3 ± 8.4 ns Duke 9 81.7 ± 4.0 de 8 73.3 ± 8.4 ns Elliott 4 60.0 ± 5.2 abcd 5 60.0 ± 13.7 ns Legacy 3 53.3 ± 10.9 abc 1 45.0 ± 12.3 ns Liberty 1 38.0 ± 8.0 a 6 66.7 ± 9.5 ns Star 8 65.0 ± 10.9 cde 7 68.3 ± 10.1 ns

p‐value 0.0020 0.3929 Combined Model p‐value Cultivar 0.0074 Trial 0.6958 Trial*cultivar 0.3924

Table 4. Root infection incidence, and relative shoot and root mass compared to non‐infected plants of the same cultivar. Trial 3.

Infected 1‐cm root Treatment Rank Relative Shoot Mass (%) Rank Relative Root Mass (%) Rank sections (%) Blue Ribbon 13 32.0 ± 4.1 a 13 10.7 ± 0.4 a 12 81.7 ± 5.4 c Cargo 10 44.7 ± 9.6 abc 7 27.1 ± 8.8 abc 10 75.0 ± 5.6 c Clockwork 4 71.5 ± 5.3 def 3 54.3 ± 7.6 de 2 50.0 ± 11.3 ab Draper 7 58.5 ± 8.4 bcde 10 18.8 ± 3.6 ab 11 78.3 ± 4.8 c FC Selection 1 5 66.7 ± 8.9 cdef 5 33.2 ± 11.2 bc 5 66.7 ± 12.0 bc FC Selection 2 8 50.8 ± 12.3 abcd 6 29.4 ± 4.6 abc 7 68.3 ± 7.9 bc FC Selection 3 6 66.0 ± 2.5 cdef 9 24.2 ± 4.8 abc 6 68.3 ± 8.7 bc Last Call 11 37.3 ± 11.1 ab 8 25.7 ± 8.8 abc 3 61.7 ± 7.9 bc Legacy 2 83.1 ± 5.3 fg 4 40.4 ± 8.1 cd 8 70.0 ± 8.6 bc Overtime 1 100.7 ± 5.6 g 1 71.2 ± 12.5 e 1 34.0 ± 11.2 a Reka 3 78.0 ± 9.9 efg 2 63.7 ± 11.2 e 4 63.3 ± 12.6 bc Top Shelf 12 33.1 ± 6.7 a 12 13.8 ± 1.9 ab 13 83.3 ± 4.9 c Ventura 9 47.1 ± 9.2 abc 11 17.6 ± 3.2 ab 9 73.3 ± 7.1 bc

p‐value <0.0001 <0.0001 0.0216

120% ‐

non 100%

cultivar

80% y = ‐0.9941x + 1.219 same

compared R² = 0.3282

the 60% of

growth

plants

40% shoot

infected 20% Relative

0% 30% 40% 50% 60% 70% 80% 90% % Infected 1‐cm root sections Figure 1. Correlation between root infection incidence and shoot mass of 10 blueberry cultivars infected with P. cinnamomi, relative to shoot mass of non‐infected plants of the same cultivar. Data from trial 1 & 2 only. Each data point represents the average of six plants evaluated in one of two trials.

60%

infected 50% ‐ y = ‐0.6252x + 0.7125

non R² = 0.4387

to 40% cultivar

same 30% compared

the

20% growth

plants root 10% Relative 0% 30% 40% 50% 60% 70% 80% 90% % Infected 1‐cm root sections

Figure 2. Correlation between root infection incidence and root mass of 10 blueberry cultivars infected with P. cinnamomi relative to root mass of non‐infected plants of the same cultivar. Data from trial 1 & 2 only. Each data point represents the average of six plants evaluated in one of two trials.

Chapter Three: Evaluation of gypsum and organic matter soil amendments for suppressing Phytophthora cinnamomi root rot disease of blueberry

John R. Yeo, Jerry Weiland, Dan M. Sullivan, and David R. Bryla

Abstract Phytophthora cinnamomi is a soilborne pathogen that causes root rot disease of highbush blueberry (Vaccinium corymbosum L.), severely reducing growth and yield. Chemical fungicides are available to suppress this disease, but these chemicals are prohibited within certified organic production systems. Other workers have demonstrated the efficacy of gypsum and organic soil amendments in suppression of Phytophthora root rot in avocado, raspberry, and frazer fir. Gypsum provides soluble Ca2+ which inhibits zoospore motility, and organic amendments may induce hyphal lysis. Three greenhouse pot experiments were conducted to evaluate efficacy of soil amendments in suppressing P cinnamomi root rot disease. 'Draper', a disease susceptible cultivar, was grown in all experiments. Soil amendments included composts (derived from yard debris, dairy solids, or municipal biosolids + bark), incorporated at 20% v/v into soil, or gypsum incorporated at 5% v/v. Amendment effects on root and shoot mass were also evaluated without P. cinnamomi inoculation, in an effort to separate amendment effects on disease from other factors that might influence plant growth. Organic amendments did not provide significant disease suppression in any of the experiments. When plants were inoculated with disease, gypsum enhanced plant growth and decreased root infection incidence in one of the experiments, but not the other two experiments. High rate gypsum application did not reduce shoot or root biomass when plants were grown in the absence of P. cinnamomi inoculation. An additional greenhouse experiment was conducted to determine the application rate of gypsum required for disease suppression. The highest gypsum rate corresponded with the 5% v/v rate used in earlier experiments. Ten incremental rates of gypsum [0 to 96 cmol (+)/ kg soil] were applied, and soluble Ca in soil solution was monitored using mini‐lysimeters (RhizonTM tubes). Gypsum addition increased soluble Ca in soil from 15 mg/L (no gypsum added) to 450 mg/ L when gypsum was added at a rate of 15.6 cmol (+) / kg soil. At greater gypsum application rates, soluble Ca concentration in soil solution did not change, remaining at a plateau value of 450 mg/L. Gypsum rates that maximized Ca in soil solution (450 mg/L) provided the greatest disease suppression, as demonstrated by reduced root infection, and increased shoot biomass. Results of these experiments suggest that gypsum may reduce blueberry root rot disease, and that blueberry is tolerant of high rate gypsum applications. The mechanism proposed by others for disease suppression by gypsum (interaction of soluble Ca in soil solution with the pathogen) is supported by our findings. The lack of disease suppression from organic amendments observed

39 in these experiments may be attributed to high disease pressure, extreme host susceptibility, or that the amendments truly did not influence pathogenesis.

Introduction Phytophthora cinnamomi (Chromista; Oomycota) is a serious pathogen of >800 plant hosts and has a global distribution (Zentmyer, 1980). Host resistance, chemical fungicides, and manipulation of the soil environment can disfavor pathogenicity. P. cinnamomi causes root rot disease of highbush blueberry (Vaccinium corymbosum L.), and is a problem facing many growing regions worldwide. Cultivars vary in susceptibility. However, the most widely planted cultivars are susceptible (Larach et al., 2009; Smith 2002; Milholland and Galletta, 1967). Chemical controls are relatively effective for disease control, including mefenoxam, and phosphite products (Brannen et al., 2009). Organic blueberry production in the Northwestern United States is increasing, but chemical tools are not available to organic growers who must rely on cultural control for disease management.

Suppression of soilborne Phytophthora species has been accomplished in some cases by calcium or organic amendment, creating a soil environment inhospitable to hyphal persistence, sporangia production, and zoospore viability. Guy Ashburner, an Australian farmer, demonstrated substantial P. cinnamomi suppression in avocado (Persea americana Mill.) by heavy application of mulch, manure, and calcium in the 1960s (Downer et al., 2002; Cook and Baker 1983).

Variations of the Ashburner system have been adapted for Phytophthora suppression in production systems worldwide including avocado, raspberry, frazer fir, and bell pepper (Downer et al., 2002; Messenger et al., 2000a; ; Nesbitt et al., 1979; Richter et al., 2011, Chellemi, 2006; Costa et al., 1996). Phytophthora suppression by mulches is reported to be due to enhanced β‐ 1,3, 1,4, and 1,6 glucanase activity causing sporangial abortion, hyphal lysis, and zoospore trapping (Downer et al., 2001; Nesbitt et al., 1979; Richter et al., 2011, Chellemi, 2006; Costa et al., 1996). Variation in chemical and biological properties exists between organic amendments, as does the degree of disease suppression (Hoitink et al., 1976; Hoitink and Fahy, 1986).

High levels of calcium in soil have been shown to decrease zoospore motility and cause premature encystment to non‐infectious status (Messenger et al., 2000b). Zoospores are released in mass when temperature and moisture permit. They seek new roots via chemical signals, and are responsible for a rapid repeating disease cycle. Management practices that reduce zoospore production and survival are critical to an integrated disease management program. Gypsum (Ca2SO4 ∙ 2 H2O) provides soluble calcium that can mediate Phytophthora root rot suppression in red raspberry and avocado (Messenger et al., 2000a; Menge et al., 1994; Pinkerton et al., 2009).

Highbush blueberry is adapted to soils with low pH, soluble salts, and high organic matter. Therefore high rates of soluble salt addition as gypsum for disease suppression could pose a risk of salt injury to the plant. The purpose of our experiments were to 1. evaluate organic amendments, with or without a high rate of gypsum amendment for suppressing root rot on diseased plants, 2. evaluate the effect of gypsum and organic amendments on growth of non‐ diseased plants, and 3.) to model calcium solubility in soil solution in response to increasing rates of gypsum.

Methods Organic and Gypsum Amendment Experiments Spring ‐ Summer 2011 Experiment; Repeated Winter 2011‐2012 Five organic soil amendments, both with and without an additional gypsum amendment, and a gypsum amendment alone, were evaluated for their ability to suppress Phytophthora cinnamomi root rot of blueberry. A total of thirteen treatments were evaluated, including two controls. Six treatments consisted of either none, or one of five organic amendments mixed with Willamette loam field soil (Fine‐silty, mixed, mesic Pachic Ultic Argixeroll) at 20% of total volume (v/v). The five organic amendments were: composted bark with biosolids (Tagro, Tacoma, WA,), composted dairy solids (Oregon State University dairy), Douglas‐fir sawdust aged 1‐yr (Lane Forest Products, Eugene, OR), canadian sphagnum peat (Lakeland®, SunGro, Belvue, WA), and municipal yard debris compost (Rexius, Eugene, OR). An additional six treatments consisted of the same six organic amendment regimes, but also included gypsum added at 5% v/v. A

41 fungicide control treatment with no gypsum or organic matter was treated with aluminum tris o‐ethyl phosphonate (Aliette WDG) soil drench (1L water and 3.53g Aliette WDG per plant).

Root and shoot biomass accumulation was evaluated both in the presence and absence of P. cinnamomi inoculum. The isolate of P. cinnamomi used for both experiments was recovered from an infected ‘Draper’ blueberry plant in Linn County, OR in 2010. Inoculum was prepared by adding 1500 mL of 10% clarified V8 juice broth (3 g CaCO3 mixed with 150 mL V8 juice filtrate and and 1350 mL deionized water) to 3 L vermiculite in an autoclavable spawn bag with a Tyvek filter patch (Fungi Perfecti, Olympia WA). After autoclaving the vermiculite mixture, a petri plate containing 20 mL potato dextrose agar (PDA) fully colonized by P. cinnamomi was sliced into > 100 pieces and added to the bags. Bags were heat sealed and incubated > 1 month at 20o C with weekly mixing by hand to evenly distribute the inoculum. Vermiculite inoculum was incorporated at 5% v/v into each of the thirteen gypsum and organic amendment regimes to establish an inoculated set of treatments, whereas sterile vermiculite was incorporated at 5% to establish the non‐inoculated set of treatments. Soil for each treatment was then distributed individually into 15‐cm diameter 3L pots, with six replicate pots for each treatment. In the summer experiment, 5‐cm‐diameter plugs of highbush blueberry cultivar ‘Draper’ were transplanted into each pot in April 2011 using plants that were just breaking dormancy. In the winter experiment, planted in October 2011, the plants still had green leaves, but may have been entering dormancy. Greenhouse temperature was regulated at 20oC for both trials, which was maintained by cooling in the summer, and heating in the winter. The greenhouse was covered in silver‐colored 60% shade cloth during the summer experiment, and supplemental lighting with HPS bulbs was adjusted throughout both experiments to maintain 16‐hr daylength. Plants were watered daily by hand, and fertilized once weekly with Miracle‐Gro Water Soluble Azalea Camellia Rhododendron fertilizer (30‐10‐10) supplying 26 mg N plant‐1 wk‐1.

The experimental design was a 6 organic matter level x 2 gypsum level x 2 P. cinnamomi infection level factorial split plot RCB design with six replications. Replications were randomly split into infected and non‐infected main plots, and the gypsum and organic matter treatment combinations were randomly assigned within the main infection plots. Additionally, a fungicide control treatment was assigned within each infection main plot.

Plants were grown in the greenhouse for 14 weeks before destructive harvest, partitioning into shoot and root components. All soil was washed from the roots, and ten 1‐cm root sections were sampled at random from each infected plant and placed on PARPH media selective for Phytophthora species (Tsao and Guy, 1977). Infection incidence was estimated based on the percentage of infected roots. Shoot and root components were dried and weighed to determine biomass. Propagules of P. cinnamomi in soil were estimated the end of the experiments by spreading 0.5 mL of 20:1 0.2% water agar:soil suspensions on petri plates containing PCH media (Erwin and Ribeiro, 2005). Plates were incubated 5 d at 25oC, washed of soil, and P. cinnamomi colonies were counted by microscopy at 100x magnification.

Chemical analyses of composts, soils, and plant tissue were only performed for the spring‐ summer 2011 experiment. Chemical analyses of composts and the soil + gypsum + compost mixes were performed by Brookside Laboratories Inc. (New Bremen, OH). Soil from three of the six replications of non‐infected plants was sampled for chemical analysis and analyzed for pH (1:1 soil:water), organic matter loss on ignition, Bray I P, and Mehlich III extractable P, Mn, Zn, B, Cu, Fe, Al, S, Ca, Mg, and K via the Brookside S001P soil test package. One representative sample of each compost was analyzed for chemical content via deionized water extraction and analysis for pH, NO3, NH4, Ca, Mg, P, K, Na, S, B, Mn, Zn, Fe, Al, Mo, and Cu via the S003 soil‐less media test package. All stems and leaves from both non‐infected and infected plants were oven dried, ground, homogenized, and analyzed at the USDA‐ARS HCRL, Corvallis OR. C and N were analyzed by dry combustion in a Leco Truspec analyzer. Tissue was digested in 67‐70% w/w nitric acid at 20 psi (190oC) in a Anton Paar Multiwave 3000 microwave , and analyzed for P, K, and micronutrients by ICP‐AES using a Perkin Elmer Optima 4300 instrument.

Summer 2013 Experiment The main effects of organic matter and gypsum incorporation were evaluated using fewer treatments than the 2011‐2012 experiments. The experiment was a 2x2 completely randomized factorial design including gypsum incorporation at 5% v/v, yard debris compost (Rexius, Eugene, OR) at 20% v/v, a combination of both amendments, and no amendment. The fungicide control treatment [fosetyl‐Al (Aliette WDG) drench (1L water and 3.53g Aliette WDG per plant)] did not

43 receive gypsum or organic soil amendment. Each soil amendment treatment was evaluated with and without 5% v/v vermiculite P. cinnamomi inoculum added. Inoculum was prepared with methods described for the spring and winter 2011 experiments.

Five‐cm diameter plugs of highbush blueberry cultivar ‘Draper’ were transplanted into 15‐cm diameter pots containing the amended soil mixes in 12‐Jun 2013 using green‐growing plants that had been sheared approximately 12‐cm high. Plants were grown in a greenhouse covered with silver‐colored 60% shade cloth and maintained at 25oC with daily irrigation and no supplemental lighting. Fertilizer was supplied using Miracle‐Gro Water Soluble Azalea Camellia Rhododendron fertilizer (30‐10‐10) supplying 26 mg N plant‐1 wk‐1. The experiment was planted in a completely randomized design, and destructively harvested after nine weeks on 15‐Aug.

Plants were partitioned into shoot and root components. All soil and potting media was washed from the rootballs, and ten 1‐cm root sections were sampled at random from each infected plant and placed on PARPH media as described above. Infection incidence was estimated based on the percentage of infected roots. Propagule density of P. cinnamomi in soil were estimated the end of the experiments by dilution and spreading on PCH media as with the spring and fall 2011 experiments. Shoot and root components were dried and weighed to determine biomass.

Minimum gypsum rate for disease suppression A final greenhouse experiment was performed to determine the minimum amendment rate of gypsum required for suppressing P. cinnamomi infection of blueberry. Malabon silty clay loam (Fine, mixed, superactive, mesic Pachic Ultic Argixeroll) was collected from Lewis Brown Horticulture Farm, Corvallis, OR and screened through 1.3‐cm mesh. Chemical analysis for pH, Ca, Mg, K, Bray I P was performed via the W1 soil test package by Kuo Labs, Othello, WA. Soil CEC was determined by ammonium acetate exchange via the Total CEC soil test package by Brookside Laboratories Inc. (New Bremen, OH). Soil CEC was 24.4 cmol (+) / kg, pH was 5.3, Ca was 10.3 cmol (+)/ kg, Mg was 5.9 cmol (+)/ kg, K was 0.57 cmol (+)/ kg, total bases were 16.8

‐ cmol (+)/ kg, Bray I P was 23 mg / kg, and NO3 was 0.5 mg / kg. Gypsum was mixed into the soil at 0, 6, 12, 18, 24, 36, 48, 60, 72, and 96 cmol (+ )/ kg soil, along with 2% v/v vermiculite P.

44 cinnamomi inoculum prepared with methods described in the gypsum and organic amendment section. Gypsum additions to soil were equal to 0, 5.7, 11.4, 17.1, 22.9, 34.3, 45.71, 57.1, 68.6 and 91.4 g gypsum / kg dry soil.

Highbush blueberry cultivar ‘Draper’ in 5‐cm diameter plugs were transplanted into 15‐cm diameter, 3‐L pots containing the gypsum amended soils on Feb 29, 2012 and arranged in a CRD with six replications in the greenhouse. Pots were maintained in saucers of standing water for the first 13 weeks, and raised on blocks for the second 13 weeks. Supplemental lighting was provided with high pressure sodium (HPS) bulbs to maintain 16‐hr daylength during spring, and silver‐colored 60% shade cloth was placed over the greenhouse during the summer. Greenhouse heaters and evaporative coolers were programmed to maintain 25oC temperature. Plants were watered daily by hand, and fertilized once weekly with Miracle‐Gro Water Soluble Azalea Camellia Rhododendron fertilizer (30‐10‐10) supplying 26 mg N plant‐1 wk‐1.

Mini‐lysimeters (Rhizon Soil Moisture Samplers, Eijkelkamp Agrisearch Equipment, The Netherlands) were installed in four of the six plants at each gypsum level . The mini‐lysimeter tubes were constructed of hydrophyllic, wire‐reinforced materials with 0.15 micron pores. They were 2.5 mm diameter x 10 cm length. Soil solution samples were collected on 10‐Mar, 23‐Mar, 5‐Apr, 16‐May, and 11‐Jun and analyzed for cation and trace element concentration by ICP spectroscopy.

Plants were destructively harvested after 26 weeks, and all soil and potting media was washed from the roots. Root infection was estimated by placing ten randomly sampled 1‐cm root sections on PARPH selective media and incubated with daily inspection at 20oC for 10 days. Shoot and root components were dried and weighed separately to determine biomass.

Statistical Analysis Gypsum and organic matter experiments Shoot and root biomass were modeled using Proc Mixed in SAS 9.2. The fixed effect model for the Spring 2011, Fall 2011, and Spring 2013 experiments with two‐way interactions included terms for Infection, Gypsum, Organic amendment type, Infection x Gypsum, Infection x Organic

45 amendment type, and Gypsum x Organic amendment type. Replication and the Replication x Infection interaction were included as random effects for the Spring 2011 and Fall 2011 split plot RBD designs, and only replication was included as a random effect for the Spring 2013 CRD design. Pairwise comparisons for main and interaction effects were performed by the Fisher’s LSD method, and SAS Macro PDMIX800 was used to convert multiple comparisons into letter code format.

Root infection incidence and pathogen propagule density in soil were modeled using Proc Mixed in SAS 9.2. Data were only collected from the plants deliberately inoculated with P. cinnamomi, and the analysis was not performed across infection level. The model included terms Gypsum, Organic amendment type, and the Gypsum x Organic amendment type interaction. Replication was included as a random effect. Pairwise comparisons for main and interaction effects were performed by Fisher’s LSD method, and SAS Macro PDMIX800 was used to convert multiple comparisons into letter code format.

Mean comparison of the full factorial Infection x Gypsum x Organic amendment with the fungicide controls was accomplished using Proc Mixed in SAS 9.2. A simplified model was created including only replication and treatment. Treatment consisted of all possible combinations of infection, gypsum, organic type, and the fungicide controls. Pairwise comparisons were performed using a Bonferroni correction to adjust for large number of treatments, and pairwise comparisons were converted into letter code format using the SAS Macro PDMIX800.

Gypsum rate experiment Soluble calcium in soil solution was modeled using a logistic function selected with nonlinear regression in StatGraphics Centurion version 16.1.03 (StatPoint Technologies, 2010). Calcium was measured at five dates, and the average of all five dates was used to compute the regression. Initial estimates were 450 for the plateau parameter, and 0.1 for the intercept and exponential increase factors. Parameter estimates converged after 10 iterations. The logistic model approximates two straight lines representing the response phase and the plateau phase, which comprise a linear‐plateau mode (Wilcutts et al., 1998).

The straight line of the plateau phase can be approximated by [Ca2+] =A. The slope of the linear response phase can be approximated by the derivative of the logistic model at [Ca2+] =A/2, and the Gyp Rate = 0 intercept is calculated using a point‐slope method. The intercept between the linear response phase and linear plateau phase represents the point of Ca2+ saturation in soil solution (Cerrato and Glackmer, 1990; Willcutts et al., 1998).

Logistic model: [Ca2+] = A/(1+(e(b‐(c*Gyp Rate)))) A (Plateau factor): 478.53 b (Intercept factor): 2.281346 c (Exponential increase factor): 0.631146 [Ca2+]: mg Ca2+ / L soil solution Gyp Rate: cmol (+ )/ kg soil

[Ca2+] = 478.513 / (1+(e(2.281346‐ (0.631146* Gyp_Rate))))

Linear Plateau Model: For Gyp Rate < 15.6: Ca = (Gyp Rate * C) + B For Gyp Rate > 15.6: Ca = A A (Plateau): 478.53 B (Intercept of linear increase phase): 0.0835 C (Slope of linear increase phase): 0.1395833

Results Organic and Gypsum Amendments Experiment, Spring ‐ Summer 2011 Inoculation with P. cinnamomi had a large impact on plant biomass, reducing shoot biomass by 32% and root biomass by 42% compared to the non‐infected plants across all treatments. Shoot biomass accumulation was not different between the organic amendment treatments, nor did infected plants respond differently than non‐infected plants to organic amendment (Table 1). Root biomass of plants not infected with P. cinnamomi was higher with sawdust than the other

47 organic amendments, however the increase in biomass did not occur on the plants infected with P. cinnamomi (Table 1). There was no difference in root infection incidence, or P. cinnamomi propagule density in soil between the organic amendments (Table 2).

The main effect of gypsum amendment, across all organic amendments, significantly increased shoot biomass by 46%, and root mass by 49% of plants infected with P. cinnamomi (Table 1). However, there was no significant difference in root or shoot biomass between gypsum levels for the non‐infected plants indicating the high level of gypsum application did not negatively impact biomass (Table 1). Root infection incidence of plants without gypsum was 45.6%, and significantly decreased to 26.4% with the addition of gypsum (Table 2). Pathogen propagule density in soil at the end of the trial was not different between the gypsum amended soil, and the soil without gypsum (Table 2).

Gypsum application increased Mehlich III extractable calcium in soil after 14 weeks 1300 mg kg‐1 in the pots without gypsum, to 10,350 mg kg‐1 with gypsum (Table 3). Plant uptake of Ca2+ by infected plants increased from 14.8 mg plant‐1 to 32.2 mg plant‐1 with gypsum addition, and Ca2+ uptake by non‐infected plants increased from 30.6 mg plant‐1 to 43.8 mg plant‐1 (Table 4). Tissue Ca2+ concentration of infected plants increased from 2600 mg Kg‐1 to 3885 mg Kg‐1 with gypsum addition, and from 2920 mg Kg‐1 to 4403 mg Kg‐1 in the non‐infected plants (Appendix 1). Tissue concentration of K+ and Mg2+ were not influenced by gypsum incorporation (Appendix 1). Total uptake of N, P, S, Ca, Mg, and K by plants infected with P. cinnamomi significantly increased with gypsum application, due to increased total biomass (Table 4).

Organic Matter and Gypsum Experiment Winter 2011‐2012 The spring‐summer experiment was repeated during winter 2011‐2012, however statistical differences were incongruent with the spring‐summer 2011 experiment. Total biomass accumulation of plants was much lower, and the effect of infection was much greater. Infected plants only accumulated marginal growth, and gypsum did not significantly reduce disease incidence or biomass accumulation. (Table 5; 6)

In contrast to the spring‐summer experiment, the main effect of organic type on shoot biomass was significant across levels of infection and gypsum. Plants amended with peat had the highest shoot biomass, and plants amended with sawdust had the lowest shoot biomass. Organic amendment type had no significant effect on root biomass (Table 5). Root infection incidence was higher for the dairy solids, sawdust, and yard debris compost than the non‐amended control (Table 6).

Gypsum had no significant effect on shoot or root biomass of infected or non‐infected plants, indicating no disease suppression on infected plants, and no effect of salt toxicity on non‐ infected plants (Table 5). The gypsum x infection interaction term was non‐significant, indicating the effect of gypsum on biomass was not different between infected and non‐infected plants. Root infection incidence was not different between the gypsum and non‐gypsum amended plants across all levels of organic amendment (Table 6). P. cinnamomi propagule density at the end of the experiment did not differ between levels of gypsum, organic amendment, or the gypsum x organic type interaction (Table 6).

Organic Matter and Gypsum Trial Summer 2013 The main effects of gypsum and one compost source were evaluated during summer 2013. Biomass differences between the infected plants appeared visually evident, and the trial was harvested after nine weeks, compared to 14 weeks for the summer and winter 2011 trials. After plants were destructively harvested, no significant treatment effect of disease suppression was revealed from either gypsum or organic amendment on shoot or root biomass (Table 7). There was a gypsum x organic matter interaction for root infection incidence, where the yard debris compost reduced infection incidence only in the absence of gypsum (Table 8). Pathogen propagule density at the end of the trial was not assessed.

Rates of gypsum required for suppression Soluble calcium in soil solution followed a linear increase with gypsum addition up to 15.6 cmol (+) / kg soil. Beyond 15.6 cmol (+)/ kg of gypsum added, calcium in soil solution did not increase further due to solubility limits. Soluble calcium in soil solution at saturation was 453.62 mg / L soil solution (Figure 1).

Root infection incidence decreased with the addition of gypsum. Gypsum amendment of 6 cmol (+) / 100 g soil was not significantly different than no gypsum amendment, however root infection at gypsum amendment of 12 cmol (+) / kg was significantly lower. Increasing gypsum beyond 12 cmol (+) / kg did not cause further decrease in root infection incidence (Figure 2).

Despite differences in root infection incidence, the differences in plant biomass accumulation were less apparent. There was no significant effect on root biomass across all levels of gypsum amendment (Figure 4). Shoot biomass tended to increase with higher gypsum application rates, however the response was variable (Figure 3).

Discussion Three experiments were performed evaluating organic and gypsum amendment for suppressing P. cinnamomi root rot disease of blueberry. In the spring 2011 experiment, gypsum decreased root infection incidence and enhanced root and shoot biomass. However, in the fall 2011 and spring 2013 experiments, gypsum failed to reduce root infection incidence or enhance biomass of infected plants. One experiment evaluated growth response and root infection incidence of P. cinnamomi inoculated plants with varying rates of gypsum. This experiment determined the minimum gypsum rate to maximize calcium solubility, showed decreased root infection and increased shoot biomass with calcium saturation, however root biomass was not influenced.

If gypsum does have a biologically significant effect on blueberry root rot disease, the effect may vary according to disease pressure, plant vigor, or may be manifested over a prolonged timeframe than the greenhouse experiments revealed. Despite the somewhat inconsistent results of infected plant response to gypsum, we consistently demonstrated that a very high rate of gypsum (5% v/v) did not appear to cause salt injury to either the infected or non‐infected plants.

The proposed mechanism of P. cinnamomi disease suppression by gypsum is zoospore encystment and subsequent germination in response to Ca2+ concentration in soil solution (Byrt et al., 1982). In pure solution studies, Ca2+ concentration of 20 cmol (+) / L induced P. parasitica

50 zoospore encystment inhibiting motility (von Broembsen and Deacon, 1997). Calcium concentration in solution of 1600 mg L‐1 significantly decreased zoospore release of P. sojae, and release was further inhibited by Ca concentration 3200 mg L‐1 (Sugimoto et al., 2008). However, translation to field gypsum application rates is difficult due to Ca2+ solubility interactions with

2‐ 3‐ 2‐ ‐ ‐ CO3 , PO4 , SO4 , NO3 , Cl , and the soil cation exchange. Most fruit crop research using gypsum for suppressing P. cinnamomi on avocado and raspberry has investigated single, rates of gypsum (Maloney et al., 2005; Messenger 2000a; 2000b; Pinkerton et al., 2009).

In three of the experiments, a single high rate (5% v/v) of gypsum was added, which is cost‐ prohibitive for blueberry growers on a whole‐field scale. The lowest rate of gypsum to maximize soluble Ca2+ was modeled using nonlinear regression with a logistically derived linear plateau model to determine when increasing gypsum application rate did not further increase Ca2+ in soil solution. The intercept of the linear increase phase and linear plateau phase occurred after 15.6 cmol (+) / kg gypsum amendment.

Maximum calcium solubility, and the gypsum rate required to maximize soluble Ca2+ likely vary between soils with different pH, CEC, P levels, base saturation, and anion speciation. The CEC of the native Malabon silty clay loam (fine, mixed, superactive, mesic Pachic Ultic Argixeroll) soil was 24.4 cmol (+ )/ kg / kg, pH was 5.3, Ca was 10.3 cmol (+ )/ kg, Mg was 5.9 cmol (+ )/kg, K was

‐ 0.57 cmol (+ )/kg, total bases were 16.8 cmol (+ )/ kg, Bray P was 23 mg / kg, and NO3 was 0.5 mg / kg.

Organic amendment did not significantly reduce disease severity or incidence in the three experiments testing various composts and organic residues for disease suppression. Other research in field settings have correlated organic residues with P. cinnamomi suppression due to increased activity of β‐1,3, 1,4, and 1,6 glucanase producing microbial populations antagonistic to Phytophthora hyphae largely composed of β‐1,3, β‐1,4 β‐1,6 glucan linkages (Downer et al., 2001; Hegnauer and Hohl, 1978; Richter et al., 2011). Lack of suppression in our greenhouse experiments may have been caused by high inoculum level, conditions conducive to infection, or high host plant sensitivity. However, suppression of soilborne diseases with organic amendment

51 appears fundamentally variable and inconsistent (Hoitink and Fahy, 1986; Maloney et al., 2005; Scheuerell et al., 2005)

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Figures and Tables

Table 1. Main and interaction effects for aerial and root biomass, and all treatment mean separation. Spring‐Summer 2011 greenhouse experiment. Vegetative Biomass (g) Root Biomass (g) Main Effects Organic Matter Gypsum Infection z z Non‐Infected 10.2 ± 0.39 A 3.4 ± 0.16 A Infected 7.0 ± 0.39 B 2.0 ± 0.16 B

z z Y9.1± 0.36 A 2.9 ± 0.16 A N8.1± 0.36 B 2.5 ± 0.16 B

y z z BSB 9.0 ± 0.53 AB 2.9 ± 0.23 AB DS 8.6 ± 0.53 AB 2.6 ± 0.23 B None 8.4 ± 0.53 AB 2.5 ± 0.23 B Pt 9.4 ± 0.53 A 3.2 ± 0.23 A SD 8.1 ± 0.53 B 2.4 ± 0.23 B YDC 8.3 ± 0.53 AB 2.5 ± 0.23 B

Gypsum X Infection Interaction z z NNon‐Infected 10.5 ± 0.48 A 3.5 ± 0.20 A YNon‐Infected 9.9 ± 0.48 A 3.3 ± 0.20 A YInfected8.3± 0.48 B 2.4 ± 0.20 B NInfected5.7± 0.48 C 1.6 ± 0.20 C

Model Parameter p‐value p‐value Infection 0.0012 0.0002 Org_Type 0.3478 0.0366 Gyp 0.0073 0.0033 Gyp*Infection <0.0001 0.0552 Org_Type*Infection 0.3534 0.0003 Org_Type*Gyp 0.5264 0.0876

All Treatment Combinations Vegetative Biomass (g) Root Biomass (g) Organic Type Gypsum Infected Non‐Infected Infected Non‐Infected y x x NA N6.0± 1.02 BCDE 8.8 ± 1.02 ABCDE 1.9 ± 0.39 CDE 2.6 ± 0.39 BCDE Pt N 7.0 ± 1.02 ABCDE 12.2 ± 1.02 A 1.7 ± 0.39 CDE 3.0 ± 0.39 BCDE BSB N 5.1 ± 1.02 DE 11.0 ± 1.02 ABC 1.6 ± 0.39 DE 4.0 ± 0.39 AB DS N 4.8 ± 1.02 E 11.4 ± 1.02 AB 1.3 ± 0.39 E 3.0 ± 0.39 BCDE SD N 5.4 ± 1.02 DE 9.7 ± 1.02 ABCDE 1.5 ± 0.39 E 5.2 ± 0.39 A YDC N 5.8 ± 1.02 CDE 10.0 ± 1.02 ABCDE 1.4 ± 0.39 E 2.2 ± 0.39 BCDE None Y 9.4 ± 1.02 ABCDE 9.4 ± 1.02 ABCDE 2.9 ± 0.39 BCDE 2.9 ± 0.39 BCDE PT Y 8.4 ± 1.02 ABCDE 10.2 ± 1.02 ABCDE 2.5 ± 0.39 BCDE 2.9 ± 0.39 BCDE BSB Y 9.5 ± 1.02 ABCDE 10.4 ± 1.02 ABCD 2.5 ± 0.39 BCDE 3.6 ± 0.39 ABCD DS Y 8.7 ± 1.02 ABCDE 9.3 ± 1.02 ABCDE 2.3 ± 0.39 BCDE 3.7 ± 0.39 ABC SD Y 7.7 ± 1.02 ABCDE 9.5 ± 1.02 ABCDE 2.0 ± 0.39 BCDE 4.0 ± 0.39 AB YDC Y 6.3 ± 1.02 BCDE 10.8 ± 1.02 ABC 1.9 ± 0.39 CDE 4.0 ± 0.39 AB Fungicide Control ‐ 12.0 ± 1.02 A 11.4 ± 1.02 AB 2.9 ± 0.39 BCDE 2.4 ± 0.39 BCDE

p‐value <0.0001 <0.0001 z Letters indicate comparisons of main and interaction effects y NA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost; Gyp=gypsum. x Letters indicate Bonferroni adjusted pairwise comparisons for all infection, gypsum, and oragnic amendment treatment combinations, and the fungicide controls.

Table 2. Root infection incidence and P. cinnamomi propagule density in soil after 14 wks. Spring‐Summer 2011 greenhouse experiment. % Root Infection P. cinn. propagules / g soil Main Effects Gypsum z z N 45.6% ± 3.3% A 0.86 ± 0.24 A Y 26.4% ± 2.9% B 0.57 ± 0.21 A

Organic Type y z z BSB 32.5% ± 5.4% A 0.86 ± 0.42 A DS 32.5% ± 7.1% A 0.49 ± 0.38 A NA 29.2% ± 5.8% A 1.10 ± 0.45 A Peat 42.5% ± 6.2% A 1.10 ± 0.48 A SD 40.8% ± 4.0% A 0.74 ± 0.42 A YDC 38.3% ± 7.4% A 0.00 ± 0.00 A

Model Parameter p‐value p‐value Gypsum <0.0001 0.3725 Organic 0.3652 0.3381 Gypsum * Organic 0.4727 0.2377

All Treatment Combinations Organic Type Gypsum % Root Infection P. cinn. propagules / g soil y x x NA N 37.7% AB 1.72 A PT N 53.6% A 1.23 A BSB N 45.0% AB 0.74 A DS N 49.2% A 0.00 A SD N 41.5% AB 1.47 A YDC N 50.0% A 0.00 A NA Y 14.0% AB 0.49 A PT Y 27.3% AB 0.98 A BSB Y 13.7% AB 0.98 A DS Y 11.6% AB 0.98 A SD Y 39.5% AB 0.00 A YDC Y 29.6% AB 0.00 A Fungicide Control ‐ 5.1% B 1.23 A

p‐value 0.0006 0.3406 z Letters indicate comparisons of main effects y NA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost; Gyp=gypsum. x Letters indicate Bonferroni adjusted pairwise comparisons for all gypsum, and oragnic amendment treatment combinations, and the fungicide controls.

Table 3. Soil chemical properties of soil‐organic matter‐gypsum blends after 14‐wk. Trial 1. Treatment pH OM Bray I P S Ca Mg K Gypsum Residue 1:1 LOIy ‐‐‐‐‐‐‐‐‐‐Mehlich III extractable‐‐‐‐‐‐‐‐‐‐ mg kg‐1 5% v/v 20% v/v water g 100g‐1 mg kg‐1 mg kg‐1 mg kg‐1 mg kg‐1 N NAz 5.27 2.7 123 30 1313 225 474 N PT 5.47 5.0 129 19 1267 242 468 N BS 5.60 6.3 259 24 1388 260 441 N DS 5.67 4.0 134 19 1277 267 588 N SD 5.83 4.4 85 12 1124 222 480 N YDC 6.00 5.7 123 23 1452 285 657 Y NA 5.47 2.7 121 5002 9527 153 459 Y PT 5.20 4.7 171 5507 10727 130 379 Y BSB 5.43 6.2 262 5334 10492 138 373 Y DS 5.50 4.3 146 5371 10548 161 470 Y SD 5.67 4.5 107 5060 9964 168 421 Y YDC 5.87 5.3 160 5373 10833 220 549 N Fungicide Control 5.03 2.8 210 200 1924 237 474 zNA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost yLoss on ignition

Table 4. Total nutrient uptake of ‘Draper’ highbush blueberry grown in different gypsum and organic matter amendments, either infected or non‐infected with P. cinnamomi root rot. Trial 1. N P S Ca Mg K

P. cinn. Infection mg mg mg mg mg mg Y N Y N Y N Y N Y N Y N Treatment NAz 70 78 6.7 7.5 5.6 6.6 15 23 4.2 5.8 28 38 PT 74 107 7.5 11.2 6.1 9.1 19 33 5.5 9.4 32 52 BSB 58 89 6.1 11.5 4.8 8.8 13 35 3.7 9.6 21 45 DS 56 96 5.6 10.8 4.6 8.2 12 30 3.3 8.4 23 50 SD 57 68 5.9 9.3 5.1 6.3 13 34 3.7 9.4 22 39 YDC 59 87 6.3 9.4 5.1 7.5 13 25 3.8 6.9 27 46 Gyp 83 87 8.6 8.2 16.3 15.1 38 37 7.8 7.6 41 42 Gyp + PT 80 93 8.2 9.9 12.7 16.1 32 46 6.3 7.8 38 47 Gyp + BSB 81 88 9.2 10.2 15.4 18.1 37 47 7.8 8.9 42 48 Gyp + DS 82 78 8.9 8.8 11.5 15.1 31 41 6.1 6.9 39 43 Gyp + SD 72 60 7.9 8.6 13.2 14.2 29 42 6.3 7.2 37 44 Gyp + YDC 63 92 6.8 9.0 10.2 19.0 24 47 5.1 8.6 31 53 Fungicide 109 115 12.5 11.8 10.3 8.9 34 34 10.2 8.5 50 49

Main Effects: Gypsum 0.3910 0.2935 0.0009 0.0855 <0.0001 <0.0001 <0.0001 <0.0001 <0.0001 0.3829 <0.0001 0.7879 Organic 0.0099 0.0036 0.7120 0.0119 0.2451 0.2114 0.1367 0.1078 0.2339 0.1292 0.5687 0.2459 Gyp*Org 0.7690 0.4676 0.5281 0.6984 0.3512 0.6570 0.3464 0.5606 0.3170 0.1016 0.3183 0.4873 zNA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost; Gyp=gypsum.

Table 5. Main and interaction effects for aerial and root biomass, and all treatment mean separation. Fall‐Winter 2011 greenhouse experiment. Aerial Mass (g) Root Mass (g) Main Effects Organic Matter Gypsum Infection z z Non‐Infected 4.3 ± 0.18 A 1.7 ± 0.05 A Infected 2.0 ± 0.18 B 0.7 ± 0.05 B

z z Y3.1± 0.14 A 1.2 ± 0.04 A N3.1± 0.14 A 1.2 ± 0.04 A

y z z BSB 3.2 ± 0.18 AB 1.1 ± 0.06 A DS 3.1 ± 0.18 AB 1.2 ± 0.06 A None 2.9 ± 0.18 B 1.2 ± 0.06 A Pt 3.5 ± 0.18 A 1.2 ± 0.06 A SD 2.8 ± 0.18 B 1.3 ± 0.06 A YDC 3.2 ± 0.18 AB 1.2 ± 0.06 A

Gypsum X Infection Interaction z z NNon‐Infected 4.36 ± 0.20 A 1.737 ± 0.05 A YNon‐Infected 4.22 ± 0.20 A 1.693 ± 0.05 A YInfected2.06± 0.20 B 0.709 ± 0.05 B NInfected1.88± 0.20 B 0.628 ± 0.05 B

Effect Infection 85.6 0.0002 369.4 <0.0001 Org_Type 2.34 0.0456 0.6 0.6751 Gyp 0.03 0.8600 2.7 0.1060 Gyp*Infection 1.79 0.1835 0.2 0.6270 Org_Type*Infection 1.65 0.1534 0.5 0.8084 Org_Type*Gyp 0.78 0.5675 3.7 0.0041

All Treatment Combinations Vegetative Biomass (g) Root Biomass (g) Organic Type Gypsum Infected Non‐Infected Infected Non‐Infected y x x NA N2.0± 0.33 D 4.3 ± 0.33 AB 0.7 ± 0.10 E 1.8 ± 0.10 A Pt N 2.0 ± 0.33 CD 5.1 ± 0.33 A 0.8 ± 0.10 DE 1.8 ± 0.10 A BSB N 1.8 ± 0.33 D 4.4 ± 0.33 AB 0.5 ± 0.10 E 1.4 ± 0.10 ABC DS N 1.9 ± 0.33 D 4.1 ± 0.33 AB 0.6 ± 0.10 E 1.7 ± 0.10 AB SD N 1.8 ± 0.33 D 3.7 ± 0.33 ABC 0.6 ± 0.10 E 1.8 ± 0.10 A YDC N 1.7 ± 0.33 D 4.5 ± 0.33 AB 0.6 ± 0.10 E 1.6 ± 0.10 AB None Y 2.0 ± 0.33 D 3.5 ± 0.33 ABCD 0.7 ± 0.10 DE 1.7 ± 0.10 AB PT Y 2.2 ± 0.33 CD 4.6 ± 0.33 AB 0.7 ± 0.10 E 1.5 ± 0.10 ABC BSB Y 2.2 ± 0.33 CD 4.5 ± 0.33 AB 0.7 ± 0.10 DE 1.9 ± 0.10 A DS Y 1.8 ± 0.33 D 4.5 ± 0.33 AB 0.6 ± 0.10 E 1.7 ± 0.10 AB SD Y 2.1 ± 0.33 CD 3.7 ± 0.33 ABC 0.9 ± 0.10 DE 1.7 ± 0.10 AB YDC Y 2.2 ± 0.33 CD 4.5 ± 0.33 AB 0.6 ± 0.10 E 1.9 ± 0.10 A Fungicide Control ‐ 2.1 ± 0.33 CD 3.0 ± 0.33 BCD 1.0 ± 0.10 CDE 1.2 ± 0.10 BCD

Table 6. Root infection incidence and P. cinnamomi propagule density in soil after 14 wks. Fall‐ Winter 2011 greenhouse experiment. % Root Infection Propagules / g soil Main Effects Gypsum z z N 56.7% ± 3.6% A 1.66 ± 0.38 A Y 52.8% ± 3.4% A 2.60 ± 0.42 A

Organic Type y z z BSB 46.7% ± 7.1% BC 2.32 ± 0.46 A DS 57.5% ± 4.8% AB 1.47 ± 0.79 A NA 40.8% ± 5.6% C 1.33 ± 0.60 A Peat 53.3% ± 6.6% ABC 2.37 ± 0.83 A SD 66.7% ± 5.0% A 3.37 ± 0.71 A YDC 63.3% ± 4.5% A 1.90 ± 0.77 A

Model Parameter p‐value p‐value Gypsum 0.3590 0.0869 Organic 0.0134 0.2870 Gypsum * Organic 0.4325 0.3298

All Treatment Combinations Organic Type Gypsum % Root Infection P. cinn. propagules / g soil y x x NA N 39.5% A 1.0 A PT N 55.8% A2.6A BSB N 48.5% A2.3A DS N 67.8% A0.3A SD N 73.6% A1.9A YDC N 57.1% A1.9A NA Y 41.1% A1.7A PT Y 51.0% A2.2A BSB Y 44.0% A2.3A DS Y 48.3% A2.7A SD Y 61.2% A4.9A YDC Y 70.6% A1.9A Fungicide Control ‐ 0.0% B1.0A

p‐value <0.0001 0.1614 z Letters indicate comparisons of main effects y NA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; x Letters indicate Bonferroni adjusted pairwise comparisons for all gypsum, and oragnic amendment treatment combinations, and the fungicide controls.

Table 7. Main and interaction effects for aerial and root biomass, and all treatment mean separation. Spring 2013 gypsum and organic matter greenhouse experiment. Vegetative Biomass (g) Root Biomass (g) Main Effects Organic Matter Gypsum Infection z z Non‐Infected 5.4 ± 0.19 A 2.5 ± 0.11 A Infected 3.0 ± 0.19 B 1.2 ± 0.11 B

z z N4.2± 0.19 A 1.9 ± 0.11 A Y4.1± 0.19 A 1.7 ± 0.11 A

y z z NA 4.3 ± 0.19 A 1.8 ± 0.11 A YDC 4.1 ± 0.19 A 1.8 ± 0.11 A

Interaction Effect z z NNon‐Infected 5.6 ± 0.27 A 2.6 ± 0.15 A YNon‐Infected 5.2 ± 0.27 A 2.4 ± 0.15 A YInfected3.1± 0.27 B 1.3 ± 0.15 B NInfected2.9± 0.27 B 1.0 ± 0.15 B

Effect p‐value p‐value Infection <0.0001 <0.0001 Org 0.4992 0.9068 Gyp 0.7489 0.1474 Gyp*Infection 0.2607 0.8638 Org*Infection 0.2088 0.0466 Org*Gyp 0.7419 0.7175

All Treatment Combinations Vegetative Biomass (g) Root Biomass (g) Organic Type Gypsum Infected Non‐Infected Infected Non‐Infected y x x NA N3.3± 0.37 B 5.5 ± 0.37 A 1.1 ± 0.21 C 2.4 ± 0.21 AB YDC N 2.5 ± 0.37 B 5.7 ± 0.37 A 1.0 ± 0.21 C 2.4 ± 0.21 AB y NA Y3.2± 0.37 B 5.2 ± 0.37 A 1.5 ± 0.21 BC 2.3 ± 0.21 AB YDC Y 3.0 ± 0.37 B 5.3 ± 0.37 A 1.0 ± 0.21 C 2.9 ± 0.21 A Fungicide Control 2.6 ± 0.37 B 2.3 ± 0.37 B 1.3 ± 0.21 C 1.2 ± 0.21 C

p‐value <0.0001 <0.0001 z Letters indicate comparisons of main and interaction effects y NA=no amendment; YDC=yard debris compost x Letters indicate Bonferroni adjusted pairwise comparisons for all infection, gypsum, and oragnic amendment treatment combinations, and the fungicide controls.

Table 8. P. cinnamomi root infection incidence after 9 wks. Spring 2013 greenhouse experiment. % Root Infection Main Effects Gypsum z N 61.7% ± 3.9% A Y 55.8% ± 7.8% A

Organic Type z None 54.2% ± 6.5% A YDC 63.3% ± 5.7% A

Interaction Effect Gypsum Organic z NN66.7% ± 8.2% A NYDC41.7% ± 25.6% B YN56.7% ± 16.3% AB YYDC70.0% ± 21.9% A

Model Parameter p‐value Gypsum 0.4345 Organic 0.2021 Gypsum * Organic 0.0283

All Treatment Combinations Organic Type Gypsum y x NA N 66.9% A YDC N 57.1% A None Y 36.7% A YDC Y 71.6% A Fungicide Control 0.3% B p‐value <0.0001 z Letters indicate comparisons of main and interaction effects y NA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost; Gyp=gypsum. x Letters indicate Bonferroni adjusted pairwise comparisons for all gypsum, and oragnic amendment treatment combinations, and the fungicide controls.

Table 9. Chemical analysis of saturated deionized water extract analysis of organic residues used in the spring and fall 2011 experiments evaluating organic amendments and gypsum for blueberry root rot disease suppression.

Organic pH EC NO3‐N NH4‐N P SO4‐S Ca Mg K Na Experiment Amendment 1:1 ms/cm mg L‐1 mg L‐1 mg L‐1 mg L‐1 mg L‐1 mg L‐1 mg L‐1 mg L‐1 Sp11 BSBz 6.5 2.8 57 5 6 998 295 142 51 101 Sp11 DS 7.7 2.8 7 < 0.5 37 76 51 27 655 89 Sp11 YDC 7.2 3.6 3 20 6 164 63 68 759 64 Sp11, F11 PT 4.8 0.7 1 9 15 216 84 34 14 13 Sp11, F11 SD 7.0 0.1 < 0.5 3 0 6 9 2 10 3 F11 BSB 6.8 2.2 96 61 10 491 91 53 77 58 F11 DS 8.2 2.8 1 1 32 206 57 28 633 138 F11 YDC 7.4 2.8 < 0.5 < 0.5 10 118 66 49 620 53 zNA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost.

Table 10. Total nutrient analysis of organic amendments used in the spring and fall 2011 experiments evaluating organic amendments and gypsum for blueberry root rot disease suppression. Nutrient concentrations are reported on a dry‐weight basis. Organic Experiment N P K Ca Mg Na S B Mn Amendment % % % % % % % ppm ppm Sp11 BSBz 0.93 0.59 0.14 1.08 0.37 0.07 0.24 7.9 373 Sp11 DS 2.45 0.37 1.25 1.12 0.38 0.14 0.30 20.5 267 Sp11 YDC 1.63 0.24 0.89 1.48 0.35 0.05 0.17 21.3 537 Sp11, F11 PT 1.29 0.05 0.07 0.98 0.19 0.01 0.29 7.6 235 Sp11, F11 SD 0.08 bdly 0.01 0.04 bdl bdl 0.01 2.0 26 F11 BSB 1.12 0.77 0.15 1.19 0.41 0.06 0.32 7.5 476 F11 DS 1.47 0.32 1.13 1.06 0.34 0.22 0.23 23.8 227 F11 YDC 1.12 0.21 0.74 1.17 0.37 0.06 0.14 14.7 494 zNA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost. yBelow detection limit.

600

500

400 Solution

Soil 300 L

200 Logistic model Ca = 453.622/ (1+(e(1.85347 ‐ (0.246* Gyp Rate)))) r2=0.79 Ca2+

For Gyp Rate < 15.65: Ca = Gyp Rate * 27.9166 + 16.700 r2=0.83 mg 100 Linear plateau model For Gyp Rate > 15.65: Ca = 453.622 0 0 20406080100 Gypsum Rate cmol + / kg soil added as Ca2SO4*2H2O

Figure 1. Effect of increasing gypsum amendment rate on soluble Ca2+ in soil solution sampled with Rhizon tube mini‐lysimeters. One cmol (+) / kg soil of gypsum incorporation is equal to 0.95 g gypsum / kg soil. Cation exchange capacity of the native soil was 24.4 cmol (+)/ kg, and total exchangeable cations (Ca, Mg, K, Na) was 16.8 cmol (+)/ kg soil.

70% a 60% a

50% Incidence

40%

30% b

Infection b b

20% b

Root b 10% b b b

0% 0 20406080100 Gypsum Rate (meq / 100 g)

Figure 2. Effect of gypsum rate on P. cinnamomi root infection incidence. Letters indicate significant differences between treatments via Fisher’s LSD test. One cmol (+) / kg soil of gypsum incorporation is equal to 0.95 g gypsum / kg soil. Maximum Ca2+ in soil solution occurred when gypsum was added at rates greater than 15.6 cmol (+) / kg soil. Cation exchange capacity of the native soil was 24.4 cmol (+)/ kg, and total exchangeable cations (Ca, Mg, K, Na) was 16.8 cmol (+)/ kg soil.

9 8 7 (g)

6 c 5 bc bc bc abc abc Biomass

4 3 abc Shoot ab 2 a ab 1 0 0 20406080100 Gypsum Rate (cmol (+) /kg)

Figure 3. Effect of gypsum rate on shoot biomass of plants infected with P. cinnamomi . Letters indicate significant differences between treatments via Fisher’s LSD test. One cmol (+) / kg soil of gypsum incorporation is equal to 0.95 g gypsum / kg soil. Maximum Ca2+ in soil solution occurred when gypsum was added at rates greater than 15.6 cmol (+) / kg soil. Cation exchange capacity of the native soil was 24.4 cmol (+)/ kg, and total exchangeable cations (Ca, Mg, K, Na) was 16.8 cmol (+)/ kg soil.

10 9 8 7 (g) 6 5 Biomass 4

Root 3 2 1 0 0 20406080100 Gypsum Rate (cmol (+) /kg)

Figure 4. Effect of gypsum rate on root biomass of plants infected with P. cinnamomi. Treatment means were not significantly different at P =0.05. Error bars show standard error of the mean. Gypsum rate experiment.

Chapter Four: Influence of gypsum, mulch, and drip irrigation line placement on Phytophthora cinnamomi root rot disease during field establishment of highbush blueberry

John R. Yeo, J. Weiland, D.M. Sullivan, and D.R. Bryla

Abstract: Phytophthora cinnamomi causes root rot disease of highbush blueberry, which decreases plant growth, yield, and profitability for producers. Fungicides are available for suppression of the disease, but are subject to pathogen resistance, and excluded from use in certified organic production systems. Alternative, non‐chemical management strategies to reduce Phytophthora root rot were evaluated in a new field transplanted with a highly susceptible blueberry cultivar, ‘Draper’. The field was deliberately infested with P. cinnamomi prior to transplanting. Treatment combinations included conventional sawdust mulch or weed mat, narrow or wide drip irrigation line placement, and no soil amendment or pre‐plant incorporation of gypsum. Two conventional fungicide treatments, mefenoxam and fosetyl‐Al, were also included with a narrow drip line placement. Over multiple sampling dates, root infection incidence was lower with sawdust mulch than with weed mat, with irrigation lines placed 20 cm from the crown than adjacent to the crown, and with pre‐plant gypsum incorporation compared to no gypsum amendment. Soil under weed mat accumulated more heat units than under sawdust, and P. cinnamomi in buried petri plate incubations grew more hyphal area under weed mat. However, after two growing seasons, plant growth was similar between the two mulch types. The effects of irrigation line placement and gypsum incorporation, on the other hand, were interactive, and the combination of widely‐spaced irrigation lines with gypsum incorporation produced the greatest plant biomass of the cultural treatments. Irrigation lines placed adjacent to the plant crown negated the disease‐suppressive effects of gypsum by moving the zoospore‐inhibiting Ca2+ from the salt away from the plant root zone. Additionally, plots with irrigation lines adjacent to the plant crown were wetter in the center of the bed, facilitating root infection by the zoospores. Despite enhanced plant biomass with gypsum amendment and widely‐spaced irrigation lines, plants treated with conventional fungicides had at least twice the biomass of plants grown in any of the treatments with no fungicide. In addition, the use of widely‐ spaced irrigation lines resulted in N deficiency during the first year after planting because the ammonium sulfate fertilizer supplied via fertigation was applied too far from the roots of the young plants. Thus, while less effective than the fungicides, the study suggests that pre‐plant soil amendment with gypsum coupled with widely‐spaced drip irrigation lines can help suppress phytophthora root rot and increase growth and survival of blueberry at sites where the pathogen is present, particularly if N is managed to avoid plant deficiency.

Introduction: Phytophthora cinnamomi is a soilborne pathogen of >900 plant species, and is a serious impediment to commercial production of susceptible hosts. Phytophthora species worldwide contribute major economic losses to almost all food crops, and necessitate large fungicide inputs to the environment. On blueberry, P. cinnamomi causes root rot disease, and has been detected in 23% of fields comprising 7800 acres in Oregon (Bryla et al., 2008). Root rot disease causes economic loss in most blueberry producing regions worldwide, with United States acreage totaling 77,700 acres in 2012 and rapidly expanding (USDA ERS, 2013).

Blueberry root rot disease is more severe when plants are irrigated by drip than by overhead sprinklers (Bryla and Linderman, 2007). However, when Phytophthora is not present, drip irrigation offers advantages over sprinklers including greater food safety, more efficient fertilizer delivery, higher water use efficiency, and enhanced plant growth (Bryla et al., 2011; Bryla and Machado, 2011). Highbush blueberry is sensitive to salinity accumulation in soil, and irrigation lines placed directly adjacent to the plant promote growth of non‐infected plants by increasing N availability and removing salts from the rootzone (Bryla and Vargas, 2014). However, increased root rot with drip irrigation compared to sprinkler irrigation may be mitigated by moving irrigation lines further from the crown to avoid periodic saturation in the rootzone during irrigation events (de Silva et al., 1999).

Organic mulches have been shown to suppress Phytophthora on eucalyptus, frazer fir, and avocado (Downer et al., 2001; Richter et al., 2011b; Menge et al., 1994). Phytophthora hyphae are largely composed of β‐1,3, β‐1,4 β‐1,6 glucan linkages, and glucanase activity associated with microbial activity in these mulches cleaves glucan linkages causing hyphal lysis and sporangial abortion (Downer et al., 2001; Hegnauer and Hohl, 1978; Richter et al., 2011). In addition to organic mulches, black plastic geotextile mulches are prevalent for weed control in many perennial cropping systems including blueberry. Black plastic mulches increase soil temperature, influencing nutrient cycling, water relations, and other chemical and physical processes related to the rootzone (Larco, 2010; Larco et al., 2013).

Inhibition of Phytophthora zoospore motility by increasing soil calcium levels with gypsum is used for root rot suppression on avocado and raspberry. Greenhouse experiments have indicated gypsum also provides some suppression root rot disease suppression on highbush blueberry (Yeo et al. 2014a). Gypsum is a soluble salt, and is translocated throughout the soil profile with soil water movement.

Fungicides are systemically and soil applied to combat root rot disease, and may account for a substantial fraction of bluebery variable production costs (Brannen et al., 2009). Organic blueberry production is expanding, but risk of crop loss from P. cinnamomi is significantly increased when fungicides are excluded. Disease suppressive cultural practices are critical when fungicides are excluded from the production system. We performed a field experiment investigating the influence of cultural practices on root rot with highly susceptible blueberry cultivar ‘Draper,’ during the 2‐yr establishment phase. Treatments included all combinations of drip irrigation line placement, mulch type, and gypsum incorporation, and also an additional fungicide control.

Methods Site description A 0.1 ha field experiment to investigate influences of irrigation tubing placement, gypsum incorporation, and geotextile mulch on Phytophthora root rot disease of blueberry was installed at Oregon State University Lewis Brown Horticultural Research Farm, Corvallis, OR. ‘Duke’ blueberry was previously planted at the site in 2004, but was removed in 2010 due to Phytophthora root rot. Soil at the site is Malabon silty clay loam (fine, mixed, mesic Pachic Ultic Argixeroll), and was acidified with elemental sulfur incorporation in 2003. A soil test in Feb. 2012 indicated soil pH was 5.4, cation exchange capacity was 24.4 cmol / kg, and Bray‐P was 23 ppm (Kuo Testing Labs, Othello, WA; Brookside Labs, New Bremen, OH).

Site preparation and field layout The site was seeded in a cover crop mix of fava beans, vetch, oat in clover in fall 2011. The cover crop was killed with herbicide in Apr. 2012, and the field was tilled into the soil using a power‐spader implement on 9‐May 2012. Douglas‐fir (Pseudotsuga menziesii) sawdust (Lane Forest Products, Eugene OR) aged for 1 year was spread approximately 2.5 cm thick across the field and incorporated by rototilling. Rows were marked, and raised beds were formed 0.3 m high and 1 m wide using a tractor‐ drawn bed shaper adjusted to pull soil from a 3.05 m swath. The area between raised beds was seeded with annual ryegrass, which was mowed regularly after grass establishment.

Eight raised‐bed rows 45 m long were formed in a 28o N – 208o S row orientation. The center six rows were each planted with 54 3‐L size ‘Draper’ blueberry plants from Fall Creek Nursery (Lowell, OR) spaced 0.76 m apart between plants. Additionally, guard rows consisting of ‘Toro’ and ‘Reka’ were planted on

either side of the ‘Draper’ rows. Experimental plots consisted of six contiguous plants in the ‘Draper’ rows, with each row having nine plots.

Inoculum preparation Despite a history of P. cinnamomi root rot infection, the field was re‐inoculated to provide active inoculum in all plots. Inoculum was produced in autoclavable bags with filter patches (Fungi Perfecti, Sheldon, WA). Each bag was filled with 3 L of medium‐grade horticultural vermiculite, and 1.5 L of nutrient broth consisting of 10% v/v V‐8 juice (Campbells Soup Co, Camden, NJ) and 2 g / L CaCO3. Bags were placed in autoclave trays, covered with aluminum foil, and autoclaved for 55 minutes at 15 psi. After cooling for 24 hours, one 100 mm diameter petri plate fully colonized with P. cinnamomi growing on potato dextrose agar was sliced into 100 pieces and added to each bag. Bags were incubated in cardboard boxes under ambient greenhouse conditions and shaken at 2‐week intervals. Inoculum from all bags was homogenized in a cement mixer, and 1.56 kg moist inoculum was spread on each plot in a 30‐cm wide band incorporated 15‐cm deep with a front‐tine rototiller. Inoculum samples were collected at the time of application, and pathogen density was quantified on a 1:20 dilution w/v of 0.2% w/w water agar, by spreading 0.5 mL aliquots on PARPH selective agar, and counting the number of P. cinnamomi colonies per plate daily for 7 days. The field‐applied inoculum rate was calculated as 2.7 cfu / cm3 soil.

Treatment description Eight treatments were selected to include all combinations of wide or narrow drip irrigation line spacing, geotextile or sawdust mulch, and gypsum incorporation or no gypsum. Gypsum (21% Ca) was spread on the bed surface in a 30 cm wide band at 22,420 kg/ha in‐band rate , and incorporated with a narrow front‐tine rototiller to a depth of 15 cm following bed formation, but before planting. Two drip irrigation lines were installed per row of plants and secured to the ground either adjacent to each side of the crown (narrow spacing treatment) or 20 cm on either side (wide spacing treatment) using wire staples. In‐line emitters on the irrigation lines were spaced 30 cm apart, providing 1.02 L / hr each. Woven black plastic geotextile mulch (Berry Hill Drip Irrigation, Buffalo Junction VA) was secured to the ground on each side of the row with wire staples, overlapping in the center with cut‐outs for the plants. Sawdust mulch was applied 8 cm thick with a side‐discharge mulch spreader.

One additional treatment was included as a fungicide control, and was planted with narrow irrigation line spacing, sawdust mulch, and no gypsum incorporation. Mefenoxam (Ridomil Gold SL) was sprayed on the soil surface in a 45 cm wide band shortly after planting in May 2012, and again in May 2013. Application rate was 0.4033 mL Ridomil Gold SL (0.1827 mL Mefenoxam) / m row, and was mixed with 57 mL water / m row for application. Fosetyl‐Al (Aliette WDG) was applied to the canopy after fully‐ expanded leaves developed on the plants) in May 2012 and May 2013 at a rate of 3.36 kg / ha (0.62 g aluminum tris o‐ethyl phosphonate / plant), using 373 L water / ha.

Plant Establishment and Fertilization Blueberry cultivar ‘Draper’ was selected for this experiment because of its high susceptibility to Phytophthora root rot (Yeo et al., 2014b). 3‐L potted plants were obtained from Fall Creek Nursery (Lowell, OR) on 30‐Mar‐2012 and maintained under daily irrigation until planting on 15‐May‐2012. Soil analysis indicated that soil P was low at the site (23ppm Bray P1), and therefore and triple superphosphate (0‐45‐0) was mixed into each planting hole equivalent to 44.85 Kg P / prior to planting.

Liquid ammonium sulfate (9‐0‐0) fertilizer was supplied to the pants via fertigation using a Dosatron fertilizer injector located in the irrigation manifold. The fertilizer was applied beginning on 30 May 2012 with four biweekly applications of 28 kg∙ha‐1 N each. The first two applications`were injected at a 2% fertilizer concentration (v/v), and the second two applications were reduced to a 0.5% concentration. The following year, liquid urea (18‐0‐0) was applied 10 times between 17 Apr. and 17 July at a rate of 18 kg∙ha‐1 N each, injected into the drip system at 1% (v/v) fertilizer concentration.

Irrigation was scheduled four times per week during the growing season, using an automated irrigation controller, and was adjusted weekly based on plant size and reference ET obtained from the Corvallis AgriMet agricultural weather station (http://www.usbr.gov/pn/agrimet/).

Root infection incidence assessment A 2‐cm‐diameter core sampler was driven diagonally through the approximated root zone of the plants to assess percent root infection over time. One core from one plant in each plot was sampled on 26 July in 2012 and 25 Mar., 26 June, and 24 Sept. in 2013. Either 10 or 20 root sections, each ≈1 cm in length, were randomly sampled from the cores, rinsed in 1% bleach solution for 30 s, and placed on PARPH media, selective for Phytophthora species. One plant from each plot was also excavated for root rot

assessment in Dec. 2012 and Dec. 2013, following the first and second growing seasons. The roots were washed with a hose, and 1‐cm sections (20/plant in 2012; 10 /plant in 2013) were randomly sampled from each plant and placed in PARPH media. On each occasion, the media was incubated at 20oC and inspected daily at 100x magnification under a compound microscopy for 10 d to mark infected roots.

Influence of mulch type on in vitro P. cinnamomi hyphal growth Pure P. cinnamomi cultures inside sealed petri plates were buried in the soil at a depth of 15 cm. The isolate was obtained from a single hyphal tip culture collected originally in 2010 from infected ‘Draper’ plants located at the Lewis Brown Farm. Cultures were cut from fully colonized PDA plates, using a 5‐ mm cork borer, and transferred to new PDA petri plates under sterile conditions. Plates were wrapped with parafilm, and six plates were placed in a sealable plastic bag in two stacks of three. Bags were buried in the wide irrigation spacing, non‐gypsum plots, under both weed mat and sawdust, in only three of the six replications. Soil temperature was measured using thermocouples buried adjacent to the petri plates and was recorded diurnally using data loggers (HOBO Pendant, Onset Computer Corporation, Bourne, MA). Plates were buried on three separate occasions including 9 Aug. 2012, 18‐ Apr. 2013, and 11‐Jul. 2013, and hyphal dimensions were assessed after 6 ‐8 d.

Soil moisture measurement Soil moisture was measured in all plots on 12‐July and 7‐August, 2013 using a Trase I time domain reflectometry system (Soilmoisture Equipment Corp., Santa Barbara, CA) equipped with 15‐cm waveguides. Moisture was measured at three locations in the in each plot, directly in the middle of the raised bed.

Soil sampling and nutrient analysis Soil samples were collected using a 2‐cm diameter stainless core sampler to a depth of 15‐cm in June 2012, July 2013, and April 2014. Six cores were subsampled from each plot, directly in the center of the raised beds. Soils were air‐dried, ground, homogenized, and analyzed by Brookside Laboratories (New Bremen, OH) via the S001P soil test package including 1:1 pH and Mehlich III extractable cations.

Experimental design and statistical analysis Treatments were arranged in a split plot RCBD. Each raised‐bed row comprised one replication, and was divided randomly in half into geotextile weed mat and sawdust main plots. The gypsum and irrigation line treatments were randomly assigned within the mulch main plots.

The influence of main and interaction effects on plant biomass accumulation, root infection incidence, soil moisture, and leaf and soil solution nutrient concentration were modeled using Proc Mixed in SAS 9.2. Root infection incidence ranged from 0 to 100%, and were transformed using an arcsine square root function before analysis. Fixed effects in the model included replication, gypsum, irrigation spacing, mulch, two‐way interactions, gypsum*mulch, gypsum*irrigation spacing, irrigation spacing*mulch, and the three‐way interaction gypsum*irrigation spacing*mulch. Random effects included replication, and the replication*mulch interaction needed to generate main plot effects. Multiple comparisons of least square means were converted into mean separation letter format using the PDMIX800 SAS Macro (Saxton, 1998). Additional ANOVAS were performed using Proc GLM in SAS 9.2 to examine Fisher’s LSD mean separation for plant biomass and root infection incidence of all treatment combinations and the fungicide control, including only replication and treatment in the model.

Heat unit accumulations beneath sawdust and weed mat were calculated based on soil temperature readings recorded every 15‐min, and a 9o C minimum threshold (Zentmyer et al., 1976). Hyphal area of P. cinnamomi cultures incubated beneath either sawdust or weed mat were calculated based on measurement in two directions. Statistical differences in soil heat unit accumulation between sawdust and weed mat, and the associated differences in hyphal growth response were revealed using a general linear model including mulch type and replication as terms.

Results

Root Infection Incidence

Mulch Plants with weed mat had significantly higher root infection than those with sawdust mulch, in July 2012 and in June 2013 (Table 3; Figure 1). At the other four root sampling dates, no significant difference existed in root infection incidence between the two mulch types. Interaction effects between mulch type and either gypsum or irrigation line placement were non‐significant at all dates except December 13, when plants with weed mat and without gypsum had significantly higher root infection than all other mulch and gypsum combinations (Table 3).

Irrigation line placement Plants with widely‐spaced drip lines had significantly less root infection than those with drip lines located adjacent to the crown, in Dec. 2012 and in Mar. 2013 (Table 3; Figure 2). At the other four sampling dates, the difference was not statistically significant. Interactions between irrigation line

placement and either gypsum or mulch type were non‐significant, except in Sept. 2013, when plants with widely‐spaced drip lines and gypsum had significantly less root infection than those with other combinations of gypsum and irrigation spacing (Table 3).

Gypsum Plants amended with gypsum had significantly less root infection than those without gypsum, in July and December in 2012 and in September and December in 2013. Gypsum had no influence on root infection on the other two sampling dates (Table 5; Figure 3). The interaction effect between gypsum and mulch type was significant in Dec. 2013 and in indicated that plants with weed mat and no gypsum had significantly more infection than those with other combinations of mulch and gypsum. The interaction effect between gypsum and drip line spacing was also significant in Sept. 2013, revealing that plants with gypsum and sawdust had less infection than those with other combinations of gypsum and mulch (Table 3).

Non‐fungicde treatments compared to fungicide control Plants with weed mat, narrow drip lines, and no gypsum had the most root infection among all of the treatments on each sampling date (Table 3). On the other hand, plants with sawdust, widely spaced drip lines, and gypsum had the least infection among all of the non‐fungicide treatments on each date. Plants with sawdust, widely‐spaced drip lines, and gypsum had a similar level of infection as the fungicide‐ treated control plants on each date (Figure 4; Table 3).

Biomass Accumulation

Mulch – Year 1 and 2 Shoot and root biomass was similar between plants grown with sawdust mulch and weed mat, after both the first and second growing seasons (Table 1;2). Biomass was not affected by interactions between mulch type and gypsum or mulch type and drip line placement in either season.

Irrigation line placement – Year 1 Plants with narrow drip lines placement had much more shoot biomass than those with widely spaced drip lines but less root biomass (Table 1). Although each treatment was fertilized with a total of 110 kg∙ha‐1 N, leaf N concentrations of plants with a wide drip line spacing were well below the sufficiency threshold while those of plants with a narrow spacing were near the threshold (Table 4).

Gypsum – Year 1 Gypsum had no influence on shoot or root biomass during the first growing season. There were no significant interactions between gypsum and the mulch or irrigation placement treatments (Table 1).

Irrigation line placement and Gypsum Interaction – Year 2 Interactions between gypsum application and drip line placement became apparent during the second growing season. In year 2, plants with wide‐spaced drip lines and gypsum had significantly higher leaf, shoot, and root dry weights than those with the same spacing and no gypsum, and all plants with narrow drip line placements (Table 2).

Non‐fungicide treatments compared to fungicide control – Year 1 and 2 Despite significant differences between the non‐fungicide treatments, every plant biomass component of every non‐fungicide treatment factorial combination was lower than the fungicide control plants in both years (Table 2). Plants with wide irrigation line placement and gypsum had the greatest amount of shoot and root biomass of the non‐chemical treatments after year 2, regardless of mulch type, but only ≈35% of the biomass in the fungicide control (Figure 4). Relative root biomass of plants with narrow irrigation lines without gypsum was 20% of the fungicide controls, and relative root biomass was 23% (Table 2).

Soil Temperature and influence on P. cinnamomi Both heat unit accumulation and P. cinnamomi hyphal growth on sealed, buried PDA petri plates were significantly higher beneath weed mat than sawdust mulch at three different monitoring periods (Table 9). Daily minimum temperatures were similar between sawdust and weed mat. However, the daily maximum temperatures beneath weed mat were generally greater than the maximum temperatures beneath the sawdust (Table 6; Figure 5).

Soil moisture Soil moisture was measured in the center of raised beds on two dates, between 1 to 2 h after a 15‐min irrigation event. Soil moisture was significantly higher near the plant crown with the narrow irrigation line placement than with wider placement (Figure 6). Despite 15% additional irrigation volume application on the weed mat treatments, soil moisture was similar between the weed mat and the sawdust treatments. Gypsum application also had no effect on soil moisture content (Figure 6).

Calcium levels in soil solution Placement of irrigation lines adjacent to the plant crown reduced Ca2+ concentration in the plant root zone by approximately 50% compared to the gypsum amended plots with wide irrigation line placement after 5 months (Figure 7). After 16 months, Ca2+ concentration in the rootzone of gypsum amended plots with narrow irrigation line placement was not significantly different than the plots without gypsum amendment. Ca2+ concentration at the plant crown in the wide irrigation line treatments with gypsum decreased between 5 and 16 months after application due to over‐winter leaching, however the difference was not significant (Figure 7).

Mehlich III extractable calcium Over the course of the experiment, the applied gypsum leached substantially. In June 2012, shortly after planting, extractable Ca was 4524 mg/kg in the gypsum‐amended plots and 1968 mg/kg in the non‐ amended plots. After 14 months extractable Ca had decreased to 2722 mg/kg, further decreased to 2453 mg/kg 23 months after planting (Table 7).

Discussion

Cultural practices including irrigation line placement, gypsum, and much had additive effects on reducing root rot disease development, and are important considerations for both organic and conventional growers. Without fungicide, plants grown with a combination of widely spaced drip lines and gypsum incorporation produced more biomass after 2 years than those with widely spaced drip lines and no gypsum or closely spaced drip lines either with or without gypsum. Surprisingly, mulch type had no effect on plant biomass after two years, even though root infection was greater with weed mat than with sawdust mulch on two out of six sampling dates, and in situ incubations indicated that P. cinnamomi hyphae grows more rapidly under weed mat. The lack of difference in biomass between mulch types could be attributed to more rapid nutrient cycling or water uptake with weed mat which enhanced plant growth overcoming the greater disease pressure (Larco et al., 2013).

Suppression of P. cinnamomi from gypsum is caused by high levels of Ca2+ in soil solution, which inhibits zoospore motility and survival (Byrt et al., 1982; von Broembsen and Deacon, 1997). While gypsum was applied equally to the amended plots, drip line placement had a profound effect on Ca2+ translocation to or away from the root zone. Placement of the lines close to the plants reduced Ca2+ concentration by

approximately 50% after 5 months compared to the wider placement, but concentrations were similar to the non‐amended plots after 16 months, regardless of drip spacing. On plots with gypsum and widely spaced irrigation lines, Ca2+ in soil solution decreased between 5 and 16 months, however the difference was not significant (Figure 7).

Frequent saturation of the plant rootzone is conducive to P. cinnamomi root rot by enabling rapidly repeating zoospore cycles of infection (DeSilva et al., 1999). Placement of drip irrigation lines 20‐cm on either side of the plant reduced the volumetric moisture content in the center of the raised bed after irrigation events compared to lines adjacent to the crown (Figure 6). Although shoot growth was greater in the first growing season with lines adjacent to the crown due to enhanced N and water availability, root infection incidence also increased. After the second growing season, root biomass of plants with narrow irrigation line placement was approximately 50% of the previous season.

Irrigation management minimizing root zone moisture near saturation while also replenishing evapotranspiration losses is critical for maximizing plant growth and minimizing root infection in the presence of Phytophthora. Drip irrigation on blueberry enhances plant growth during establishment compared to sprinkler irrigation in the absence of Phytophthora infection (Bryla et al., 2011). However plant growth is significantly reduced by increased root infection with drip irrigation placed near the plant crown when plants are infected by Phytophthora during establishment (Bryla and Linderman, 2007). Under the paradigm of drip irrigation, this experiment demonstrated no dramatic effect of drip line placement relative to the plant crown on Phytophthora root rot disease. However, the influence of placing dual irrigation lines 20‐cm from the crown compared to adjacent to the crown made a significant difference in reducing disease and enhancing biomass if gypsum was incorporated to the soil before planting.

Despite the significant differences in root infection incidence and plant biomass between the cultural treatments, plants in the fungicide control plots with narrow irrigation line placement and sawdust mulch had dramatically more shoot and root biomass, after both the first and second growing seasons (Table 2; 4; Figure 4). Although root infection incidence between the fungicide control treatment and the treatment with widely‐spaced irrigation lines, gypsum incorporation, and sawdust, was equivalent, biomass of plants was dramatically greater with fungicide and narrow drip irrigation lines than gypsum and wide irrigation lines (Table 6; Figure 4). Reduced water and nitrogen availability, and possibly

salinity accumulation at the crown for plants with widely placed irrigation lines likely contributed to these growth differences, given equivalent root infection incidence (Bryla and Vargas, 2014; Patten et al., 1988). Plants with widely‐spaced irrigation lines were severely nitrogen deficient during the first

+ growing season due to low mobility of NH4 fertilizer injected through the drip system, far from the rootzone (Table 4). Furthermore, minor drought stress symptoms (shoot dieback) occurred in some of the plants with wide irrigation lines, as lateral diffusion of water was inhibited by the freshly‐tilled soil structure favoring rapid infiltration downward. By the second season, the soil structure had settled, rootzones were larger, N was supplied with urea, and none of the N or water deficiencies recurred (Table 5). Perhaps if plants were installed in the fall to allow soil settling before the beginning of the growing season, drought stress may not have occurred, and N fertilizer could be supplied more judiciously to avoid the N deficiency.

Suppression of Phytophthora root rot with gypsum on blueberry in this experiment was not as effective as reports of Phytophthora suppression with gypsum on avocado and red raspberry (Maloney et al., 2005; Messenger et al., 2000a; Pinkerton et al., 2009). Blueberries have a fine, fibrous, shallow root system requiring frequent irrigation during the growing season for replenishing evapotranspiration losses, which may inherently predispose blueberries to enhanced levels of root rot disease compared to avocado and raspberry with more deep root systems requiring less frequent irrigation (de Silva et al., 1999). Gypsum was also subjected to leaching from the rootzone during the course of the study. The experiment was terminated after 23‐months at which time extractable Ca was approximately half of the original levels in the gypsum‐amended plots, although the majority of Ca leaching occurred after one growing season (Table 7).

Cultivars of blueberry vary widely in susceptibility or tolerance to P. cinnamomi, and a highly susceptible cultivar (‘Draper’) was chosen for this experiment to provide a worst‐case scenario for root rot disease in combination with silty clay loam soil at the site (Yeo et al., 2014b). Control of blueberry root rot disease is difficult. An integrated disease management plan requires a suite of strategies including site selection, pathogen exclusion, cultivar selection, scrutinous irrigation scheduling, irrigation line placement, fungicides, gypsum, and mulch selection, all providing cumulative additive effects of reducing disease. With more tolerant cultivars or on more rapidly draining soils, widely‐spaced drip irrigation lines and gypsum may provide disease suppression more comparable to fungicide treatment than in this experiment, and future research efforts could evaluate the effects of mulch, drip irrigation

line placement on gypsum on more tolerant cultivars, especially maintaining sufficient N levels during the first year.

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Table 1. Biomass of ‘Draper’ blueberry grown in Phytophthora cinnamomi infested soil after one growing season. Mulch Drip Gypsum Leaf mass (g) Stem mass (g) Shoot mass (g) Root mass (g) Main Effect Means Geotextile 32.6 ±4.91 az 53.6 ±4.32 a 86.1 ±8.99 a 45.5 ±3.48 a Sawdust 33.1 ±4.13 a 62.8 ±4.33 a 95.9 ±7.64 a 42.8 ±3.41 a

Narrow 50.2 ±3.66 a 70.9 ±4.20 a 121.1 ±7.16 a 39.1 ±2.80 b Wide 15.5 ±1.26 b 45.5 ±2.76 b 61.0 ±3.39 b 49.1 ±3.73 a

N 29.5 ±4.10 a 54.8 ±4.43 a 84.3 ±7.73 a 42.6 ±3.62 a Y 36.2 ±4.84 a 61.6 ±4.32 a 97.8 ±8.81 a 45.6 ±3.27 a

2‐Way Interaction Means Geotextile Narrow 49.9 ±6.59 a 69.8 ±5.08 a 119.7 ±11.05 a 40.3 ±3.52 a Geotextile Wide 15.2 ±1.61 b 37.4 ±2.16 c 52.6 ±3.37 b 50.7 ±5.78 a Sawdust Narrow 50.5 ±3.52 a 72.0 ±6.90 a 122.5 ±9.58 a 38.0 ±4.50 a Sawdust Wide 15.8 ±2.02 b 53.6 ±3.91 b 69.4 ±4.89 b 47.6 ±4.92 a

Geotextile N 28.5 ±6.59 a 50.0 ±5.82 b 78.4 ±11.85 b 44.8 ±5.40 a Geotextile Y 36.7 ±7.37 a 57.2 ±6.47 ab 93.8 ±13.68 ab 46.2 ±4.64 a Sawdust N 30.6 ±5.15 a 59.6 ±6.63 ab 90.2 ±10.16 ab 40.4 ±4.96 a Sawdust Y 35.7 ±6.59 a 66.1 ±5.70 a 101.7 ±11.60 a 45.1 ±4.80 a

Narrow N 45.0 ±4.80 a 65.0 ±6.87 a 110.0 ±10.16 a 36.2 ±3.56 b Narrow Y 55.4 ±5.29 a 76.8 ±4.49 a 132.2 ±9.43 a 42.0 ±4.31 ab Wide N 14.0 ±1.87 b 44.6 ±3.98 b 58.6 ±5.14 b 49.1 ±5.86 ab Wide Y 16.9 ±1.67 b 46.5 ±3.99 b 63.4 ±4.54 b 49.2 ±4.87 ab

3‐Way Interaction Means Geotextile Narrow N 44.6 ±9.09 a 64.0 ±7.49 ab 108.6 ±15.07 a 38.2 ±5.59 a Geotextile Narrow Y 55.2 ±9.86 a 75.6 ±6.65 a 130.8 ±16.17 a 42.3 ±4.64 a Geotextile Wide N 12.3 ±2.04 b 36.0 ±3.79 c 48.3 ±5.11 b 51.5 ±8.92 a Geotextile Wide Y 18.1 ±1.99 b 38.8 ±2.30 c 56.9 ±4.06 b 50.0 ±8.20 a Sawdust Narrow N 45.4 ±4.32 a 66.0 ±12.30 ab 111.4 ±15.04 a 34.1 ±4.80 a Sawdust Narrow Y 55.5 ±5.07 a 78.1 ±6.62 a 133.6 ±11.35 a 41.8 ±7.75 a Sawdust Wide N 15.8 ±3.17 b 53.1 ±5.07 bc 68.9 ±6.89 b 46.7 ±8.33 a Sawdust Wide Y 15.8 ±2.80 b 54.1 ±6.45 bc 69.9 ±7.59 b 48.4 ±6.08 a Sawdust Narrow Fungicide 65.4 ± 5.53 y 110.0 ± 10.34 y 175.4 ± 12.58 y 71.9 ± 9.29 y

Model Term p‐value p‐value p‐value p‐value Mulch 0.8918 0.1144 0.2680 0.6264 Drip <0.0001 <0.0001 <0.0001 0.0199 Gypsum 0.1037 0.1667 0.0967 0.4711 Mulch*Drip 0.9977 0.1589 0.3807 0.9174 Mulch*Gypsum 0.6953 0.9422 0.8091 0.6874 Drip*Gypsum 0.3538 0.3151 0.2794 0.4834 Mulch*Drip*Gypsum 0.744 0.9002 0.8094 0.9761 z Letters indicate LSD between treatment combinations yThe fungicide control was not part of the factorial design, and was not included in the model, thus LSD comparisons do not apply

Table 2. 2013 Main and interaction effects of mulch, drip irrigation spacing, and gypsum incorporation on plant biomass after two growing seasons. Mulch Drip Gypsum Leaf mass (g) Stem mass (g) Shoot mass (g) Root mass (g) Main Effect Means Geotextile 34.2 ±4.51 a 50.7 ±4.99 b 84.9 ±9.36 a 41.0 ±6.16 a Sawdust 41.8 ±3.74 a 64.6 ±4.24 a 106.4 ±7.69 a 35.8 ±4.97 a

Narrow 31.3 ±2.40 b 55.4 ±3.44 a 86.7 ±5.62 a 26.0 ±1.83 b Wide 44.7 ±5.10 a 59.9 ±5.90 a 104.6 ±10.87 a 50.8 ±6.83 a

N 27.8 ±2.08 b 46.1 ±3.13 b 73.9 ±4.94 b 33.0 ±3.98 a Y 48.2 ±4.71 a 69.2 ±5.07 a 117.4 ±9.54 a 43.8 ±6.70 a

2‐Way Interaction Means Geotextile Narrow 25.7 ±2.90 b 45.5 ±3.11 b 71.3 ±5.79 b 25.2 ±2.95 c Geotextile Wide 42.7 ±7.97 a 56.0 ±9.47 ab 98.6 ±17.28 a 56.8 ±10.23 a Sawdust Narrow 36.9 ±3.15 ab 65.3 ±4.70 a 102.2 ±7.41 a 26.9 ±2.27 bc Sawdust Wide 46.7 ±6.66 a 63.8 ±7.29 a 110.5 ±13.75 a 44.7 ±9.15 ab

Geotextile N 24.2 ±2.06 c 36.6 ±2.11 c 60.8 ±3.84 c 33.8 ±7.76 a Geotextile Y 44.2 ±7.91 ab 64.8 ±7.98 ab 109.1 ±15.67 ab 48.3 ±9.43 a Sawdust N 31.4 ±3.39 b 55.6 ±4.51 b 87.0 ±7.48 b 32.2 ±2.45 a Sawdust Y 52.2 ±5.23 a 73.5 ±6.35 a 125.8 ±11.08 a 39.3 ±9.75 a

Narrow N 27.2 ±3.30 b 50.6 ±4.59 bc 77.7 ±7.60 b 25.9 ±2.28 b Narrow Y 35.5 ±3.16 b 60.2 ±4.91 b 95.7 ±7.71 b 26.2 ±2.96 b Wide N 28.4 ±2.67 b 41.7 ±4.05 c 70.1 ±6.44 b 40.1 ±7.21 b Wide Y 61.0 ±7.30 a 78.1 ±8.29 a 139.1 ±15.36 a 61.4 ±11.06 a

3‐Way Interaction Means Geotextile Narrow N 22.0 ±3.74 d 39.0 ±3.53 c 61.0 ±7.08 c 21.1 ±1.35 c Geotextile Narrow Y 29.5 ±4.19 cd 52.0 ±3.62 bc 81.5 ±7.46 bc 29.4 ±5.43 bc Geotextile Wide N 26.4 ±1.67 cd 34.3 ±2.19 c 60.6 ±3.86 c 46.5 ±14.09 abc Geotextile Wide Y 59.0 ±13.05 ab 77.6 ±14.20 a 136.7 ±26.84 a 67.2 ±14.80 a Sawdust Narrow N 32.3 ±4.84 cd 62.1 ±5.19 ab 94.4 ±9.63 bc 30.8 ±3.42 bc Sawdust Narrow Y 41.5 ±3.43 bc 68.4 ±8.14 ab 110.0 ±11.15 ab 22.9 ±2.20 c Sawdust Wide N 30.5 ±5.17 cd 49.1 ±6.74 bc 79.5 ±11.47 bc 33.7 ±3.72 bc Sawdust Wide Y 62.9 ±7.94 a 78.6 ±10.05 a 141.5 ±17.76 a 55.7 ±17.49 ab Sawdust Narrow Fungicide 125.7 ± 12.08 y 206.5 ± 10.26 y 332.2 ± 15.74 y 130.2 ± 16.82 y

Model Term p‐value p‐value p‐value p‐value Mulch 0.1689 0.0392 0.0622 0.4553 Drip 0.0033 0.3732 0.0552 0.0007 Gypsum <0.0001 <0.0001 <0.0001 0.1073 Mulch*Drip 0.4007 0.2431 0.2968 0.2978 Mulch*Gypsum 0.9262 0.3118 0.6008 0.5717 Drip*Gypsum 0.0071 0.0116 0.0078 0.1148 Mulch*Drip*Gypsum 0.9053 0.723 0.8002 0.5087 z Letters indicate LSD between treatment combinations yThe fungicide control was not part of the factorial design, and was not included in the model, thus LSD comparisons do not apply

Table 3. Root infection incidence at five sampling dates over two growing seasons. Jul 2012 Dec 2012 Mar 2013 Jun 2013 Sep 2013 Dec 2013 Main Effect Contrasts: Mulch Weed mat 15.4%x ± 3.2% azy 26.9% ± 3.0% a 20.4% ± 2.1% a 20.4% ± 0.0 a 15.0% ± 2.7% a 17.1% ± 2.7% a Sawdust 6.9% ± 2.2% b 20.7% ± 2.2% a 16.7% ± 2.7% a 7.5% ± 0.0 b 10.4% ± 2.8% a 10.0% ± 2.2% a

Drip Narrow 10.8% ± 3.3% a 27.2% ± 2.6% a 24.6% ± 2.3% a 17.5% ± 3.7% a 14.2% ± 2.3% a 14.6% ± 2.3% a Wide 11.5% ± 2.5% a 20.4% ± 2.6% b 12.5% ± 1.8% b 10.4% ± 2.6% a 11.3% ± 3.1% a 12.5% ± 2.7% a

Gypsum No Gypsum 18.1% ± 3.3% a 26.5% ± 2.7% a 20.2% ± 2.5% a 15.8% ± 3.1% a 19.6% ± 3.0% a 17.5% ± 2.8% a Gypsum 4.2% ± 1.3% b 21.1% ± 2.6% b 16.9% ± 2.3% a 12.1% ± 3.5% a 5.8% ± 1.6% b 9.6% ± 1.9% b

Interaction Effect Contrasts: Mulch Gypsum Weed Mat N 24.6% ± 4.8% b 32.5% ± 4.7% a ± 2.9% a 24.2% ± 5.0% a 20.8% ± 4.2% a 25.0% ± 3.1% a Weed Mat Y 6.3% ± 2.1% c 21.3% ± 3.1% b 20.0% ± 3.1% a 16.7% ± 6.1% ab 9.2% ± 2.6% b 9.2% ± 2.9% b Sawdust N 11.7% ± 3.9% a 20.4% ± 1.6% b 19.6% ± 4.1% a 7.5% ± 1.3% b 18.3% ± 4.4% ab 10.0% ± 3.7% b Sawdust Y 2.1% ± 1.3% bc 21.0% ± 4.2% b 13.8% ± 3.4% a 7.5% ± 3.0% b 2.5% ± 1.3% c 10.0% ± 2.5% b

Drip Gypsum Narrow N 18.3% ± 5.6% a 28.8% ± 4.0% a 27.1% ± 3.5% a 16.7% ± 4.8% a 18.3% ± 3.7% a 15.8% ± 3.6% a Narrow Y 3.3% ± 1.8% b 25.6% ± 3.5% a 22.1% ± 3.0% ab 18.3% ± 5.9% a 10.0% ± 2.5% a 13.3% ± 3.1% ab Wide N 17.9% ± 3.9% a 24.2% ± 3.7% a 13.3% ± 2.2% bc 15.0% ± 4.0% a 20.8% ± 4.8% a 19.2% ± 4.5% a Wide Y 5.0% ± 1.9% b 16.7% ± 3.5% b 11.7% ± 2.9% c 5.8% ± 2.9% b 1.7% ± 1.1% b 5.8% ± 1.5% b

Mulch Drip Weed Mat Narrow 15.8% ± 5.3% a 30.4% ± 4.2% a 26.7% ± 2.3% a 25.8% ± 6.2% a 19.2% ± 3.6% a 17.5% ± 3.7% a Weed Mat Wide 15.0% ± 3.9% a 23.3% ± 4.2% ab 14.2% ± 2.4% bc 15.0% ± 4.5% ab 10.8% ± 3.8% b 16.7% ± 4.0% a Sawdust Narrow 5.8% ± 3.4% b 24.0% ± 3.0% ab 22.5% ± 4.0% ab 9.2% ± 2.6% b 9.2% ± 2.3% b 11.7% ± 2.7% ab Sawdust Wide 7.9% ± 3.0% ab 17.5% ± 3.0% b 10.8% ± 2.7% c 5.8% ± 1.9% b 11.7% ± 5.2% b 8.3% ± 3.4% b

Fungicide ‐ Narrow, sawdust, no gypw 0% ± 0.0% 15% ± 3.2% 2% ± 1.8% 4% ± 3.7% 14% ± 5.5% 4% ± 2.2%

Model Term p‐value p‐value p‐value p‐value p‐value p‐value Mulch 0.0375 0.0985 0.1949 0.0420 0.1039 0.0907 Drip 0.5061 0.0150 0.0005 0.0966 0.0592 0.4617 Gypsum <0.0001 0.0486 0.2092 0.1054 <.0001 0.0333 Mulch*Drip 0.5531 0.8236 0.9468 0.5486 0.1474 0.4607 Mulch*Gypsum 0.3558 0.2689 0.3374 0.4571 0.2709 0.0075 Drip*Gypsum 0.6819 0.2797 0.6502 0.1578 0.0406 0.1681 Mulch*Drip*Gypsum 0.4311 0.1893 0.8931 0.2668 0.7130 0.4600 z Mean separation by Fisher's LSD method. Mean separation letters only indicate differences within each main or interaction effect, for one date y All statistics were performed on arcsine square root transformed data x Means and standard error were calculated from the non‐transformed data wThe fungicide control was not included in the factorial design, therefore LSD comparisons are not provided.

Table 4. Leaf tissue nutrient concentration Aug 2, 2012, 10‐wks after planting (Year 1). Mulch Drip Gypsum N (g 100g‐1)Ca (g 100g‐1) K (g 100g‐1) Mg (g 100g‐1) S (g 100g‐1) P (g 100g‐1) Main Effect Means Geotextile 1.28 ±0.087 a 0.54 ±0.013 a 0.49 ±0.014 b 0.17 ±0.003 a 0.23 ±0.015 b 0.09 ±0.003 b Sawdust 1.35 ±0.088 a 0.55 ±0.015 a 0.53 ±0.017 a 0.17 ±0.004 a 0.28 ±0.018 a 0.10 ±0.003 a

Narrow 1.69 ±0.037 a 0.54 ±0.013 a 0.48 ±0.007 a 0.17 ±0.003 a 0.26 ±0.010 a 0.10 ±0.002 a Wide 0.94 ±0.040 b 0.55 ±0.015 a 0.54 ±0.020 b 0.17 ±0.004 a 0.25 ±0.022 a 0.09 ±0.004 b

N 1.30 ±0.090 a 0.52 ±0.012 b 0.46 ±0.007 b 0.17 ±0.003 b 0.19 ±0.011 b 0.10 ±0.004 a Y 1.33 ±0.085 a 0.58 ±0.013 a 0.57 ±0.015 a 0.18 ±0.004 a 0.31 ±0.013 a 0.10 ±0.003 a

2‐Way Interaction Means GeotextileNarrow 1.68 ±0.034 a 0.56 ±0.013 ab 0.47 ±0.011 c 0.17 ±0.003 b 0.24 ±0.010 b 0.10 ±0.003 a GeotextileWide 0.87 ±0.035 b 0.52 ±0.023 b 0.52 ±0.025 b 0.17 ±0.006 b 0.22 ±0.029 c 0.08 ±0.003 b Sawdust Narrow 1.71 ±0.068 a 0.53 ±0.022 b 0.50 ±0.009 b 0.17 ±0.005 b 0.28 ±0.017 a 0.10 ±0.003 a Sawdust Wide 1.00 ±0.069 b 0.57 ±0.018 a 0.56 ±0.031 a 0.18 ±0.006 a 0.28 ±0.032 a 0.10 ±0.006 a

Geotextile N 1.24 ±0.128 a 0.50 ±0.017 b 0.44 ±0.009 d 0.16 ±0.005 b 0.17 ±0.014 d 0.09 ±0.004 b Geotextile Y 1.31 ±0.123 a 0.58 ±0.014 a 0.55 ±0.017 b 0.17 ±0.005 ab 0.29 ±0.012 b 0.09 ±0.005 b Sawdust N 1.36 ±0.130 a 0.53 ±0.016 b 0.48 ±0.009 c 0.17 ±0.004 b 0.21 ±0.013 c 0.10 ±0.006 a Sawdust Y 1.34 ±0.123 a 0.57 ±0.023 a 0.59 ±0.025 a 0.18 ±0.007 a 0.34 ±0.020 a 0.10 ±0.004 ab

Narrow N 1.69 ±0.036 a 0.53 ±0.018 bc 0.46 ±0.010 c 0.17 ±0.004 b 0.23 ±0.009 c 0.10 ±0.002 a Narrow Y 1.69 ±0.067 a 0.55 ±0.019 b 0.51 ±0.008 b 0.17 ±0.004 b 0.28 ±0.015 b 0.10 ±0.003 a Wide N 0.91 ±0.071 b 0.50 ±0.016 c 0.45 ±0.010 c 0.16 ±0.004 b 0.15 ±0.008 d 0.10 ±0.007 ab Wide Y 0.96 ±0.039 b 0.60 ±0.017 a 0.63 ±0.014 a 0.19 ±0.006 a 0.34 ±0.016 a 0.09 ±0.005 b

3‐Way Interaction Means GeotextileNarrow N 1.65 ±0.062 a 0.54 ±0.023 bc 0.44 ±0.011 e 0.17 ±0.006 b 0.21 ±0.008 e 0.10 ±0.004 ab GeotextileNarrow Y 1.71 ±0.031 a 0.57 ±0.014 abc 0.50 ±0.007 c 0.17 ±0.003 bc 0.27 ±0.010 cd 0.10 ±0.003 ab GeotextileWide N 0.83 ±0.032 b 0.46 ±0.013 d 0.44 ±0.014 e 0.15 ±0.005 c 0.13 ±0.007 g 0.08 ±0.002 c GeotextileWide Y 0.92 ±0.061 b 0.58 ±0.026 ab 0.59 ±0.018 b 0.18 ±0.008 ab 0.30 ±0.020 b 0.08 ±0.007 c Sawdust Narrow N 1.74 ±0.036 a 0.53 ±0.029 c 0.49 ±0.009 cd 0.17 ±0.006 bc 0.25 ±0.012 d 0.10 ±0.002 ab Sawdust Narrow Y 1.68 ±0.137 a 0.54 ±0.037 bc 0.51 ±0.015 c 0.17 ±0.008 bc 0.30 ±0.029 bc 0.11 ±0.006 ab Sawdust Wide N 0.99 ±0.136 b 0.54 ±0.018 bc 0.47 ±0.014 de 0.17 ±0.005 bc 0.17 ±0.005 f 0.11 ±0.011 a Sawdust Wide Y 1.00 ±0.049 b 0.61 ±0.022 a 0.66 ±0.009 a 0.19 ±0.008 a 0.38 ±0.012 a 0.09 ±0.005 bc

Target Levely 1.76‐2.0 0.41 ‐ 0.8 0.40‐0.70 0.13‐0.25 0.11‐0.16 >0.10 Deficiency Levelx 1.50 0.3 0.4 0.2 0.05 0.1

Model Term p‐value p‐value p‐value p‐value p‐value p‐value Mulch 0.2299 0.3192 0.0075 0.2746 0.0035 0.0265 Drip <0.0001 0.7574 <0.0001 0.1059 0.1834 0.0148 Gypsum 0.6818 <.0001 <0.0001 0.0059 <0.0001 0.6354 Mulch*Drip 0.3847 0.0052 0.2089 0.0322 0.1364 0.0072 Mulch*Gypsum 0.4024 0.1948 0.8201 0.5989 0.5225 0.3867 Drip*Gypsum 0.6498 0.0034 <0.0001 0.0004 <0.0001 0.0898 Mulch*Drip*Gypsum 0.8771 0.4900 0.0209 0.8060 0.3096 0.1104 z Letters indicate LSD between treatment combinations y Hart et al., 2006 x Erb, 1998

Table 5. Leaf tissue nutrient concentration August 1, 2013, 14‐months after planting (Year 2). Mulch Drip Gypsum N (g 100g‐1)Ca (g 100g‐1) K (g 100g‐1) Mg (g 100g‐1) S (g 100g‐1) P (g 100g‐1) Main Effect Means Geotextile 1.60 ±0.063 az 0.42 ±0.052 a 0.52 ±0.053 a 0.16 ±0.011 a 0.11 ±0.007 a 0.10 ±0.004 a Sawdust 1.89 ±0.076 b 0.48 ±0.019 a 0.47 ±0.013 a 0.19 ±0.008 a 0.12 ±0.004 a 0.10 ±0.002 a

Narrow 2.04 ±0.050 a 0.46 ±0.052 a 0.47 ±0.052 a 0.19 ±0.010 a 0.12 ±0.006 a 0.10 ±0.004 a Wide 1.45 ±0.042 b 0.44 ±0.021 a 0.53 ±0.012 a 0.17 ±0.010 a 0.12 ±0.005 a 0.10 ±0.003 a

N 1.69 ±0.086 a 0.38 ±0.019 b 0.46 ±0.018 a 0.18 ±0.010 a 0.10 ±0.002 b 0.10 ±0.002 a Y 1.80 ±0.063 b 0.52 ±0.048 a 0.53 ±0.050 a 0.18 ±0.010 a 0.13 ±0.006 a 0.11 ±0.004 a

2‐Way Interaction Means Geotextile Narrow 1.86 ±0.039 b 0.46 ±0.101 a 0.50 ±0.105 a 0.18 ±0.014 a 0.12 ±0.010 a 0.10 ±0.008 a Geotextile Wide 1.35 ±0.056 d 0.39 ±0.028 a 0.54 ±0.023 a 0.15 ±0.016 b 0.11 ±0.008 a 0.09 ±0.004 a Sawdust Narrow 2.22 ±0.052 a 0.45 ±0.031 a 0.43 ±0.017 a 0.20 ±0.016 a 0.12 ±0.005 a 0.10 ±0.003 a Sawdust Wide 1.56 ±0.044 c 0.50 ±0.021 a 0.51 ±0.011 a 0.19 ±0.007 a 0.12 ±0.005 a 0.11 ±0.004 a

Geotextile N 1.52 ±0.103 c 0.34 ±0.021 b 0.46 ±0.030 a 0.16 ±0.013 a 0.10 ±0.004 b 0.10 ±0.004 a Geotextile Y 1.68 ±0.070 b 0.51 ±0.097 a 0.58 ±0.100 a 0.17 ±0.019 a 0.13 ±0.012 a 0.10 ±0.008 a Sawdust N 1.85 ±0.124 a 0.43 ±0.028 ab 0.46 ±0.020 a 0.19 ±0.015 a 0.10 ±0.003 b 0.10 ±0.003 a Sawdust Y 1.93 ±0.093 a 0.53 ±0.016 a 0.49 ±0.016 a 0.19 ±0.008 a 0.13 ±0.002 a 0.11 ±0.004 a

Narrow N 2.03 ±0.080 a 0.36 ±0.027 b 0.39 ±0.018 b 0.18 ±0.016 ab 0.11 ±0.004 b 0.10 ±0.004 a Narrow Y 2.05 ±0.063 a 0.55 ±0.094 a 0.54 ±0.101 a 0.21 ±0.013 a 0.13 ±0.010 a 0.11 ±0.008 a Wide N 1.35 ±0.062 c 0.40 ±0.028 ab 0.52 ±0.015 ab 0.18 ±0.014 ab 0.10 ±0.003 b 0.09 ±0.003 a Wide Y 1.56 ±0.038 b 0.49 ±0.027 ab 0.53 ±0.020 ab 0.16 ±0.013 b 0.13 ±0.007 a 0.11 ±0.004 a

3‐Way Interaction Means Geotextile Narrow N 1.84 ±0.049 b 0.34 ±0.029 b 0.38 ±0.028 b 0.17 ±0.012 abc 0.11 ±0.006 bcd 0.10 ±0.006 ab Geotextile Narrow Y 1.88 ±0.064 b 0.58 ±0.195 a 0.62 ±0.205 a 0.19 ±0.026 ab 0.12 ±0.020 abc 0.10 ±0.015 ab Geotextile Wide N 1.21 ±0.073 d 0.34 ±0.032 b 0.53 ±0.030 ab 0.15 ±0.023 bc 0.10 ±0.004 d 0.09 ±0.003 b Geotextile Wide Y 1.48 ±0.038 c 0.43 ±0.037 ab 0.54 ±0.036 ab 0.14 ±0.024 c 0.13 ±0.014 ab 0.10 ±0.006 ab Sawdust Narrow N 2.22 ±0.105 a 0.38 ±0.045 ab 0.40 ±0.024 ab 0.18 ±0.031 abc 0.11 ±0.007 bcd 0.10 ±0.005 ab Sawdust Narrow Y 2.23 ±0.030 a 0.52 ±0.022 ab 0.46 ±0.018 ab 0.22 ±0.005 a 0.13 ±0.003 a 0.11 ±0.004 ab Sawdust Wide N 1.49 ±0.061 c 0.47 ±0.027 ab 0.51 ±0.010 ab 0.20 ±0.006 a 0.10 ±0.003 cd 0.10 ±0.003 ab Sawdust Wide Y 1.64 ±0.050 c 0.54 ±0.025 ab 0.52 ±0.021 ab 0.17 ±0.008 abc 0.13 ±0.004 a 0.11 ±0.006 a

Target Levely 1.76‐2.0 0.41 ‐ 0.8 0.40‐0.70 0.13‐0.25 0.11‐0.16 >0.10 Deficiency Levelx 1.50 0.3 0.4 0.2 0.05 0.1

Model Term p‐value p‐value p‐value p‐value p‐value p‐value Mulch 0.0010 0.3606 0.5156 0.0705 0.5787 0.3542 Drip <0.0001 0.8270 0.2319 0.0612 0.7587 0.5353 Gypsum 0.0100 0.0141 0.1255 0.7580 <0.0001 0.0813 Mulch*Drip 0.0867 0.2468 0.6483 0.2980 0.9517 0.2321 Mulch*Gypsum 0.3790 0.5189 0.3846 0.8182 0.7458 0.7016 Drip*Gypsum 0.0334 0.3222 0.1820 0.0801 0.4656 0.7128 Mulch*Drip*Gypsum 0.6004 0.6752 0.3817 0.4819 0.6458 0.6640 z Letters indicate LSD between treatment combinations y Hart et al ., 2006 x Erb, 1998

Table 6. Soil heat unit accumulation, and P. cinnamomi hyphal growth on sealed and buried PDA petri plates, beneath sawdust and black woven plastic geotextile weed mat mulch. Degree day P. cinnamomi colony area Date Mulch accumulationz (mm2) 9‐Aug to 16‐Aug 2012 Sawdust 51.7 ± 1.13 a 2978 ± 92.3 a Weed Mat 64.3 ± 4.12 b 3706 ± 158.7 b

18‐Apr to 28‐Apr 2013 Sawdust 97.2 ± 2.63 a 544 ± 16.7 a Weed Mat 111.7 ± 2.02 b 730 ± 26.2 b

11‐July to 19‐July 2013 Sawdust 114.3 ± 0.96 a 2593 ± 79.8 a Weed Mat 117.8 ± 1.06 b 2831 ± 89.8 b z o Degree days were calculated above 9 C baseline (Zentmyer et al., 1976)

Table 7. Soil pH and Mehlich III extractable cations from plots treated with all combinations of gypsum, drip irrigation line placement, and mulch type. pH Ca Mg K Date Mulch Drip Gypsum 1:1mg / kg mg / kg mg / kg June 2012 Weed Mat Wide Y 5.07 ± 0.04 4997 ± 563 592 ± 18 201 ± 10 Weed Mat Wide N 5.40 ± 0.11 2173 ± 78 692 ± 23 200 ± 5 Weed Mat Narrow Y 4.07 ± 0.03 4330 ± 1083 499 ± 44 179 ± 14 Weed Mat Narrow N 4.05 ± 0.02 1746 ± 102 624 ± 29 177 ± 9 Sawdust Wide Y 4.88 ± 0.03 4836 ± 590 527 ± 23 219 ± 7 Sawdust Wide N 5.25 ± 0.08 2118 ± 82 654 ± 36 216 ± 17 Sawdust Narrow Y 4.07 ± 0.05 3935 ± 574 529 ± 40 186 ± 10 Sawdust Narrow N 4.08 ± 0.09 1835 ± 99 609 ± 24 190 ± 11 Sawdust Narrow Fungicide 4.27 ± 0.21 1953 ± 109 650 ± 43 199 ± 9

July 2013 Weed Mat Wide Y 5.35 ± 0.12 2973 ± 142 643 ± 35 182 ± 10 Weed Mat Wide N 5.70 ± 0.12 2284 ± 79 780 ± 23 206 ± 17 Weed Mat Narrow Y 4.88 ± 0.15 2424 ± 63 669 ± 35 160 ± 4 Weed Mat Narrow N 5.17 ± 0.21 2042 ± 41 717 ± 15 162 ± 5 Sawdust Wide Y 5.25 ± 0.07 3260 ± 241 657 ± 27 199 ± 9 Sawdust Wide N 5.68 ± 0.09 2300 ± 62 764 ± 27 204 ± 9 Sawdust Narrow Y 4.73 ± 0.09 2230 ± 132 603 ± 26 131 ± 8 Sawdust Narrow N 4.87 ± 0.14 1983 ± 43 693 ± 17 146 ± 8 Sawdust Narrow Fungicide 4.80 ± 0.13 2059 ± 76 707 ± 37 133 ± 3

April 2014 Weed Mat Wide Y 5.42 ± 0.19 2637 ± 117 553 ± 27 156 ± 20 Weed Mat Wide N 5.92 ± 0.07 2098 ± 50 746 ± 17 168 ± 9 Weed Mat Narrow Y 5.87 ± 0.14 2229 ± 103 669 ± 18 113 ± 7 Weed Mat Narrow N 6.00 ± 0.12 2169 ± 83 804 ± 23 137 ± 12 Sawdust Wide Y 5.45 ± 0.07 2735 ± 142 447 ± 36 148 ± 6 Sawdust Wide N 5.77 ± 0.10 2012 ± 94 710 ± 24 160 ± 16 Sawdust Narrow Y 5.78 ± 0.09 2210 ± 32 679 ± 888± 7 Sawdust Narrow N 5.72 ± 0.10 1980 ± 70 743 ± 30 94 ± 7 Sawdust Narrow Fungicide 5.73 ± 0.09 1912 ± 90 707 ± 52 86 ± 8

35% Weed mat 30%

Sawdust 25% Incidence

20%

15% Infection

10% Root 5%

0% Jul 2012 Dec 2012 Mar 2013 Jun 2013 Sep 2013 Dec 2013

Figure 1. Differences in root infection incidence between geotextile weed mat and sawdust mulch at five sampling dates over two growing seasons. The December 2012 and 2013 samples were randomly sampled from whole excavated root balls. At all other dates, a single core sample was taken from one rootball per plot, and root sections were randomly sampled from the core. Data are in Table 3.

35% Narrow 30% Wide 25% Incidence

20%

15% Infection 10% Root 5%

0% Jul 2012 Dec 2012 Mar 2013 Jun 2013 Sep 2013 Dec 2013

Figure 2. Influence of drip irrigation line spacing on P. cinnamomi root infection incidence over two growing seasons. Narrow lines were adjacent to the crown of the plants, and wide lines were placed 20 cm on either side of the crown. The December 2012 and 2013 samples were randomly sampled from whole excavated root balls. At all other dates, a single core sample was taken from one rootball per plot, and root sections were randomly sampled from the core. Data are in Table 3.

35% No Gypsum 30% Gypsum 25% Incidence

20%

15% Infection

10% Root

0% Jul 2012 Dec 2012 Mar 2013 Jun 2013 Sep 2013 Dec 2013

Figure 3. Influence of gypsum amendment on P. cinnamomi root infection incidence over two growing seasons. The December 2012 and 2013 samples were randomly sampled from whole excavated root balls. At all other dates, a single core sample was taken from one rootball per plot, and root sections were randomly sampled from the core. Data are in Table 3.

50% Narrow No Gypsum Weed Mat 45% Wide Gypsum Sawdust 40% Fungicide Control ‐ Narrow, No Gyp, Sawdust 35% 30% Incidence

25% 20% Infection

15%

Root 10% 5% 0% Jul 2012 Dec 2012 Mar 2013 Jun 2013 Sep 2013 Dec 2013

400 Leaf Stem Root 200 grams

‐200 Narrow, No Gypsum, Wide, Gypsum, Sawdust Fungicide Control ‐ Weed Mat Narrow, No Gyp, Sawdust

Figure 4. Root infection incidence and plant biomass after two growing seasons of cultural treatments with the consistently highest and lowest root infection incidence, and the fungicide control. The December 2012 and 2013 samples were randomly sampled from whole excavated root balls. At all other dates, a single core sample was taken from one rootball per plot, and root sections were randomly sampled from the core. Data are in Table 3.Error bars indicate standard error (n=6).

30 A Sawdust 28 Weed Mat

20 9‐Aug 10‐Aug 11‐Aug 12‐Aug 13‐Aug 14‐Aug 15‐Aug 16‐Aug

22 B 20 C o

16 Temperature 14 Soil 12

10 18‐Apr 20‐Apr 22‐Apr 24‐Apr 26‐Apr 28‐Apr 30

28 C

20 11‐Jul 13‐Jul 15‐Jul 17‐Jul 19‐Jul

Figure 5. Diurnal soil temperature fluctuation at 15‐cm beneath either sawdust or woven black geotextile weed mat. Three dataloggers were installed per mulch type. A:9‐Aug to 16‐Aug 2012 B:18‐Apr to 28‐Apr 2013 C: 11‐Jul to 19‐Jul 2013.

40 a Dual driplines adjacent to crown A 35 Dual driplines 20‐cm away from crown 30 b a 25

20 b 15

0 7/12/2013 8‐7‐2013 35 aa

30 Weed Mat Mulch Sawdust Mulch B

25 a a Moisture 20 Soil

10 Volumetric

% 5

0 7/12/2013 8‐7‐2013 35 aa

30 No Gypsum Gypsum C

25 a a 20

0 7/12/2013 8‐7‐2013 Figure 6. Mean main effects of (A) drip irrigation line placement, (B) mulch type, and (C) gypsum incorporation on volumetric soil moisture in the center of raised beds1‐hr after a 15‐min irrigation event on 12‐Jul. and 8‐Aug. 2013. Error bars indicate standard error (n=6)

350 5‐Months 16‐Months a 300 L) 250 / ab

(mg

2+ 200 Ca

bc bcd 150 Solution cd 100 Soil cd

50 d cd

0 Gypsum Wide No Gypsum Gypsum No Gypsum Dripline Wide Dripline Narrow Dripline Narrow Dripline

Figure 7. Calcium levels in soil solution at the plant crown 5 and 16 months after planting. Gypsum was incorporated in a 30‐cm band at 10 tons / acre (22,420 kg/ha) in‐band rate (2,242 kg/ha whole‐field rate). Drip irrigation lines were placed either adjacent to the crown, or 20‐cm on either side of the crown. Letters indicate Fisher’s LSD between all treatments across both sampling dates. Error bars indicate standard error (n=6)

Chapter Five: Cation‐specific salinity effects from calcium and potassium on highbush blueberry

John R. Yeo, Dan M. Sullivan, and Dave R. Bryla

Abstract

‐1 High gypsum (CaSO4) application rates increase soil electrical conductivity (EC) > 2.0 dS∙m , which may pose threat of salt damage to highbush blueberry (Vaccinium corymbosum L.). Nitrogen and potassium fertilizer salts cause stunted growth, leaf necrosis, and plant death of highbush

blueberry when applied beyond EC 2.0 dS/m, however the risk of plant damage from high CaSO4 rates has not been studied. Gypsum provides some suppression of Phytophthora root rot disease on bluberry, and very high rates are required for disease suppression. We compared the

effects of increasing soil salinity with either gypsum or potassium sulfate (K2SO4) on highbush blueberry cultivar ‘Draper’ plant biomass accumulation, leaf nutrient concentration, soil solution, and Mehlich III extractable soil cations.. Plants were grown in pots in loam soil in a greenhouse trial for six months. Soil EC was monitored periodically using Rhizon mini lysimeters and adjusted to target EC levels with dilute salt solutions as needed. Plants generally exhibited linear decreases in aboveground biomass and root biomass in response to elevated EC. Compared to the lowest EC treatment (0.3 dS m‐1), aboveground biomass at the highest soil solution EC concentration (2.6 dS m‐1) was 10 to 15% lower with gypsum and 60 to 70% lower with potassium sulfate. Similarly, root dry weight was unaffected by elevated EC from gypsum, but it was reduced by 70 to 80% at the highest rate of potassium sulfate addition. Only plants exposed to high EC from potassium sulfate exhibited marginal leaf necrosis. Leaf necrosis was absent across all gypsum rates. The K concentration in leaf tissue increased from 0.6 to 2.5 % as

‐1 EC increased from 0.3 to 2.6 dS m with K2SO4. With gypsum, Ca concentration was similar across all rates, ranging from 0.7 to 0.8 %. Potassium sulfate addition decreased both leaf Ca and Mg concentration, but gypsum addition had no effect on leaf K or Mg concentration. The differential influence of K and Ca at equivalent EC values indicates the effects of soil salinity on highbush blueberry are ion‐specific effects, rather than osmotic stress. Potassium accumulation in plants was associated with reduced growth, and leaf Ca and Mg concentrations that approached deficiency levels. Although gypsum application dramatically increased Ca in soil solution, Ca did not accumulate in leaves, and had minimal effects on plant growth. Findings reported here suggest that blueberry can tolerate higher EC than the reported salt damage threshold (2.0 dS m‐1) when Ca is the major cation contributing to EC. In contrast, the EC threshold for plant damage may be lower than 2.0 dS m‐1 when K is the dominant cation. Findings suggest that soluble Ca can be increased in soil solution to concentrations that will inhibit P. cinnamomi root rot disease without causing severe salt damage to plants.

Introduction Control of Phytophthora spp. in horticultural crops can be partially accomplished by soil application of high levels of Ca2+ suppressive to asexual motile zoospores, thus disrupting the rapid repeating disease cycle (Zentmyer, 1980). Gypsum (Ca2SO4∙2H2O) is a soluble salt commonly applied in agriculture for enhancing Ca and S levels in soil, and provides suppression of Phytophthora root rot on highbush blueberry (Yeo et al., 2014a; 2014b). Rates of gypsum application for suppressing Phytophthora root rot disease in research trials are incredibly high compared to typical rates applied for supplemental Ca or S nutrition, and range from 1% to 5% of total soil volume (Messenger et al., 2000a; 2000b; Yeo et al., 2014a) or 13.5 Mg∙ha‐1 (Maloney et al., 2005; Pinkerton et al., 2009) to 22.4 Mg∙ha‐1 (Yeo et al., 2014b).

Salinity damage in blueberry is reported to occur at soil electrical conductivity (EC) thresholds of ≈1.5‐ 2.0 dS∙m‐1, which is similar to many salt‐sensitive plants (Bryla et al., 2014). High soil EC reduces growth and yield in blueberry, and, in extreme cases, results in leaf necrosis and plant death (Bryla and Machado, 2011; Bryla and Vargas, 2014; Bryla et al., 2014; Haby et al., 1986; Townsend, 1973; Wright et al., 1992; 1994). Salinity in cultivated blueberry soil can occur from applications of ammonium fertilizer, composts containing high K concentrations, and gypsum (Bryla et al., 2011; Sullivan et al., 2014; Yeo et al., 2014a). Salinity damage thresholds are based only on EC, and do not account for the ion‐specific

+ + + ‐ effects of salinity (Grattan and Grieve, 1999). Blueberry sensitivity to salinity from Na , NH4 , and K , Cl is well documented, however sensitivity to salinity from Ca2+ has not been well studied to determine if EC is a reliable predictor of salt stress when the salinity complex is dominated by gypsum. Application of high quantities of gypsum on blueberry for Phytophthora suppression increases soil EC above recommended limits for blueberry, and the objective of this study was to determine if high EC from gypsum is detrimental to blueberry plant growth, compared to high EC from K2SO4.

Materials and Methods

The effects of gypsum or K2SO4 on highbush blueberry and nutrient uptake were studied in a potted greenhouse experiment using Willamette silt loam soil (fine‐silty, mixed, mesic Pachic Ultic Argixeroll) collected from the southeast corner of the OSU North Willamette Research and Extension Center, Aurora, OR. A non‐replicated laboratory bench trial indicated soil electrical conductivity (EC) did not increase beyond approximately 2500 mg gypsum / kg soil, at which point soil EC was 2.6 dS/cm, and

100 equivalent mass additions of either gypsum or K2SO4 below this level resulted in approximately ‐1 equivalent EC. Ten rates of either gypsum (Ca2SO4●2H2O; molecular weight 172.17 g mol ) or potassium ‐1 sulfate (K2SO4; molecular weight 174.26 g mol ) ranging from 0 to 2500 mg salt / kg soil were added to the soil and mixed in a cement mixer, and 5‐cm diameter nursery plugs of highbush blueberry cultivar ‘Draper’ were transplanted into 2.6 L, 15‐cm diameter pots containing the soil‐salt mixes on 8‐Oct 2012. Pots were arranged in a CRD on a greenhouse bench with three replications per salt treatment. Plants were grown in a greenhouse maintained at 25oC, with supplemental HPS lighting providing 16‐hr daylength during the winter. Silver reflective shadecloth (60% shade) was placed over the greenhouse in spring and summer. Nitrogen fertilizer was applied every two weeks with urea dissolved in water, supplying 10 mg N plant‐1 wk‐1.

Hydrophyllic porous plastic wire reinforced 0.15 micron pore size mini‐lysimeters 2.5 mm diameter 10 cm length (Rhizon Soil Moisture Samplers, Eijkelkamp Agrisearch Equipment, The Netherlands) were installed in each pot and equipped with 10 mL syringes to collect soil solution samples for EC monitoring for the duration of the experiment. Syringe plungers were withdrawn under vacuum, and wooden blocks were inserted between the plunger and syringe body to maintain vacuum until >3 mL soil solution was collected. Soil solution samples were collected approximately one hour after hand irrigation, and EC was measured with a VWR Symphoy SB80 PC combination pH and EC meter and VWR Symphony model 11388‐382 four‐cell conductivity probe.

Plants were maintained in saucers and judiciously hand‐watered in an attempt to avoid leaching salts. However, after 3.5 months, EC in pots had decreased up to 50% in some pots with high variability, and a salinity monitoring and replenishment program was subsequently implemented to maintain EC levels. Target EC values were established in ten equal increments between 0.3 and 2.6 dS m‐1, which were approximately equal to EC values after soil mixing, with the 0.3 dS m‐1 rate corresponding to zero initial salt addition, and 2.6 dS m‐1 corresponding to the maximum initial rate of each salt. EC was measured weekly starting three months after planting, and when EC dropped 10% below the target value, dilute saline solutions of either gypsum or K2SO4 mixed to target EC levels were added to the soil. Pots were salinized, followed by hand irrigation, and soil solution samples were collected one hour later to determine if further EC adjustment was needed. In the event salinity exceeded the target value by 10%, the pot was leached with water until EC was within 10% of the target value.

101

All plants were destructively harvested 8.5 months after planting, and soil and potting media were washed from the roots. Plants were partitioned into stem, leaf, and root components which were dried and weighed. Stems and leaves were ground, homogenized, and subsamples were digested with 67‐70% w/w nitric acid in an Anton Paar Multiwave 3000 microwave and analyzed for Ca, K, Mg, P, and S concentration by ICP‐AES using a Perkin Elmer Optima 4300 instrument. Subsamples of dried and ground stem and leaf tissue were analyzed for C and N concentration by dry combustion using a Leco Truspec analyzer. Soil solution samples were collected from each pot shortly before harvest, and were analyzed for Ca, K, Mg, and S by ICP‐AES. All leaf and soil solution analysis was performed at the USDA‐ ARS HCRL, Corvallis, OR. Soil samples were collected from each pot, dried and ground, and analyzed for Mehlich III extractable Ca, K, Mg, and S by Brookside Laboratories, New Bremen, OH (Mallarino 2003).

Statistical Analysis Responses of plant biomass, nutrient status, and soil nutrient status to increasing EC were modeled separately for gypsum and K2SO4 using the REG procedure in SAS 9.2. Linear models were fitted to each response variable, so that the slope response could be compared between CaSO4 and K2SO4. Furthermore, most of the data appeared somewhat linear, and there was no biological justification to select nonlinear models.

To determine if responses differed between the two salt sources, Interactions of the slope response between gypsum and K2SO4 were compared using the GLM procedure in SAS 9.2. Model terms included

EC as a numerical variable, salt source (K2SO4 or CaSO4) as categorical, and the salt source x EC interaction (Table 1).

Results

High soil EC from both gypsum and K2SO4 reduced vegetative biomass compared to plants grown in non‐ salinized soil. However, plants grown with high EC from gypsum had more vegetative biomass compared to plants grown in high EC from K2SO4. Vegetative biomass decreased by 5.3 g from 17.58 g to 12.22 g as ‐1 EC from gypsum salinization increased from 0.30 to 2.6 dS m . Over the same EC range from K2SO4 salinization, vegetative biomass decreased by 8.53 g from 13.95 to 5.42 g.

Root biomass did not change significantly with increasing EC from gypsum, however decreased

‐1 significantly from 8.36 to 3.12 g as soil was salinized with K2SO4 between 0.30 to 2.6 dS m (Table 1;

102

Table 2; Figure 1). Plants subjected to the high EC levels from K2SO4 exhibited marginal leaf necrosis and leaf senescence, while the plants subjected to high EC from CaSO4 lacked necrotic symptoms.

Soluble cations in soil solution collected from the mini‐lysimeters responded differently to the different

‐1 ‐1 2+ salt additions. Increasing soil EC with CaSO4 from 0.30 dS m to 2.6 dS m increased soluble Ca in soil ‐1 ‐1 solution by 25.01 meq L from 1.15 to 26.17 meq L . Increasing soil EC with K2SO4 over the same range increased K+ in soil solution by 12.33 meq L‐1 from 0.12 to 12.45 meq L‐1. The impact of Ca addition on K and Mg concentration in soil solution differed from the impact of K addition on Ca and Mg in soil solution. Addition of K increased both Ca and Mg in the solution pool, however Ca addition only increased solution Mg concentration, and had no significant influence on solution K concentration (Table 1).

While soil solution ionic distribution tends to be transient and rapidly altered, extractable ions represent the exchangeable and solid phase reservoirs of soil ions. Changes in extractable ion distribution occur more slowly than changes in the soil solution, and represent more permanent alteration of the soil

‐1 ‐1 chemistry. Increasing soil EC with CaSO4 from 0.30 dS m to 2.6 dS m increased Mehlich III extractable 2+ ‐1 ‐1 Ca by 3.3 cmol kg from 11.0 to 14.3 cmol kg . Increasing soil EC with K2SO4 over the same salinity + ‐1 ‐1 range increased extractable K by 4.0 cmol kg from 0.7 to 4.7 cmol kg . Salinization with Ca significantly reduced both extractable K and Mg, and salinization with K significantly reduced extractable Ca and Mg.

Leaf K concentration increased dramatically more with K2SO4 addition, than did leaf Ca concentration with CaSO4 addition. Leaf K concentration increased by 1.84 % from 0.62 to 2.46 % as EC increased from ‐1 0.30 to 2.6 dS m with K2SO4. However leaf Ca concentration only increased by 0.10 % from 0.67 to 0.77

% as EC increased over the same range with CaSO4 (Table 1; Figure 2). Increasing salinity from 0.30 to 2.6 ‐1 dS m with K2SO4 reduced leaf concentration of Ca by 0.52% from 0.89 to 0.37 %, and reduced leaf Mg concentration by 0.13% from 0.31 to 0.18 %. However, increasing salinity over the same EC range with

CaSO4 had no effect on leaf K concentration, or leaf Mg concentration.

Although the relative sink partitioning of K or Ca between the Mehlich III extract, soil solution, and leaf tissue was different between the K 2SO4 and CaSO4 application treatments, distribution of SO4 in these pools had less variation between the two salt treatments. Soil solution S concentration increased from

‐1 ‐1 ‐1 1.7 to 25.4 meq L with CaSO4 salinization between 0.30 to 2.6 dS m , and from 1.8 to 26.9 meq L with

103 salinization by K2SO4. Although soil solution SO4 between K2SO4, and CaSO4 did not differ, plants treated with K2SO4 had higher leaf S concentration than plants treated with CaSO4. Soil extractable SO4 was higher with CaSO4 salinization than with K2SO4 salinization.

Discussion

The results of this study indicate that effects of salinity from K2SO4 salinization on blueberry differ greatly than the effects of CaSO4 salinization. Addition of both salts reduced shoot dry mass, although the reduction was greater with K2SO4 than CaSO4. Root dry mass decreased with addition of K2SO4, but not with CaSO4 salinization.

Studies differentiating specific‐ion toxicity from reduced osmotic potential have not been conducted for blueberry, however NaCl damage on Vaccinium myrtillus appears to be caused by specific Na+ ion toxicity rather than osmotic limitation on water uptake (Tahkokorpi et al., 2012). The salt effect of K2SO4 in this experiment reduced plant biomass much more dramatically than the salt effect of gypsum,

+ ‐ 2+ indicating the effects of salinity are ion‐specific. Blueberries are highly sensitive to Na and Cl , but Ca 2‐ and SO4 appear to have little impact plant growth (Eaton et al., 1999; Muralitharan et al., 1992; Patten et al, 1989). Few studies have evaluated salinity effects from K+ specifically, however high K+ accumulation in tissue and subsequent biomass reduction in this study appears to mimic non‐ exclusionary uptake of Na+.

Leaf tissue concentration response to added soil K was much greater than to added Ca. Whereas leaf Ca concentration did not increase with CaSO4 application, K leaf concentration increased from 0.62 to 2.46 ‐1 % as EC as K2SO4 salinization increased between 0.30 to 2.6 dS m greatly exceeding levels considered excessive beyond 0.9 % (Hansen, 1996). Increased levels of calcium in soil rarely correlate with plant Ca uptake, however K concentration in leaves responds strongly to soil K levels (Levula et al., 2000; Sanderson et al., 1996; Townsend, 1973). Leaf Mg2+ and Ca2+ concentration did not reach deficiency levels in this study K2SO4 salinization, however leaf concentration of both decreased with trend approaching deficiency levels. Additionally, K2SO4 salinization decreased plant biomass for much lower Mg and Ca total uptake.

104

Addition of K or Ca to the soil altered the distribution of Mehlich III extractable cations due to replacement on the soil cation exchange. Cation exchange in the soil depends both on the lyotropic

3+ + 2+ 2+ + + + series predicting strength of adsorption where Al > H > Ca > Mg > K = NH4 > Na , and on ionic composition of the soil solution. CaSO4 salinization decreased soil extractable K which follows the lyotropic series. Salinization with K2SO4 decreased extractable Ca which does not follow the lyotropic series, and the exchange was likely controlled by high soil solution K (Table 1). Both CaSO4 and K2SO4 salinization decreased extractable Mg. Thus, soil salinization with high K compost, or gypsum can have a long term effect on Mg availability. Growth suppressive effects of salinization with K2SO4 were not, however, attributed to Mg deficiency in this study, as leaf tissue concentration did not reach critical deficiency levels.

Mehlich‐III extractable K+ and Mg2+ decreased with gypsum application due to higher affinity for Ca2+ adsorption on the soil cation exchange based on the lyotropic series (Lindsay, 1979). However only Mg2+ increased in soil solution, as K+ is more mobile and likely leached from the soil solution pool. Although Ca2+ and Mg2+ are more strongly adsorbed to cation exchange sites than K+, Mehlich‐III extractable Ca2+

2+ + and Mg decreased with K2SO4 addition due to high K concentration in solution altering equilibrium between soil solution and the cation exchange (Lindsay, 1979).

The primary implication of this research, is that EC levels do not accurately represent salinity thresholds for blueberries, as the salinity thresholds differ depending on ionic composition of the soil solution. The

‐2 effects of salinity ranged from highly deleterious with K2SO4, to benign with CaSO4. The SO4 anion has been shown to have little damaging effect on blueberry, indicating the difference in salt effect between

K2SO4 and CaSO4 addition is due to the different cations (Muralitharan et al., 1992). Plants did not accumulate large quantities of Ca in leaf tissue when CaSO4 was added, but accumulated levels of K in leaf tissue from K2SO4 addition were substantially higher than thresholds considered excessive. Calcium moves into plant roots primarily by mass flow, however the excessive K uptake suggests a more active uptake mechanism that is not downregulated when K approaches toxicity. Although K is typically not applied to phytotoxic levels, some manure‐based composts can supply excessive K if the compost is applied for its N value. Blueberry producers may want to deliberately apply large quantities of gypsum for Phytophthora suppression, and this research indicates there is little risk of plant injury.

105

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Yeo, J.R., D.M. Sullivan, J. Weiland, and D.R. Bryla. 2014b. Influence of gypsum, mulch, and irrigation line placement on Phytophthora cinnamomi root rot disease of blueberry in a 2‐year field study. Chapter 2. in Cultural controls for suppressing Phytophthora cinnamomi root rot disease of highbush blueberry. Ph.D. Dissertation. Oregon State University.

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Table 1. Relationships between electrical conductivity (EC) and plant biomass, nutrient status, and soil nutrient status vary between gypsum and K2SO4. Gypsum Potassium sulfate Salt x EC Interaction Response Variable Slope p‐value R‐square Relationship Slope p‐value R‐square Relationship p‐value Plant Biomass (g) Vegetative ‐1.5944 0.0279 0.16 Negative ‐4.4429 <0.0001 0.61 Negative 0.0045 Root ‐0.6148 0.1063 0.09 ns ‐2.2304 <0.0001 0.52 Negative 0.0045

Leaf Concentration N 0.1112 0.0007 0.34 Positive 0.1674 <0.0001 0.62 Positive 0.1483 (%) Ca 0.0316 0.0928 0.10 ns ‐0.1729 <0.0001 0.56 Negative <0.0001 K 0.0219 0.0558 0.12 ns 0.9134 <0.0001 0.87 Positive <0.0001 Mg ‐0.0089 0.1762 0.06 ns ‐0.0351 0.0045 0.25 Negative 0.0501 P ‐0.0008 0.8349 0.00 ns ‐0.0115 0.0908 0.10 ns 0.1591 S 0.0344 0.0547 0.13 ns 0.0877 0.0008 0.34 Positive 0.0704

Soil Solution Ca 10.9463 <0.0001 0.99 Positive 3.4018 <0.0001 0.90 Positive <0.0001 Concentration (meq K 0.0584 0.1648 0.07 ns 5.2342 <0.0001 0.94 Positive <0.0001 / L) Mg 0.0639 <0.0001 0.88 Positive 0.0688 <0.0001 0.97 Positive 0.3472 S 9.7066 <0.0001 0.88 Positive 10.4440 <0.0001 0.97 Positive 0.3472

Mehlich III Ca 1.2128 <0.0001 0.43 Positive ‐1.1688 <0.0001 0.52 Negative <0.0001 Extractable (cmol / Mg ‐1.0264 <0.0001 0.75 Negative ‐0.7443 <0.0001 0.56 Negative 0.0999 Kg soil) K ‐0.0506 0.0001 0.42 Negative 1.8727 <0.0001 0.95 Positive <0.0001 Na ‐0.0772 <0.0001 0.59 Negative ‐0.0534 <0.0001 0.45 Negative 0.1504 S 6.3199 0.0009 0.33 Positive 4.6318 <0.0001 0.47 Positive 0.3897

Soil pH 1:1 water ‐0.1542 0.0061 0.239 Negative ‐0.2443 <0.0001 0.48 Negative 0.2063

109

Gypsum 25 Potassium Sulfate Linear (Gypsum) 20 Linear (Potassium Sulfate) (g)

15 Mass

10 Vegetative 5

0 00.511.522.53 EC (dS/m)

10 9 8 7 (g) 6 5 Mass

4 Root 3 2 1 0 0 0.5 1 1.5 2 2.5 3 EC (dS/m)

Figure 1. Influence of increasing EC with either gypsum or K2SO4 on vegetative biomass of ‘Draper’ highbush blueberry.

110

1.0 0.9 0.8 (%) 0.7 0.6 0.5 concentration

0.4 Ca 0.3 Leaf 0.2 0.1 0.0 0 0.5 1 1.5 2 2.5 3 EC (dS/m) 30 Gypsum ) 1 ‐ Potassium Sulfate L

25 Gypsum (Linear) (meq Potassium Sulfate (Linear) 20

15 concentration 2+ Ca 10 Solution

5 Soil 0 00.511.522.53 EC (dS/m)) 18 ) 1

‐ 16 Kg 14

(cmol 12 2+ Ca 10 8 extractable 6 III

Mehlich 2 0 00.511.522.53 EC (dS/m)

Figure 2. Calcium leaf concentration, soil solution concentration, and exchangeable Ca2+ with increasing salinity added with either gypsum or K2SO4.

111

3.5

3.0

(%) 2.5

2.0

1.5 concentration

1.0 Leaf

0.5

0.0 0 0.5 1 1.5 2 2.5 3 EC (dS/m)

14 Gypsum

) Potassium Sulfate 1 ‐ L

12 Gypsum (Linear) Potassium Sulfate (Linear) (meq 10

concentration 6 + K

4 Solution 2 Soil

0 00.511.522.53 EC (dS/m) 6 ) 1 ‐ 5 Kg

(cmol 4 + K

3 extractable 2 III

1 Mehlich

0 00.511.522.53 EC (dS/m) Figure 3. Potassium leaf concentration, soil solution concentration, and exchangeable K+ with increasing salinity added with either gypsum or K2SO4.

112

0.3 (%)

0.2 concentration

0.1 Leaf

0.0 0 0.5 1 1.5 2 2.5 3 EC (dS/m) 0.25 Gypsum ) 1 ‐

L Potassium Sulfate

0.20 Gypsum (Linear) (meq Potassium Sulfate (Linear)

0.15 concentration

2+ 0.10 Mg

0.05 Solution

Soil 0.00 00.511.522.53 EC (dS/m) 7 ) 1 ‐ 6

(cmolKg 5 2+ Mg 4

3 extractable

III

1 Mehlich

0 00.511.522.53 EC (dS/m))

Figure 4. Magnesium leaf concentration, soil solution concentration, and exchangeable Mg2+ with increasing salinity added with either gypsum or K2SO4.

113

0.5

0.4 (%)

0.3

concentration 0.2

Leaf 0.1

0.0 0 0.5 1 1.5 2 2.5 3 EC (dS/m) 35 Gypsum ) 1

‐ Potassium Sulfate

L 30 Gypsum (Linear) (meq 25 Potassium Sulfate (Linear)

concentration 15

10 Solution

5 Soil

0 00.511.522.53 EC (dS/m) 20 ) 1 ‐

15 (cmolKg

S ‐ 4 SO

10 extractable

III 5 Mehlich

0 00.511.522.53 EC (dS/m)

Figure 5. S leaf concentration, soil solution SO4‐S concentration, and Mehlich III extractable SO4‐S with increasing salinity added with either gypsum or K2SO4.

Chapter Six: General Conclusion

John R. Yeo

115

Phytophthora cinnamomi is a soilborne pathogen that causes root rot disease of highbush blueberry

(Vaccinium corymbosum L.). When new installations of susceptible blueberry cultivars are infected with

P. cinnamomi, plants generally fail to grow significant new tissue, greatly reducing yields over the life of the planting. Chemical fungicides are available for disease suppression including mefenoxam and phosphonate compounds. However these tools are not available for organically certified growers. The purpose of research described in this dissertation was to investigate non‐chemical control strategies for blueberry root rot disease, including cultivar selection, gypsum, organic amendments, mulch, and drip irrigation line placement.

Cultivar resistance to P. cinnamomi is a critical component of non‐chemical disease control. Eighteen cultivars and advanced breeding selections were evaluated for susceptibility or resistance in three greenhouse experiments. Cultivars varied widely in susceptibility, with Duke, Draper, Bluetta, Blue

Ribbon, Cargo, Last Call, Top Shelf, and Ventura exhibiting high levels of susceptibility to the disease.

More resistant cultivars included Legacy, Liberty, Aurora, Overtime, Reka, and Clockwork. By selecting cultivars with superior resistance, growers may avoid yield losses associated with the disease, enhancing production profitability. When consumer markets require cultivars with desirable fruit quality characteristics, but high susceptibility to Phytophthora, cultural controls and fungicides become essential components of profitable production in on infested sites.

Gypsum and organic amendment provide suppression of Phytophthora root rot in crops other than blueberry. Preliminary greenhouse screening trials were conducted using highly susceptible highbush blueberry cultivar ‘Draper’ evaluating gypsum in combination with organic amendments for root rot suppression. A single high rate of gypsum (5% v/v) suppressed disease in one of the experiments, and organic amendments (peat, sawdust, dairy solids compost, yard debris compost, composted bark with biosolids) provided no disease suppression in any of the three experiments. The effects of gypsum on disease suppression appeared variable, however could be beneficial for reducing root rot disease severity.

116

Adequate rates of gypsum amendment are required for successful disease suppression. One greenhouse experiment was performed to determine the minimum gypsum application rate required to maximize soluble Ca2+ in soil solution for. Soluble Ca2+ in soil solution followed a linear‐plateau model with increasing rates of gypsum. Rates of gypsum required for disease suppression occurred when Ca2+ was maximized (~454 mg Ca2+ L‐1 soil solution) with gypsum addition of 15.6 meq gypsum 100g‐1 soil.

Increasing gypsum addition beyond this point did not increase soil solution Ca2+, or provide additional disease suppression.

Greenhouse studies indicated gypsum may provide root rot disease control, and a field experiment was installed to investigate disease suppression over two growing seasons with highly susceptible cultivar

‘Draper’ grown on a site with clay loam soil, and inoculation with P. cinnamomi. Treatments included all combinations of gypsum incorporation (2,242 kg ha‐1 whole‐field rate, applied only in 30‐cm bands in the center of raised beds) or no gypsum, woven black geotextile weed mat or sawdust, and dual drip irrigation lines placed either adjacent to the plant crown or 20‐cm on either side of the crown.

Additionally a fungicide control treatment with sawdust mulch and irrigation lines adjacent to the crown provided a comparison for efficacy of the cultural treatments Mulch type had no significant effect on plant biomass after two years, however plants grown under sawdust tended to have slightly higher biomass and less root infection. The disease suppressive effect of gypsum amendment depended on the placement of irrigation lines. When drip irrigation lines were placed adjacent to the plant crown, Ca2+ was translocated away from the rootzone, negating any disease suppression. When drip irrigation lines were placed 20‐cm on either side of the crown, Ca2+ was translocated into the rootzone, which reduced root infection incidence and enhanced plant biomass. Despite significant increases in biomass with gypsum and widely‐placed irrigation lines, the conventional fungicide control treated plants had approximately twice as much biomass after two years.

Rates of gypsum required for disease suppression increase soil electrical conductivity (EC) beyond current recommended salinity guidelines for blueberries (> 2.0 mS/cm). The effects of salinity from gypsum were compared to effects of salinity from potassium sulfate in a greenhouse experiment with

117 plants not infected with Phytophthora. Salinity was added as either gypsum or potassium sulfate at ten equivalent EC levels ranging from 0.3 to 2.6 mS/cm. Plant biomass decreased significantly with increasing EC from potassium sulfate, however vegetative biomass only decreased slightly with increasing EC from gypsum, and root biomass was not affected by gypsum. Despite increasing soil EC with gypsum, the salt effect on reducing plant growth appeared minimal, indicating high rates of gypsum are not detrimental to blueberry.

An integrated control program is required for cultural suppression of blueberry root rot disease, including cultivar resistance, gypsum, widely‐spaced irrigation lines, and careful irrigation scheduling. On highly susceptible cultivar ‘Draper,’ wide irrigation lines and gypsum increased vegetative biomass, however growth was inferior to the chemical fungicide control treatment. Organic production of highly susceptible cultivars in the presence of P. cinnamomi is difficult, and may not produce equivalent yields to conventional production. With more resistant cultivars, cultural controls could have greater efficacy at improving yields in the presence of P. cinnamomi.

118 References

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Appendix 1. Nutrient concentration of ‘Draper’ highbush blueberry grown in different gypsum and organic matter amendments, either infected or non‐infected with P. cinnamomi root rot. N P S Ca Mg K g 100g‐1 g 1000g‐1 g 1000g‐1 g 1000g‐1 g 1000g‐1 g 1000g‐1 P. cinn. Infection Y N Y N Y N Y N Y N Y N Treatment† NA 1.18 0.88 1.12 0.87 0.94 0.74 2.7 2.7 0.70 0.67 4.8 4.3 PT 1.03 0.88 1.06 0.93 0.86 0.75 2.7 2.8 0.77 0.77 4.6 4.4 BSB 1.17 0.82 1.21 1.05 0.96 0.80 2.7 3.8 0.72 0.88 4.3 4.1 DS 1.16 0.87 1.17 0.95 0.99 0.73 2.7 2.7 0.70 0.74 4.6 4.4 SD 1.09 0.71 1.10 0.96 0.98 0.66 2.4 3.6 0.67 0.96 4.2 4.1 YDC 1.04 0.88 1.10 0.94 0.88 0.75 2.4 2.5 0.66 0.68 4.8 4.6 Gyp 0.89 0.93 0.91 0.87 1.72 1.62 4.2 4.0 0.81 0.82 4.4 4.6 Gyp + PT 0.99 0.90 1.01 0.94 1.52 1.52 3.7 4.3 0.77 0.74 4.7 4.4 Gyp + BSB 0.89 0.86 0.99 0.98 1.67 1.76 4.0 4.6 0.84 0.88 4.4 4.7 Gyp + DS 0.94 0.85 1.01 0.95 1.35 1.62 3.6 4.5 0.70 0.73 4.6 4.6 Gyp + SD 0.94 0.67 1.03 0.92 1.70 1.46 3.8 4.5 0.81 0.76 4.8 4.6 Gyp + YDC 1.00 0.85 1.07 0.84 1.59 1.75 3.9 4.5 0.81 0.80 4.8 4.9 Fungicide 0.92 1.03 1.03 1.00 0.86 0.80 2.9 3.2 0.83 0.76 4.1 4.3 †NA=no amendment; PT=peat; BSB=biosolids composted bark; DS=dairy solids; SD=sawdust; YDC=yard debris compost; Gyp=gypsum.

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Appendix 2. Vegetative and root biomass of ‘Draper’ highbush blueberry grown in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m‐1. Salt EC (dS m‐1) Stem mass (g) Leaf mass (g) Vegetative mass (g) Root mass (g) Gypsum 0.3 10.3 ± 0.81 7.3 ± 6.43 17.6 ± 0.78 8.4 ± 0.53 0.555 7.0 ± 0.75 5.6 ± 3.11 12.6 ± 1.28 6.0 ± 0.31 0.811 7.3 ± 0.43 4.8 ± 2.91 12.2 ± 0.89 5.4 ± 1.02 1.066 6.8 ± 1.28 5.3 ± 2.86 12.1 ± 1.89 5.9 ± 1.06 1.322 7.4 ± 0.20 6.1 ± 2.63 13.6 ± 0.94 5.8 ± 0.70 1.577 6.8 ± 0.90 5.3 ± 2.26 12.1 ± 1.90 6.0 ± 0.93 1.833 6.7 ± 0.55 5.1 ± 2.24 11.8 ± 1.58 5.6 ± 1.14 2.088 5.8 ± 0.97 5.4 ± 1.99 11.2 ± 2.23 4.5 ± 1.06 2.344 6.2 ± 1.36 4.8 ± 2.57 10.9 ± 2.14 6.1 ± 0.15 2.6 7.1 ± 1.15 5.2 ± 2.75 12.2 ± 1.83 5.9 ± 0.82

Potassium Sulfate 0.3 8.2 ± 1.23 5.8 ± 4.04 14.0 ± 1.17 8.3 ± 1.59 0.555 8.9 ± 0.75 7.0 ± 3.75 15.9 ± 1.12 6.9 ± 1.08 0.811 6.9 ± 2.14 4.9 ± 2.40 11.8 ± 3.03 5.8 ± 1.03 1.066 7.1 ± 1.30 5.8 ± 2.40 12.9 ± 2.20 5.7 ± 1.85 1.322 7.1 ± 0.45 5.0 ± 2.91 12.0 ± 1.02 5.7 ± 0.90 1.577 6.0 ± 0.34 4.2 ± 9.43 10.1 ± 0.39 5.6 ± 0.57 1.833 4.6 ± 1.42 3.3 ± 1.78 7.9 ± 3.05 3.3 ± 0.74 2.088 4.5 ± 0.37 3.4 ± 3.62 7.9 ± 0.62 3.2 ± 0.38 2.344 3.3 ± 0.36 2.1 ± 2.94 5.4 ± 0.96 3.1 ± 0.20 2.6 3.8 ± 0.61 1.7 ± 4.36 5.4 ± 0.81 3.1 ± 0.79

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Appendix 3. Total nutrient uptake of ‘Draper’ highbush blueberry grown in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m‐1. Salt EC (dS m‐1) N (g) P (mg) Ca (meq) K (meq) Mg (meq) S (meq) Gypsum 0.3 0.17 ± 0.007 14.6 ± 1.32 3.6 ± 0.20 1.9 ± 0.07 1.7 ± 0.07 1.3 ± 0.06 0.555 0.14 ± 0.014 13.0 ± 0.91 2.9 ± 0.40 1.5 ± 0.12 1.5 ± 0.17 1.3 ± 0.16 0.811 0.13 ± 0.004 10.8 ± 1.59 2.6 ± 0.40 1.3 ± 0.11 1.2 ± 0.15 1.2 ± 0.16 1.066 0.14 ± 0.023 11.2 ± 1.47 2.7 ± 0.43 1.4 ± 0.16 1.3 ± 0.12 1.5 ± 0.14 1.322 0.16 ± 0.008 11.4 ± 0.83 3.0 ± 0.34 1.6 ± 0.13 1.5 ± 0.15 1.7 ± 0.28 1.577 0.14 ± 0.023 11.8 ± 1.56 2.5 ± 0.27 1.4 ± 0.21 1.2 ± 0.14 1.5 ± 0.06 1.833 0.15 ± 0.021 8.7 ± 1.32 2.7 ± 0.43 1.4 ± 0.21 1.1 ± 0.17 1.5 ± 0.19 2.088 0.14 ± 0.023 9.3 ± 1.21 2.5 ± 0.42 1.4 ± 0.32 1.2 ± 0.23 1.4 ± 0.23 2.344 0.14 ± 0.031 9.9 ± 1.45 3.0 ± 0.52 1.4 ± 0.28 1.1 ± 0.16 1.3 ± 0.12 2.6 0.15 ± 0.025 9.8 ± 1.19 3.1 ± 0.46 1.4 ± 0.19 1.2 ± 0.09 1.4 ± 0.13

Potassium Sulfate 0.3 0.14 ± 0.004 14.4 ± 1.68 3.5 ± 0.44 1.7 ± 0.19 1.8 ± 0.20 1.3 ± 0.15 0.555 0.18 ± 0.006 15.3 ± 0.68 3.0 ± 0.17 2.0 ± 0.11 1.5 ± 0.10 1.4 ± 0.05 0.811 0.14 ± 0.033 10.2 ± 1.77 2.2 ± 0.41 1.8 ± 0.46 1.1 ± 0.25 1.3 ± 0.27 1.066 0.16 ± 0.025 11.2 ± 1.85 2.3 ± 0.54 2.4 ± 0.34 1.2 ± 0.18 1.4 ± 0.28 1.322 0.15 ± 0.017 11.4 ± 0.62 2.1 ± 0.06 2.5 ± 0.29 1.0 ± 0.06 1.5 ± 0.13 1.577 0.12 ± 0.007 8.9 ± 0.59 1.7 ± 0.04 2.5 ± 0.09 0.8 ± 0.05 1.3 ± 0.02 1.833 0.10 ± 0.041 5.6 ± 2.76 1.3 ± 0.63 2.3 ± 0.87 0.7 ± 0.26 0.9 ± 0.33 2.088 0.10 ± 0.006 5.9 ± 0.46 1.1 ± 0.08 2.2 ± 0.15 0.6 ± 0.05 1.1 ± 0.09 2.344 0.07 ± 0.016 4.1 ± 1.02 0.8 ± 0.11 1.9 ± 0.35 0.5 ± 0.09 0.7 ± 0.08 2.6 0.06 ± 0.005 4.0 ± 0.21 0.7 ± 0.14 1.7 ± 0.15 0.4 ± 0.09 0.8 ± 0.28

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Appendix 4. Leaf nutrient concentration of ‘Draper’ highbush blueberry grown in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m‐1. Salt EC (dS m‐1) N (g 100g‐1)P (g 100g‐1)S (g 100g‐1)Ca (g 100g‐1) K (g 100g‐1)Mg (g 100g‐1) Gypsum 0.3 1.51 ± 0.113 0.08 ± 0.005 0.20 ± 0.013 0.67 ± 0.036 0.48 ± 0.028 0.22 ± 0.009 0.555 1.65 ± 0.066 0.12 ± 0.012 0.27 ± 0.041 0.71 ± 0.072 0.53 ± 0.002 0.25 ± 0.023 0.811 1.73 ± 0.099 0.11 ± 0.001 0.28 ± 0.029 0.75 ± 0.039 0.51 ± 0.021 0.25 ± 0.010 1.066 1.69 ± 0.033 0.10 ± 0.009 0.35 ± 0.030 0.79 ± 0.017 0.57 ± 0.028 0.25 ± 0.016 1.322 1.71 ± 0.039 0.10 ± 0.004 0.33 ± 0.039 0.71 ± 0.044 0.55 ± 0.029 0.24 ± 0.014 1.577 1.74 ± 0.051 0.12 ± 0.006 0.38 ± 0.073 0.74 ± 0.046 0.55 ± 0.023 0.24 ± 0.020 1.833 1.83 ± 0.052 0.09 ± 0.005 0.35 ± 0.039 0.75 ± 0.064 0.59 ± 0.013 0.21 ± 0.016 2.088 1.86 ± 0.077 0.10 ± 0.012 0.31 ± 0.012 0.70 ± 0.036 0.54 ± 0.027 0.22 ± 0.010 2.344 1.80 ± 0.102 0.10 ± 0.006 0.31 ± 0.039 0.82 ± 0.009 0.58 ± 0.030 0.23 ± 0.015 2.6 1.80 ± 0.028 0.09 ± 0.005 0.30 ± 0.015 0.77 ± 0.026 0.53 ± 0.014 0.23 ± 0.019

Potassium Sulfate 0.3 1.57 ± 0.042 0.13 ± 0.034 0.28 ± 0.055 0.89 ± 0.154 0.62 ± 0.133 0.31 ± 0.055 0.555 1.71 ± 0.035 0.11 ± 0.008 0.23 ± 0.009 0.58 ± 0.016 0.54 ± 0.035 0.20 ± 0.002 0.811 1.73 ± 0.042 0.10 ± 0.008 0.29 ± 0.012 0.57 ± 0.012 0.75 ± 0.047 0.21 ± 0.002 1.066 1.84 ± 0.035 0.09 ± 0.001 0.28 ± 0.011 0.50 ± 0.023 0.98 ± 0.061 0.19 ± 0.004 1.322 1.92 ± 0.042 0.12 ± 0.012 0.35 ± 0.095 0.53 ± 0.085 1.17 ± 0.070 0.19 ± 0.034 1.577 1.90 ± 0.077 0.10 ± 0.003 0.36 ± 0.009 0.45 ± 0.020 1.58 ± 0.117 0.18 ± 0.007 1.833 1.89 ± 0.050 0.07 ± 0.010 0.36 ± 0.069 0.39 ± 0.022 1.98 ± 0.223 0.17 ± 0.007 2.088 1.89 ± 0.038 0.10 ± 0.003 0.35 ± 0.009 0.39 ± 0.024 1.79 ± 0.025 0.16 ± 0.005 2.344 2.11 ± 0.037 0.10 ± 0.014 0.42 ± 0.069 0.39 ± 0.008 2.44 ± 0.271 0.19 ± 0.013 2.6 1.94 ± 0.063 0.10 ± 0.026 0.48 ± 0.122 0.37 ± 0.050 2.46 ± 0.218 0.18 ± 0.032 Target Levelz 1.76‐2.0 >0.10 0.11‐0.16 0.41 ‐ 0.8 0.40‐0.70 0.13‐0.25 Deficiency Levely 1.50 0.1 0.05 0.3 0.4 0.2 Excess Levelx 2.30 0.60 na 1.00 0.90 na z Hart et al ., 2006 y Erb, 1988 x Hansen et al. , 1996

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Appendix 5. Soil solution nutrient concentration in soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m‐1. Salt EC (dS m‐1) Ca (meq L‐1) K (meq L‐1)Mg (meq L‐1)S (meq L‐1) Gypsum 0.3 1.15 0.23 0.10 0.05 0.01 0.00 1.7 0.26 0.555 2.84 0.04 0.11 0.07 0.04 0.00 5.5 0.38 0.811 4.67 0.38 0.12 0.03 0.05 0.00 8.0 0.72 1.066 8.30 0.49 0.11 0.02 0.08 0.01 11.9 0.86 1.322 10.54 0.33 0.22 0.04 0.10 0.01 15.3 1.49 1.577 13.55 0.44 0.18 0.02 0.13 0.01 19.8 0.89 1.833 17.27 0.52 0.42 0.26 0.14 0.01 21.2 0.92 2.088 19.55 0.95 0.18 0.03 0.13 0.00 20.2 0.13 2.344 21.69 0.82 0.21 0.04 0.13 0.02 20.1 2.40 2.6 26.17 0.08 0.16 0.03 0.17 0.00 25.4 0.76

Potassium Sulfate 0.3 1.01 0.15 0.12 0.01 0.01 0.00 1.8 0.30 0.555 2.35 0.43 0.59 0.02 0.03 0.00 4.8 0.65 0.811 3.88 0.19 1.36 0.20 0.05 0.00 7.6 0.60 1.066 4.70 0.60 2.47 0.15 0.06 0.01 9.6 0.76 1.322 5.37 0.57 3.61 0.28 0.08 0.01 12.6 0.91 1.577 5.69 0.04 5.19 0.35 0.10 0.00 14.6 0.66 1.833 7.82 0.46 5.70 0.06 0.11 0.01 17.0 1.41 2.088 8.04 0.45 8.27 0.24 0.14 0.01 20.5 0.84 2.344 8.64 0.83 9.72 0.87 0.15 0.01 22.8 1.18 2.6 8.67 0.27 12.45 0.41 0.18 0.01 26.9 1.34

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Appendix 6. Mehlich III extractable nutrients and pH of soil amended with either gypsum or potassium sulfate at levels of salinity ranging from 0.3 to 2.6 dS m‐1. Salt EC (dS m‐1) Ca (cmol Kg‐1)Mg (cmol Kg‐1)pHK (cmol Kg‐1) Na (cmol Kg‐1)S (cmol Kg‐1) 1:1 Gypsum 0.3 11.0 ± 0.52 5.9 ± 0.38 0.6 ± 0.02 0.4 ± 0.01 2.3 ± 0.65 5.27 ± 0.07 0.555 12.2 ± 0.91 5.9 ± 0.28 0.6 ± 0.03 0.4 ± 0.01 4.8 ± 0.62 5.13 ± 0.07 0.811 12.0 ± 0.44 5.7 ± 0.12 0.6 ± 0.01 0.4 ± 0.01 6.7 ± 1.52 5.00 ± 0.06 1.066 12.5 ± 1.09 5.6 ± 0.33 0.6 ± 0.06 0.4 ± 0.07 10.0 ± 5.31 5.03 ± 0.18 1.322 12.8 ± 0.16 5.0 ± 0.14 0.5 ± 0.00 0.3 ± 0.02 7.0 ± 1.37 5.03 ± 0.12 1.577 13.2 ± 0.48 4.8 ± 0.20 0.5 ± 0.03 0.3 ± 0.01 11.2 ± 1.63 4.97 ± 0.12 1.833 12.6 ± 0.24 4.1 ± 0.07 0.5 ± 0.01 0.2 ± 0.00 9.1 ± 2.26 5.03 ± 0.22 2.088 13.5 ± 1.00 4.2 ± 0.25 0.5 ± 0.02 0.3 ± 0.01 9.0 ± 3.31 5.07 ± 0.15 2.344 14.2 ± 0.68 3.7 ± 0.21 0.5 ± 0.00 0.2 ± 0.01 18.5 ± 8.91 4.77 ± 0.15 2.6 14.3 ± 0.73 4.1 ± 0.37 0.5 ± 0.02 0.3 ± 0.02 19.8 ± 7.56 4.80 ± 0.10

Potassium Sulfate 0.3 10.6 ± 0.46 5.9 ± 0.28 0.7 ± 0.03 0.3 ± 0.01 1.5 ± 0.16 5.53 ± 0.03 0.555 10.3 ± 0.28 5.6 ± 0.15 1.2 ± 0.07 0.4 ± 0.02 5.1 ± 0.91 5.20 ± 0.00 0.811 10.7 ± 0.47 5.9 ± 0.17 1.4 ± 0.07 0.3 ± 0.01 3.9 ± 0.45 5.27 ± 0.12 1.066 10.0 ± 0.08 5.3 ± 0.11 2.1 ± 0.04 0.3 ± 0.03 6.1 ± 1.55 5.17 ± 0.03 1.322 10.1 ± 0.13 5.2 ± 0.16 2.4 ± 0.10 0.3 ± 0.03 10.0 ± 3.83 5.10 ± 0.17 1.577 10.8 ± 0.49 5.7 ± 0.44 2.9 ± 0.28 0.3 ± 0.03 9.8 ± 2.03 5.00 ± 0.00 1.833 8.9 ± 0.80 4.8 ± 0.32 3.5 ± 0.07 0.3 ± 0.02 8.5 ± 1.84 4.97 ± 0.09 2.088 8.8 ± 0.43 4.5 ± 0.29 4.0 ± 0.29 0.2 ± 0.02 7.5 ± 1.01 5.23 ± 0.03 2.344 8.4 ± 0.20 4.4 ± 0.21 4.8 ± 0.28 0.3 ± 0.04 14.1 ± 4.55 4.67 ± 0.12 2.6 8.0 ± 0.73 4.3 ± 0.51 4.7 ± 0.26 0.2 ± 0.03 13.4 ± 1.99 4.90 ± 0.10