CLONAL PROPAGATION OF WALNUT ROOTSTOCK GENOTYPES FOR GENETIC IMPROVEMENT 2012

CLONAL PROPAGATION OF WALNUT ROOTSTOCK GENOTYPES FOR GENETIC IMPROVEMENT 2012 Chuck Leslie, Wes Hackett, Reid Robinson, Morgan McMahon, Joe Grant, Bruce Lampinen, Kathy Anderson, Bob Beede, Rick Buchner, Janet Caprile, Carolyn DeBuse, Rachel Elkins, Janine Hasey, Maryam Fazel, Nicolas Manterola, Dan Kluepfel, Greg Browne, Richard Evans, Mike McKenry, John Preece, Malli Aradhya, and Dianne Velasco ABSTRACT This year we produced more than 7300 liner-sized plantlets of 114 genotypes for use in greenhouse and field pest and disease resistance screening, growth in nurseries, and development of orchard trials. These included 3357 plants of 33 new Juglans microcarpa x Serr paradox genotypes and 675 plants of 54 additional J. cathayensis x Serr or Serr x J. cathayensis hybrid clones. Microshoots rooted and acclimated in the greenhouse were grown to appropriate size and condition for pathogen testing and provided to cooperators for greenhouse and field pest and disease trials. Additional plants of six crown gall resistant transgenic lines and controls were produced for use in further field testing and in nursery and orchard performance trials. The first field trial of transgenic crown gall resistant lines was established in 2008 at UC Davis and grafted to Chandler. Trees in this trial were used this year to supply bark and nut material for studies of trans-graft union movement of macromolecules. Two additional field trials, one in the UC Davis Plant Pathology facility and one in a commercial nursery were initiated. Greenhouse stab tests of seedlings of wingnut accession DPTE 1.09 continued to show segregation for crown gall resistance in second year re-testing, with five of nine tested seedlings producing no galls and four seedlings producing large galls both years. Hardwood cuttings were used to produce clonal replicates of several seedling selections exhibiting potential crown gall resistance. We showed that mouse-ear leaf symptoms appearing in some genotypes and soil combinations in the greenhouse, particularly after chilling, could be remedied by application of nickel, we improved the efficiency of methods for introducing seed from rootstock crosses into culture, and we continued developing methods for budding or grafting small container-grown plants. Field trials of clonal rootstocks continue to be established state-wide by farm advisors and we continue to supply plant material, assistance, and technical support to commercial laboratories and nurseries producing and selling improved clonal rootstocks. GOAL AND OBJECTIVES The goal of this project is to provide the California walnut industry with new clonal rootstocks selected or designed to combat the most threatening pests and diseases. Objective include improved efficiency in clonal propagation of walnut, devising clonal propagation methods for new candidate genotypes, and to providing clonal plantlets for greenhouse and field replicated disease and pest challenge tests. California Walnut Board 97 Walnut Research Reports 2012

PROCEDURES AND RESULTS Propagation methods: We have used two approaches to clonally propagate candidate rootstock genotypes for nematode, crown gall, Phytophthora, Armillaria and blackline tolerance or resistance testing: A. Tissue culture micropropagation with in vitro and ex vitro rooting of microshoots. B. Dormant hardwood cuttings on bottom heated beds. Tissue culture micropropagation: This year we successfully rooted microshoots of 121 individual genotypes (Table1) and produced more than 7300 fully acclimated greenhouse plants (Table 2). In addition to controls, PDS clones, English cultivars, CLRV tolerant selections, and crown gall resistant transgenics, we produced over 4000 plants of 44 new paradox genotypes initiated into culture over the last two years from immature zygotic embryos. These include 33 J. microcarpa (DJUG 29.11) x Serr, 20 J. cathayensis (DJUG 11.3) x Serr, and 34 English x J. cathayensis (selection #21) paradox hybrid genotypes (Table 2) Plants produced were grown to appropriate size and condition for use in replicated disease and pest screening tests. These included greenhouse Phytophthora trials by Greg Browne, Mike McKenry s field trials for nematode response, and crown gall testing by Dan Kluepfel (Table 3). Plants produced also included about 1000 plantlets of transgenic lines expressing an RNAi construct for resistance to crown gall and controls for use in greenhouse, nursery, and orchard trials (Table 2). Improved method for extracting and culturing zygotic embryos for rootstock development from hybrid seed of black walnut and Asian butternut species Breeding and selection for resistance to rootstock pathogens frequently requires introduction of candidate genotypes into tissue culture to create multiple clonal copies for pathogen testing. The most efficient way to introduce new candidate rootstock genotypes into culture is to extract immature embryos from nuts. Because these are still enclosed in the shell and protected by the hull from external contamination, they can be much more easily disinfested from external contaminants than can nodal cuttings taken from germinated seedlings or mature trees. Often embryos can be extracted cleanly from nuts using sterile instruments even if the shell is not completely clean. Embryos of English walnuts, Juglans regia, can usually be extracted prior to shell hardening and nuts can be opened easily with a pocket knife. An encircling cut part way through the hull about 1/3 of the total distance from the blossom end followed by a twist of the blade will flip the blossom end of the hull off and cleanly expose the immature embryo. Unfortunately, shells of black walnuts and Asian butternuts, species currently of interest and in use for developing pathogen-resistant rootstocks, harden before the embryos become visible in the interior cavity. Embryos have been successfully extracted at this stage by removing the liquid or jellied contents and growing in culture until they become visible but reliability is low and the possibility of developing triploid plants from the endosperm exists. We have achieved reasonable success in the past by cracking nuts near or at maturity, first disinfesting them with a 10-20% bleach solution, then wrapping them in sterile paper towels and cracking with a vice in a laminar flow hood. California Walnut Board 98 Walnut Research Reports 2012

One factor limiting efficiency and reducing the percent success in this process has been inability to crack them so the embryo is consistently exposed and accessible. Nuts of these species contain solid woody inner septa and the cavities are normally completely filled with kernel. Depending on the direction and location of the crack that develops it is sometimes relatively easy to identify and extract the embryo from among cotyledon and shell fragments and at other times impossible to identify the embryo among the debris or it remains inaccessible in an uncracked portion of the shell and the more it is necessary to pick through fragments or to culture extraneous cotyledon pieces the more chances for contamination in the process increase. This year Maryam Fazel, a student in our lab, through careful observation determined the optimal placement of nuts of J. microcarpa and J. cathayensis during cracking, resulting in considerably increased reliability and efficiency in extracting embryos. For J. microcarpa, the first step is to note and keep track of the location of the blossom end while wrapping the nut in a clean and disinfected paper towel as shown in Figure 1. Once the nut is completely wrapped, it needs to be placed in the vice so that only the stem-end half of the nut is in the clamps of the vice and the blossom-end half is above the clamps as shown in Figure 2. Figure 1. Photo of a J. microcarpa nut being wrapped in a sterilized paper towel Figure 2. The orientation of the J. microcarpa nut inside the vice The vice is closed only until hearing the first two cracking sounds. The vice is then opened and the towel unrolled, touching only the edges of the towel or using a sterile forceps, to allow access to the cracked walnut. Figure 3, shows the exposed embryo after this process. The embryo is then extracted using a clean set of instruments that have not touched the exterior of the nut and is transferred onto the desired medium for culture. California Walnut Board 99 Walnut Research Reports 2012

Figure 3. The intact uncrushed embryo after cracking a J. microcarpa nut This approach was then adapted for an Asian butternut, Juglans cathayensis. For this species, unlike J. microcarpa, we were able to determine the suture location while the nut was still I the hull by observing the indentations at the stem-end of the hull. For best results, the nut should be placed so the suture sides of the shell are in contact with the clamps of the vice, the blossom end of the nut up and only the stem-end half of the nut between the clamps as shown in Figure 4. This results in the correct amount of pressure and direction of cracking to consistently expose the embryo end of the kernel when the cracked shell is opened as shown in Figure 5. In practice, the nut is wrapped in a sterile paper towel during this process. Cheek of walnut shell Vice clamp Suture side of walnut shell Figure 4. The suture side of the walnut shell in contact with vice clamps California Walnut Board 100 Walnut Research Reports 2012

Figure 5. The whole uncrushed embryo exposed after cracking a J. cathayensis nut Tissue culture media improvement: We have routinely used Driver Kuniyuki Walnut (DKW) medium as our standard propagation medium. Over time it has become evident that this medium is not optimal for all the species and genotypes we propagate. Although wingnut grows rapidly and is very hardy in the greenhouse and field, it has long struggled in culture, multiplying slowly and prone to vitrification. In a trial this year we tested a media with the A Stock (CaNO2, NH4NO3, and ZnNO3) component increased to 150% of the standard formulation. This had the effect of increasing the nitrogen concentration in the medium but also increased zinc and calcium as a byproduct. We test various genotypes on this 855 medium including wingnut WNxW, Juglans hindsii selection W17 which exhibits spindly growth, and several J. cathayensis clones that elongate poorly. This medium was much softer than DKW and we increased the amount of gel to strengthen it to normal consistency. Several genotypes tested showed marginally improved growth but wingnut responded most clearly to this medium, exhibiting improved elongation, reduced vitrification, and more normal leaf size and appearance. We now culture wingnut exclusively on this medium. We then attempted to determine if a single component of the A stock could be modified to advantage. Four different media were tested as shown in the table below using microshoots of Vlach, J. hindsii W17, wingnut WNxW and Chandler, four magentas of each, and in each medium doubling only one of the three components.. A Configuration NH 4 NO 3 (mg/l) Ca(NO 3 ) 2 (mg/l) Zn(NO 3 ) 2 ( mg/l) #1 (DKW = control) 1416 1968 17 #2 2832= (2x) 1968 17 #3 1416 3936 =(2x) 17 #4 1416 1968 34=(2x) Media #2 (2x ammonium nitrate) always performed best overall. W17 looked better on all of these media when compared to DKW. Media #3 (2x calcium nitrate) appeared best for wingnut but the media was too soft to use. It was clear that calcium nitrate is the ingredient responsible for making the gel less effective and the media too soft in 855 (DKW with 50% increased A stock). Plants on media #2 and #3 (highest in nitrogen) grew best and also went downhill the California Walnut Board 101 Walnut Research Reports 2012

slowest when left for a long transfer interval. This experiment showed that increasing the nitrogen without increasing the amount of calcium could potentially improve the quality of the plants but this was a small experiment and results need to be confirmed.. We next compared the growth of six diverse genotypes on six published and commercially available media and compared these to growth on DKW. Genotypes employed were Chandler and RX1 as standards, J. hindsii W17 and wingnut WNxW as two genotypes that need a medium for improved shoot quality and Serr seedling S8 and J. cathayensis x #21 seedling 3S17 as two that elongate poorly on DKW. Media tested were Quorin & Lepoivre, Rugini Olive, Broadleaf Tree, Gamborg s, Schenk & Hildebrandt, and Nistch &Nitsch. In all cases these media were clearly inferior to DKW for growth and shoot quality for all genotypes tested. Experiment 1: Average height in cm at completion DKW Quorin & Lepoivre Rugini Broadleaf Chandler 2.9 1.3 1.8 1.3 RX1 3.5 1.3 1.3 1.1 W17 3.9 1.9 4.8 0.9 WNxW 6.0 4.0 3.1 1.2 S8 3.0 1.9 2.4 1.6 3S17 1.1 1.5 1.1 0.7 Experiment 2: Average height in cm at completion DKW Gamborg s Schenk &Hildebrandt Nitsch & Nitsch Chandler 3.3 0.8 1.1 0.6 RX1 6.4 2.1 2.0 1.5 W17 1.2 2.0 2.3 2.5 WNxW 2.3 0* 3.0 0.5 S8 3.8 1.0 1.4 0.9 3S17 1.1 0.5 0.5 0.5 * All died In addition, we tested use of carrageenan as gelling agent in place of Gellan gum (Gelzan). Carrageenan gives a very clear medium with a bit stickier composition when used with standard DKW. When we tested this with diverse genotypes we saw no significant difference in growth or rooting capabilities and discontinued use because it costs more than Gelzan. When we used carrageenan with the high nitrogen medium, 855, it resulted in a somewhat cloudier medium that was a disadvantage for detecting contamination. Finally, we tested elimination of CaCl 2 from the medium. DKW includes CaCl 2 as the D stock but there would appear to be no advantage to adding chloride and this represents a relatively small portion of the total calcium in the medium. Medium without CaCl 2 was tested using genotypes AX1, RX1, Chandler, W17, wingnut WNxW, and J. cathayensis x Serr JCS1. Removal of CaCl 2 resulted in no clear differences in growth, in vitro, plant appearance, greenhouse survival, or rooting. Hardwood cuttings: Hardwood cuttings were used to clonally propagate open pollinated seedlings of mother trees at the USDA National Clonal Germplasm Repository, primarily J. microcarpa, that that had shown no previous gall formation or only small gall formation in testing by Dan Kluepfel s lab. A total of 272 cuttings were made in early February using the protocol described in the 2009 report. Only 56 of these were successfully rooted and then grown in 3 liter avocado pots for re-testing. California Walnut Board 102 Walnut Research Reports 2012

Use of Etiolation and Air Layering to Enhance Clonal Propagation: As indicated in the previous section, success in propagation using hardwood cuttings is problematic and degree of success is dependent on species and genotype. So for research purposes it would be advantages to have an alternate method for rooting greenhouse or field-grown material. This past year we have experimented with etiolation and air layering to enhance rooting potential and root initiation. Etiolation is the growth of shoots in total darkness and in many species enhances rooting potential. Air layering allows adventitious roots to form on shoots without detaching the shoot from the mother plant which can be a great advantage for species that are difficult or slow to root like walnut. The two processes can be used independently or in combination. In combination, air layering is a way to capture the benefits of enhanced rooting potential of etiolation treatment. We have found that shoots of fully chilled, containerized plants of several walnut species grow very well in total darkness to provide etiolated shoots for rooting. However, these shoots do not generally form many adventitious roots until treated with potassium indolebutyric acid through longitudinal incisions and given a moist environment. The moist environment for air layering can be provided by using a sleeve filled with moist wood shavings to cover the whole etiolated stem or to specific locations by using a rooting cube composed of plasticized peat moss. When rooted air layered stems have been detached and potted into containers under greenhouse conditions they have survived and grown. The air layering techniques even have worked for non-etiolated stems but the rooting takes longer. These procedures have worked very well for rooting all the species we ve tried except J. regia which is still recalcitrant for rooting. These procedures need to be improved to make them more efficient. Developing a source of scion material for bench-grafting micropropagated rootstock liners: Some walnut rootstock micropropagation laboratories and nurseries would like to be able to routinely graft English scions onto micropropagated clonal rootstock liners. One of the limitations to being able to do this is a source and adequate supply of suitable-diameter shoots from which to take scions or buds. Orchard-derived shoot material is generally much too large for this purpose. During the past year we have. During the past year we continued to work on procedures to produce English walnut scion wood of an appropriate diameter (1/4 inch or less) to bud onto micropropagated clonal walnut rootstock liners. Using own rooted Chandler trees growing under field conditions, and in containers in a greenhouse, we have been using various treatments to try to maximize production of small diameter shoots. Our premise is that by restricting root volume (container size) and/or increasing the number or density of shoots we can maximize the number of small diameter shoots. Last year we reported that with plants growing in 1.5 liter containers increased branching and decreased diameter could be obtained by pruning off the terminal bud and treating the remaining axillary buds with Promalin after chilling was fulfilled in the spring. However, with Chandler plants growing in the field this treatment didn t work well to increase the number or density of shoots growing and therefore the diameter of shoots developed was much too large to provide bud wood for use on liner-sized rootstock. This year we used some different approaches on field grown trees. The one that worked the best involved heading back existing shoots on previously hedged trees in late March and using an opaque tarp on frames to cover the trees so new shoots develop and grow in the dark. Actively growing shoots were then headed back again in June and allowed to grow in either full sun or shade provided by Saran shade cloth structures. These developed many shoots that were small diameter (~ ¼ inch) and had sufficient lignification (hardness) for potential use as bud wood. California Walnut Board 103 Walnut Research Reports 2012

Budding small containerized walnut rootstock Efficiency of establishing orchards on clonal rootstocks exhibiting improved pathogen resistance would be greatly enhanced by development of reliable bench grafting techniques for small containerized walnut plants so that clonal rootstock can be routinely budded in a greenhouse or lath house and sold already budded by nurseries, eliminating the need to plant these in a nursery for sizing, reducing expose to soil pathogens in the process, and increasing uniform orchard establishment if sold directly to growers. This year we conducted a large budding trial using 190 tissue culture-produced greenhouse plantlets, all of which had been through one dormancy cycle, had leafed out, and had been transplanted into a larger pot between 3/15/2012 and 5/15/2012. Plants were budded the week of 7/13/2012-7/17/2002. We used two kinds of pots, one a standard 10 x10x36cm, 2.63 liter tree pot with a slotted open bottom, the other a 24 cm deep round pot with a circular mesh bottom. We tested three watering regimes followed by three acclimatization regimes. The watering regimes were wet, dry and medium, always watered with half strength Hoagland s solution (referred to here as water ). For the wet treatment, the bases of the pots were submerged in approximately 2 of water, and the level was maintained by manually replenishing daily when low. For the mid-water treatment the pots were placed into approximately 3 of water daily for approximately 30 minutes and then drained them. For the dry treatment, all water was withheld for two days before budding and re-instated a minimum of two days after budding, depending on the acclimatization treatment, at which point 300ml of replacement water was applied at differing intervals, depending on acclimatization treatment. The amount was determined from rough measurements of average evapotranspiration obtained by weighing several pots from which water was withheld in each of the treatments to establish an average amount of water lost per day (Table 1). A rough rule of thumb developed was two days without water on the greenhouse bench was the equivalent of 3 days without water on the shaded bench and 5 days in the cold. Our intention was to dry plants to a point not approaching the permanent wilting point (PWP) but to a point that slowed metabolism, and then slowly add slightly more than ET over a period of time until the soil in the pots returned to saturation. We knew, from a previous experiment that plants which were deprived water lost water rapidly for the first 1-3 days, then much more slowly from that point until they reached PWP, which could take as long as 2-3 weeks from initial withholding. We established and maintained these watering regimes from two days prior to budding until at least 28 days afterwards. We simultaneously established three different acclimatization regimes; greenhouse, shade, and cold. For the first of these, plants went directly to a greenhouse bench uncovered and open. For the second, plants were place on a bench covered completely with 75% black shade cloth for the 10 days immediately after budding, then moved to an uncovered greenhouse bench. For the third, plants were placed in a growth chamber maintained at 65 F for 10 days and then moved to the heavily shaded bench for five days. The length of time for each of these treatments was designed to allow the buds to heal in before we attempted to subject them to full greenhouse conditions. Prior to budding all plants were held in a lath house for at least two days. California Walnut Board 104 Walnut Research Reports 2012

We also used three different sources of scion material. The first source (GR) was plants grown in the greenhouse in pots that had been repeatedly pruned to maintain small diameter. The second (TOP) was field-grown stock plants which we had also been pruned repeatedly to induce small diameter growth. These were also etiolated during the spring growing period by enveloping them under a tarp, which was later replaced by a shade cloth in early June (see Scion Development section above). The third source (CGR) was scion wood from an unpruned 3 rd leaf Chandler orchard, primarily new lateral growth developing from one-year-old shoots. We used a T-budding technique established in a series of small experiments prior to this larger experiment. This involved making a small T-shaped incision on the rootstock with an Exacto knife. A small shield of slipping bark was cut from the scion and slid into the rootstock cut until the horizontal cut at the top of the shield matched the horizontal cut on the rootstock and the two flaps on the rootstock held the shield firmly in place. We then wrapped it with Parafilm 2-3 times around completely and tightly, especially tightly around the base of the incision. We then used the Exacto knife to make a small hole in the Parafilm for the bud, to allow it to respire freely, and a horizontal incision at the base of the rootstock to relive any sap pressure that might develop. All puts were placed into this year s growth on the rootstock and most plants received two buds. Scion calipers were measured immediately above the buds and rootstock calipers were measured above the 2 nd bud of the new growth. We also scored the maturity and ability of bark to slip both the rootstock and scion on a one to five with the larger number meaning less mature and more likely to slip respectively. We also took periodic soil temperature readings and weights to estimate soil saturation of a sub-set of plants. On all plants we removed the rootstock apical buds 7 days after budding to stimulate lateral bud growth and healing. After 21 days we cut the Parafilm longitudinally on the side of the rootstock opposite the bud, but did not remove it fully until 28 days. At 28 days we pruned the rootstock two nodes above the bud and removed any buds above the budded scion. After 35 days we assessed the bud survival, health, and elongation. Buds which remained green and/or had begun to swell and grow were counted as successful. The best of the treatments tested, pots immersed in water and kept on an open greenhouse bench during healing, gave us 82% bud-take (see Tables 4-12). Pots immersed and kept in the shade was the next best treatment with 72% take. The number of plants budded in each individual treatment was relatively small and the plants were relatively mature for containerized plants but difference between some treatments and trends across combined treatments were clear and did not match conventional assumptions. A key observation from this work was the surprising improvement in bud-take we obtained by submerging the base of the pots constantly in water. This drastically improved bud-take in all treatment except for the cold acclimatization in which take was poor across treatments. One explanation possibility is that plants not sitting in water are actually receiving a mild amount of water stress when watered daily. A more likely explanation is that soil water buffers soil temperature and independent temperature data confirms this. Pot temperatures do get high in the greenhouse, in excess of 90ºF in some cases when not immersed. Another cause could be release of plant hormones associated with inundation of roots. California Walnut Board 105 Walnut Research Reports 2012

The open greenhouse bench appeared to be the best bud-healing condition, leading us to believe that there is nothing to be gained from gradual acclimatization after budding. The next important factor was scion source and maturity. Field-grown buds from unpruned young trees gave the best results. This correlated with our maturity scoring, which indicated that more mature scion material gave the best take. The unpruned young trees gave us the most mature wood. The opposite was true for rootstocks. Take was best on the least mature rootstocks. This work was undertaken to find ways to improve bench budding of containerized trees for commercial nursery production but findings will also assist us in propagating promising breeding program selections in the greenhouse, well before we would normally begin to move them to selection blocks in the field. We also hope to use the methods developed expedite CLRV patch testing by moving this to the greenhouse. We plan to repeat this experiment again next year and will attempt to improve further by testing additional budding methods, hormone use, and timing, and intend to transition towards a comprehensive method for micro-grafting smaller liner plants. Mouse Ear Symptoms on Walnut Rootstock Liners: A condition referred to as Mouse Ear has been observed during the production of clonal walnut rootstock liners in commercial greenhouses and our research greenhouse. The symptoms of reduced size and cupping of new leaves and leaflets have been observed in peat-perlite (Sunshine #4 mix) grown liner plantlets resuming growth in spring after receiving natural winter chilling and in our research greenhouse when the peat-perlite soil ph slightly above 7. Last year we presented results showing that including Micro Max at 1 lb/yd3 in the peat-perlite soil or reducing the ph to 6.5 reduced or eliminated the symptoms. However, these results didn t tell us the cause of the symptoms but did suggest that a micronutrient deficiency might be involved. Nickel deficiency has been shown to be the cause of similar leaf symptoms in pecan and some other tree species. This year we did an experiment to see if we could eliminate the symptoms in walnut rootstock liners (RX1, VX211, Vlach) by incorporating Micro Max, nickel or a combination of the two in Sunshine #4 mix at ph 6.5. As shown in Table 13, Micro Max was added at the rate of 1 lb/yd3 and Ni S04. 6 H2O at the rate of 0.0224 lbs/yd3. During initial growth of the liner plants after imposing the treatments no Mouse Ear symptoms developed. However, after resuming growth in the greenhouse after exposure to natural winter chilling almost 50% of untreated control plants showed Mouse Ear symptoms (Table 13). More than 25% of those receiving Micro Max showed symptoms but none of those receiving nickel or nickel plus Micro Max showed symptoms. These results indicate the nickel deficiency is the cause of Mouse Ear symptoms in walnut rootstock liners and can be corrected by adding nickel to peat-perlite soil mixes. However, our results (Table 14) indicated that the rate of nickel alone that we added was inhibitory to growth in VX211 and RX1. The optimal rate of nickel treatment to correct symptoms and give maximum growth needs to be investigated further. There was also some suggestion that Micro Max may have had some beneficial effects on growth especially in overcoming the inhibitory effects of nickel. Crown gall resistance in wingnut Last year we examined the relative susceptibility or resistance to crown gall of wingnut seedlings grown from seed of five different NCGR wingnut mother trees. All seedlings from four of those mother trees showed significant galling (see 2011 Propagation Report) but results, now based on California Walnut Board 106 Walnut Research Reports 2012

4 stab inoculations per seedling per year with repeat inoculations in two consecutive years on the same seedlings, show clear segregation for crown gall resistance among the seedlings of NCGR accession DPTE 1.09 (Table 15). Five of the offspring of this wingnut mother-tree developed no galls in two years of testing while the other four seedlings tested produced consistently large galls, including some of the largest measured in a trial last year that included Paradox genotypes. These results suggest further investigation of the crown gall resistance of wingnut accessions and genetic segregation for resistance in this accession in particular. Field Trials: Transgenic crown gall resistant rootstock trials: A field trial of transgenic line expressing resistance to crown gall, non-transformed background genotypes as controls, and other rootstock genotypes of interest, was established on the UCD campus in 2008 and most trees have been successfully budded to Chandler. This trial continues to be evaluated for horticultural performance and natural occurrence of crown gall. The block includes budded trees of the six best genotypes selected to move forward in the testing and potential release process. Scion and rootstock material from this trial have been, and will continue to be, used for work now in progress to assess any possible trans-graft union movement of DNA, RNA or other macromolecules. Both bark and nut samples were collected for use in these evaluations this year. In addition, a trial of the six best transgenic genotypes and the two control background genotypes was established at the Armstrong Field Station on a site previously used for crown gall testing. These trees will be exposed to the crown gall causing organism likely already present in the soil trees at this could be used in the future for trials using applied bacteria in the field if permits are approved. A third planting is anticipated early next year using plants currently in the greenhouse. This planting would be at a commercial nursery to produce trees grafted to Chandler scion that could then be used for a first commercial orchard trial. Farm advisor/grower county rootstock trials: Currently established clonal rootstock field trials in grower orchards in the various counties are summarized in Table 16. Most of the initial clonal rootstock field trials were established in replant situations but farm advisors have now established a number of clonal rootstock trials at newly planted orchard sites in diverse counties, several with replicated plot designs, and with a variety of scions and soil types. In addition, trials have been established on several challenging situations including unfumigated, walnut following walnut, nematode, and Phytophthora infested sites. Rick Buchner established a trial of RX1, VX211, and Vlach in a Tehama County orchard (H. Crain) in 2009 using nursery-grown trees developed from commercially produced liners. The clonal rootstock genotypes were planted in a replicated design and budded to Howard (see separate report in this volume). Rachel Elkins has initiated four field plantings in Lake County comparing VX211, Vlach, and seedling paradox planted in freshly prepared but un-fumigated sites thought likely to subject trees to nematode pressure (see report in this volume). California Walnut Board 107 Walnut Research Reports 2012

Joe Grant and Joe Conant established plantings near Wheatland of RX1, VX211, and Vlach grafted or budded to Ivanhoe, Sexton, Howard, and selection 91-077-40. A portion of these plantings are under power lines and evaluations will include observation of rootstock and scion effects on tree height. This trial also provides an opportunity to observe rootstock-scion interactions for several combinations, particularly effects on Howard performance, and further observation may give insight into current Howard yellowing problems seen in many orchards. Janine Hasey continued to observe a trial in Sutter County on a site with known and long-term Phytophthora cinnamomi problems. This replicated trial in a replant situation includes RX1 containerized and bare-root trees, VX211 bare root, and Paradox seedling trees (see report in this volume). A replicated clonal rootstock trial in Solano County (Cilker) was established by Carolyn DeBuse and Bruce Lampinen in 2009 on a good soil site. This trial includes RX1, VX211, Burbank and Vlach grafted to Tulare (see report in this volume). A VX211 planting in Kings County (Verboon) established by Bob Beede includes VX211 grafted to Tulare in a fumigation treatment trial with Vlach planted in the buffer rows. Trees at this location initially exhibited excessive vigor which may have contributed to reduced numbers of the secondary axillary buds on lower trunks needed for low scaffold development. Four trials which include the cherry leafroll-tolerant WIP rootstocks and own-rooted Chandler have been established by Janet Caprile and Joe Grant and are currently under evaluation. These are located in Contra Costa County (see Caprile report in this volume) and San Joaquin County (see San Joaquin County section of this report). Several additional rootstock trials to evaluate performance of English cultivars on their own roots have also been established by farm advisors in Butte, Yuba, and Stanislaus counties. Observations for these have been reported in recent volumes. See also summary of rootstock trial locations in Table 16. San Joaquin County rootstock trials - Joe Grant: Concar Ranch Trial: Performance of clonal Paradox and blackline tolerant walnut rootstocks in San Joaquin County Project leader: Joe Grant Cooperating personnel: Brett Lagorio, Concar Ranch, Linden Bonilla Nursery, Oakdale Location: North of Flood Road, East of Escalon-Bellota Road, approximately 5 miles east of Linden, San Joaquin County (Approximate GIS location: 38 1 32 N 120 59 0 W) Soil at the site is Redding Gravelly Loam. The site was previously farmed as irrigated pasture. Prior to planting the soil was ripped (two directions) to six foot depth. Scion variety is Chandler. Rootstocks include Own-rooted Chandler, RX1 clonal Paradox, VX211 clonal Paradox, WIP3, Vlach clonal paradox (June-budded and standard 2 year-old budded), AZ025, and seedling Paradox (June-budded, J. hindsii X J. regia per Bonilla Nursery). Spacing is 17 X 22. Irrigation was by single-line above-ground drip through the third year, then double-line drip to present drip. The trial is configured as a randomized complete block design with five 4 to 8-tree replications. California Walnut Board 108 Walnut Research Reports 2012

All trees were hand-planted in February 2008 as finished bare-root nursery trees. First- through fourth -year vegetative growth has been vigorous and uniform. Trees have been pruned and trained using the Barton multiple scaffold system. Trunk diameters measured at the end of the second through fifth growing seasons were largest for own-rooted Chandlers, smallest for VX211, with other rootstocks intermediate. Beginning with trunk diameter and yield measurements in 2011, data for June-budded and standard 2- year old trees were pooled because early tree growth and canopy development rate were similar for these rootstocks. Mid- Mid-summer midday canopy photosynthetically active radiation interception values (PAR), a measure of canopy size, ranged from 43% to 50% in 2011 and 67% to 75% in 2012. There were small but statistically significant differences in canopy size among rootstocks, with VX211 and WIP among the smallest and RX1 and Paradox seedlings among the largest. Nut production in 2011 did not differ among rootstocks, except for own-rooted Chandler trees, which had the lowest average yield. More statistically significant separation among rootstocks was apparent in 2012, with own-rooted Chandlers continuing to produce the least and RX1, Vlach, Paradox seedlings, and AZ025 the most. Avg. trunk diameter, inches 1 Midday light interception 2, % Avg. in-shell yield 3, lbs/a Rootstock Dec-09 Dec-10 Nov-11 Nov-12 2011 2012 2011 2012 Own-rooted Chandler 2.7 a 4 4.3 a 5.4 a 6.2 a 45.4 bc 70.6 bc 753 b 2545 d RX1 2.5 b 3.8 b 4.9 b 5.8 b 50.3 a 75.2 a 1967 a 4990 a Paradox seedling 3.7 (J. hindsii X J. 2.4 bc bcd regia) 4.7 bc 5.6 c 47.4 ab 73.2 ab 1654 a 4480 ab Vlach (June & Std. budded) 5 2.4 cd 3.6 cd 4.6 cd 5.5 cd 45.6 bc 71.5 abc 1996 a 4740 a WIP3 2.4 cd 3.8 bc 4.8 bc 5.6 c 42.6 c 67.4 c 1829 a 3807 c AZ025 2.3 de 3.5 de 4.5 de 5.4 d 44.5 bc 73.1 ab 1796 a 4380abc VX211 2.3 e 3.4 f 4.4 e 5.2 e 43.3 bc 67.9 c 1673 a 4044 bc 1 Measured 24 inches above soil line; 2 Measured 29 July 2011 and 8 September 2012; 3 8% Wet basis moisture content; 4 Values within columns followed by different letters are statistically different, Fisher's Protected LSD, P<0.05; 5 Data for Vlach June-budded and Vlach standard 2 year-old budded trees pooled. Chiappi Farms Trial Performance of RX1 clonal Paradox walnut rootstock in San Joaquin County Project leader: Joe Grant Cooperating personnel: Tony Chiappe, Chiappe Farms, Stockton, CA Greg Browne, USDA-ARS, Davis Burchell Nursery, Oakdale, CA Location: South Highway 4, east of Hewitt Road, approximately 1.8 miles west of Farmington, San Joaquin County California Walnut Board 109 Walnut Research Reports 2012

Approximate GIS location: 37 55 39 N 121 1 59 W Soil at the site is Archerdale Clay Loam. The site was previously planted to walnuts which died from Phytophthora root rot (isolated and identified from root and soil samples as P. cinnamomi), and were removed one or two years prior to planting. Tree sites were pre-plant fumigated with 1 pound methyl bromide. Rootstocks include RX1 clonal Paradox and seedling Paradox (J. hindsii X J. regia per Burchell Nursery). Tree site spacing is 28 X 28. Each tree site is planted to two trees - one RX1 and one Paradox seedling - spaced about 2 apart, paired roughly by tree size at planting. Irrigation is by impact sprinklers. Experimental design: Randomized complete block design with ten ten-tree (paired tree) replications. All trees were hand-planted in March 2010 as bare-root un-grafted nursery whips and budded in August 2010 to Serr. Trees not successfully top-worked by this initial budding were grafted to Serr in spring 2011, and a small number of these trees were re-budded in August 2011. RESULTS There was no first-year (2010) tree mortality. Tree growth, as measured by trunk circumference increment, was similar for the two rootstocks in 2010 (all trees measured). Because of differences in tree growth due to budding and/or grafting success, 2011and 2012 tree circumference data were only recorded for trees that had been successfully budded in 2010 and both trees of the RX1-seedling pairs had survived (n=51 and 45 in 2011 and 2012, respectively). There were no statistically significant trunk circumference differences in 2010 and 2011 but, by the end of 2012, surviving trees on RX1 had significantly outgrown those on Paradox seedlings (Single factor ANOVA, P<0.001). Trunk circumference, cm* July 2010 Dec. 2010 Nov. 2011 Nov. 2012 Paradox seedling 5.4 7.4 12.4 12.9 RX1 5.5 7.7 12.4* 20.7 *Trunk circumference measured 30 cm above soil. 2010 data include all trees. November 2011 data include only trees in intact 2010 budded seedling-rx1 pairs. By late September 2011, 17% of trees on Paradox seedlings were dead and an additional 6% were growing very poorly compared to other trees. By November 2012, 31% of seedling trees had died and an additional 17% of seedling trees remaining after the 2011 season were growing poorly. No RX1 trees have died to date and all were growing well at the end of the 2012 season. Root samples were collected November 2011 and again on August 2012 from dead and poorly growing Paradox seedling trees and subsequently cultured on Phytophthora selective PARP medium. Phytophthora cinnamomi was isolated from one or more root pieces cultured from 63% of dead trees and 40% of poorly growing trees in 2011, and from 54% of dead trees and 21% of poorly growing trees in 2012, indicating that P. cinnamomi infection was a principal cause of tree death and decline in the trial. California Walnut Board 110 Walnut Research Reports 2012

Nursery Propagation and Commercialization: We continue to be prepared to provide cultures of microshoots to any commercial laboratory or nursery that requests them for licensed production of plants. Appendix 1 of this report includes a list of laboratories currently licensed for in vitro production of clonal rootstocks RX1 and VX211 and for sale of clonal rootstock plantlets as liners for nursery or orchard planting. California Walnut Board 111 Walnut Research Reports 2012

Table 1 (part1). Ex vitro rooting percentage for rootstock genotypes. # Attempted # Rooted % Rooted PDS Paradox AX1 1149 856 74 Px1 552 404 73 RX1 1415 911 64 Vlach 651 445 68 VX211 438 344 79 4205 2960 70 Black W17 1285 437 34 Wingnut WNxW 10.05 b 464 371 80 CLRV Tolerant WIP3 153 94 61 English Chandler 617 76 12 J. microcarpa JMOP 2 76 59 78 3S 14 K3 K5 325 32 23 230 23 16 71 72 70 456 328 72 J. microcarpa x Serr JMS 3 144 104 72 JMS 4 207 156 75 JMS 5 89 82 92 JMS 5A 129 118 91 JMS 7 158 134 85 JMS 9 187 150 80 JMS 11 208 173 83 JMS 11A 229 188 82 JMS 12 117 83 71 JMS 13 147 102 69 JMS 15 137 98 72 JMS 18 156 125 80 California Walnut Board 112 Walnut Research Reports 2012

JMS 19 169 125 74 Table 1 (cont.). Ex vitro rooting percentage for rootstock genotypes. # Attempted # Rooted % Rooted JMS 20 228 146 64 JMS 21 156 93 60 JMS 24 110 86 78 29 JM 1 270 227 84 29 JM 2 91 65 71 29 JM 3 195 165 85 29 JM 4 76 67 88 29 JM 5 361 226 63 29 JM 8 169 133 79 29 JM 10 99 68 69 29 JM 11 99 83 84 29 JM 12 126 109 87 29 JM 17 246 196 80 29 JM 22 124 109 88 STJM 4 126 110 87 STJM 6 182 115 63 STJM 7 164 88 54 STJM 11 153 103 67 5052 3827 76 J. cathayensis x Serr JCS 1 32 20 63 JCS 2 16 14 88 JCS 3 131 94 72 JCS 6 36 34 94 JCS 7 1 0 0 3S 3 8 4 50 3S 5 63 50 79 3S 6 2 2 100 3S 8 3 3 100 3S 9 17 12 71 3S 10 8 1 13 3S 12 2 0 0 3S 13 1 1 100 3S 16 3 2 67 3S 17 95 60 63 3S 18 79 50 63 3SH 1 1 0 0 3S 19 8 1 13 California Walnut Board 113 Walnut Research Reports 2012

3SH 2 44 32 73 Table 1 (cont.). Ex vitro rooting percentage for rootstock genotypes. # Attempted # Rooted % Rooted 3SH 3 3 1 33 3SH 4 11 4 36 3SH 5 5 2 40 3SH 6 17 12 71 3SH 7 77 40 52 680 447 66 Serr x J. cathayensis #21 SC 1 42 2 5 SC 5 77 22 29 SBB7 218 57 26 SM 1 169 37 22 SM 5 229 29 13 S 1 93 13 14 S 8 184 13 7 S 10 113 8 7 S 13 78 11 14 S 15 131 3 2 S 16 11 4 36 S 17 23 1 4 S 21 114 14 12 S 25 127 1 1 S 32 7 5 71 S 35 69 6 9 S 38 38 5 13 S 39 73 15 21 S 40 57 5 9 S 42 69 3 4 S 45 54 1 2 S 49 22 1 5 S 54 25 1 4 S 58 55 5 9 S 59 31 8 26 S 61 78 26 33 2187 296 14 California Walnut Board 114 Walnut Research Reports 2012

Table 1 (cont.). Ex vitro rooting percentage for rootstock genotypes. UC 95 007 13 x J. cathayensis #21 # Attempted # Rooted % Rooted 7 1 90 5 6 7 2 65 3 5 7 3 97 1 1 7 4 70 5 7 7 10 43 37 86 7 11 3 2 67 7 12 2 2 100 7 13 11 8 73 7 14 13 12 92 394 75 19 Transgenic # Attempted # Rooted % Rooted L 33 139 83 60 L 66 164 110 67 L 194 71 50 70 L K 136 74 54 L J 143 111 78 L U 212 146 69 L V 163 95 58 J1 L 118 87 74 J1 1A 385 220 57 J1 19A 433 298 69 J1 20A 260 129 50 RR4 1A 198 137 69 RR4 8A 87 75 86 RR4 10A 176 111 63 RR4 11A 56 44 79 RR4 12A 147 102 69 J1a control 399 247 62 RR4 control 143 108 76 2402 1558 65 Total 18868 11099 59 California Walnut Board 115 Walnut Research Reports 2012

Table 2. (Part 1) Greenhouse survival of rooted clonal microshoots 2012 # Rooted # Survived % Survival PDS Paradox AX1 717 547 76 Px1 287 191 67 RX1 757 594 78 Vlach 337 246 73 VX211 258 214 83 2356 1792 76 Black W17 349 153 44 Wingnut WNxW 10.05 b 309 280 91 CLRV Tolerant WIP3 57 28 49 English Chandler 131 28 21 J. microcarpa JMOP 2 56 40 71 3S 14 201 184 92 257 224 87 J. microcarpa x Serr JMS 3 74 69 93 JMS 4 127 113 89 JMS 5 99 91 92 JMS 5A 114 98 86 JMS 7 124 112 90 JMS 9 144 95 66 JMS 11 132 114 86 JMS 11A 170 149 88 JMS 12 101 81 80 JMS 13 90 70 78 JMS 15 77 65 84 JMS 18 103 83 81 JMS 19 116 104 90 JMS 20 145 122 84 California Walnut Board 116 Walnut Research Reports 2012

Table 2. (cont.) Greenhouse survival of rooted clonal microshoots 2012 # Rooted # Survived % Survival JMS 21 114 77 68 JMS 24 90 74 82 29 JM 1 229 208 91 29 JM 2 89 77 87 29 JM 3 177 154 87 29 JM 4 101 85 84 29 JM 5 232 132 57 29 JM 8 145 121 83 29 JM 10 97 71 73 29 JM 11 81 73 90 29 JM 12 97 82 85 29 JM 17 218 173 79 29 JM 22 80 63 79 STJM 4 105 96 91 STJM 6 111 99 89 STJM 7 90 66 73 STJM 11 151 116 77 3823 3133 82 J. cathayensis x Serr JCS 1 20 17 85 JCS 2 15 9 60 JCS 3 100 79 79 JCS 6 25 21 84 3S 3 1 1 100 3S 5 42 36 86 3S 8 3 3 100 3S 9 3 3 100 3S 13 1 1 100 3S 16 2 2 100 3S 17 66 50 76 3S 18 41 32 78 3S 19 1 1 100 3SH 2 19 18 95 3SH 3 1 1 100 3SH 4 4 3 75 3SH 5 1 1 100 3SH 6 8 6 75 3SH 7 25 23 92 3SH 8 8 8 100 386 315 82 California Walnut Board 117 Walnut Research Reports 2012

Table 2. (cont.) Greenhouse survival of rooted clonal microshoots 2012 # Rooted # Survived % Survival Serr x J. cathayensis #21 SC 1 7 1 14 SC 5 16 13 81 SBB7 62 50 81 SM 1 39 24 62 SM 5 27 18 67 S 1 28 20 71 S 2 1 1 100 S 8 25 18 72 S 10 25 18 72 S 13 12 7 58 S 15 21 19 90 S 17 3 3 100 S 21 11 6 55 S 25 4 2 50 S 32 5 2 40 S 35 20 11 55 S 38 13 7 54 S 39 27 13 48 S 40 17 5 29 S 42 8 8 100 S 45 4 4 100 S 49 11 3 27 S 54 5 3 60 S 58 6 3 50 S 59 11 7 64 S 61 26 16 62 434 282 65 95 007 13 x J. cathayensis #21 7 1 8 5 63 7 2 24 21 88 7 3 14 10 71 7 4 13 6 46 7 10 19 17 89 7 11 2 2 100 7 13 9 7 78 7 14 10 10 100 99 78 79 California Walnut Board 118 Walnut Research Reports 2012

Table 2. (cont.) Greenhouse survival of rooted clonal microshoots 2012 Transgenics # Rooted # Survived % Survival L 33 80 60 75 L 66 60 26 43 L 194 28 22 79 L K 66 62 94 L J 99 82 83 L U 101 88 87 L V 61 39 64 J1 L 55 43 78 J1 1A 93 58 62 J1 19A 160 125 78 J1 20A 105 77 73 RR4 1A 123 99 80 RR4 8A 39 28 72 RR4 10A 73 49 67 RR4 11A 21 18 86 RR4 12A 75 44 59 J1a control 123 94 76 RR4 control 65 51 78 1427 1065 75 Total 9628 7378 77 California Walnut Board 119 Walnut Research Reports 2012

Table 3 (part 1). Allocation and distribution in 2011-2012 of clonal walnut plants produced for rootstock pathogen resistance testing. Genotype Browne Phytophthora Tests McKenry Nematode Tests Kluepfel Crown Gall Tests Controls AX1 125 PX1 108 RX1 152 16 20 Vlach 0 16 20 VX211 27 16 20 W17 black 94 Wingnut 10.05 100 Nematode selection RX032 35 Transgenics and controls L 33 10 8 L 66 10 8 L 194 10 8 L K 10 8 L J 10 8 L U 10 8 L V 10 8 J1 L 10 8 J1 1A J1 3A J1 19A 5 J1 20A 5 J1a control 17 RR4 1A RR4 4A 4 RR4 6C RR4 10A RR4 11A 5 RR4 12A RR4 control 5 California Walnut Board 120 Walnut Research Reports 2012

Table 3 (cont.). Allocation and distribution in 2011-2012 of clonal walnut plants produced for rootstock pathogen resistance testing. Genotype Browne Phytophthora Tests McKenry Nematode Tests Kluepfel Crown Gall Tests J. microcarpa x Serr JMS 3 63 8 12 JMS 4 25 8 12 JMS 5 57 8 12 JMS 5A 67 8 12 JMS 7 76 8 12 JMS 9 50 8 12 JMS 11 50 8 12 JMS 11A 50 8 12 JMS 12 54 8 12 JMS 13 53 8 12 JMS 15 51 8 12 JMS 18 13 8 12 JMS 19 53 8 12 JMS 20 50 8 12 JMS 21 50 8 12 JMS 24 70 8 12 29JM 1 34 8 12 29JM 2 60 8 12 29JM 3 50 8 12 29JM 4 77 8 12 29JM 5 25 8 12 29JM 8 50 8 12 29JM 10 79 8 12 29JM 11 54 8 12 29JM 12 48 8 12 29JM 17 50 8 12 29JM 22 47 8 12 STJM 4 53 8 12 STJM 6 80 8 12 STJM 7 30 8 12 STJM 11 73 8 12 California Walnut Board 121 Walnut Research Reports 2012

Table 3 (cont.). Allocation and distribution in 2011-2012 of clonal walnut plants produced for rootstock pathogen resistance testing. Genotype Browne Phytophthora Tests McKenry Nematode Tests Kluepfel Crown Gall Tests J, cathayensis x Serr JCS 1 8 12 JCS 2 7 JCS 3 9 8 12 JCS 6 8 12 JCS 7 4 6 3S 3 3S 5 8 8 3S 8 3S 9 3 3S 10 1 3S 12 6 3S 13 1 3S 16 1 3S 17 8 8 3S 18 8 8 3S 19 1 3SH 2 8 11 3SH 3 1 3SH 4 3 3SH 5 1 3SH 6 6 3SH 7 8 7 3SH 8 4 Serr x J. cathayensis #21 SC 1 1 SC 5 7 SBB7 8 8 SM 1 8 8 SM 5 8 4 S 1 8 S 2 S 8 8 S 10 3 California Walnut Board 122 Walnut Research Reports 2012