CLONAL PROPAGATION OF WALNUT ROOTSTOCK GENOTYPES …walnutresearch.ucdavis.edu/2012/2012_97.pdf ·...

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.


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.


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.

Figure 4. The suture side of the walnut shell in contact with vice clamps

Vice clamp

Suture side of walnut shell Cheek of walnut shell


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 NH4NO3 (mg/l) Ca(NO3)2 (mg/l) Zn(NO3)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 CaCl2 from the medium. DKW includes CaCl2 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 CaCl2 was tested using genotypes AX1, RX1, Chandler, W17, wingnut ‘WNxW’, and J. cathayensis x Serr ‘JCS1’. Removal of CaCl2 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.


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.


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.


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 3rd 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 2nd 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.


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


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).


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.


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, inches1 Midday light interception2, %

Avg. in-shell yield3, lbs/a

Rootstock Dec-09 Dec-10 Nov-11 Nov-12 2011 2012 2011 2012 Own-rooted Chandler 2.7 a4 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 (J. hindsii X J. regia)

2.4 bc 3.7 bcd 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 1Measured 24 inches above soil line; 2Measured 29 July 2011 and 8 September 2012; 38% Wet basis moisture content; 4Values 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


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.


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.


Table 1 (part1). Ex vitro rooting percentage for rootstock genotypes.

#Attempted #Rooted %Rooted

PDSParadox

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

WNxW10.05b 464 371 80

CLRVTolerant

WIP3 153 94 61

English

Chandler 617 76 12

J.microcarpa

JMOP2 76 59 783S14K3K5

3253223

2302316

717270

456 328 72

J.microcarpaxSerr

JMS3 144 104 72

JMS4 207 156 75

JMS5 89 82 92

JMS5A 129 118 91

JMS7 158 134 85

JMS9 187 150 80

JMS11 208 173 83

JMS11A 229 188 82

JMS12 117 83 71

JMS13 147 102 69

JMS15 137 98 72

JMS18 156 125 80


JMS19 169 125 74

Table 1 (cont.). Ex vitro rooting percentage for rootstock genotypes. #Attempted #Rooted %Rooted

JMS20 228 146 64

JMS21 156 93 60

JMS24 110 86 78

29JM1 270 227 84

29JM2 91 65 71

29JM3 195 165 85

29JM4 76 67 88

29JM5 361 226 63

29JM8 169 133 79

29JM10 99 68 69

29JM11 99 83 84

29JM12 126 109 87

29JM17 246 196 80

29JM22 124 109 88

STJM4 126 110 87

STJM6 182 115 63

STJM7 164 88 54

STJM11 153 103 67

5052 3827 76

J.cathayensisxSerr

JCS1 32 20 63

JCS2 16 14 88

JCS3 131 94 72

JCS6 36 34 94

JCS7 1 0 0

3S3 8 4 50

3S5 63 50 79

3S6 2 2 100

3S8 3 3 100

3S9 17 12 71

3S10 8 1 13

3S12 2 0 0

3S13 1 1 100

3S16 3 2 67

3S17 95 60 63

3S18 79 50 63

3SH1 1 0 0

3S19 8 1 13


3SH2 44 32 73

Table 1 (cont.). Ex vitro rooting percentage for rootstock genotypes. #Attempted #Rooted %Rooted

3SH3 3 1 33

3SH4 11 4 36

3SH5 5 2 40

3SH6 17 12 71

3SH7 77 40 52

680 447 66SerrxJ.cathayensis#21

SC1 42 2 5

SC5 77 22 29

SBB7 218 57 26

SM1 169 37 22

SM5 229 29 13

S1 93 13 14

S8 184 13 7

S10 113 8 7

S13 78 11 14

S15 131 3 2

S16 11 4 36

S17 23 1 4

S21 114 14 12

S25 127 1 1

S32 7 5 71

S35 69 6 9

S38 38 5 13

S39 73 15 21

S40 57 5 9

S42 69 3 4

S45 54 1 2

S49 22 1 5

S54 25 1 4

S58 55 5 9

S59 31 8 26

S61 78 26 33

2187 296 14


Table 1 (cont.). Ex vitro rooting percentage for rootstock genotypes. UC95‐007‐13xJ.cathayensis#21


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 19Transgenic


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

J11A 385 220 57

J119A 433 298 69

J120A 260 129 50

RR41A 198 137 69

RR48A 87 75 86

RR410A 176 111 63

RR411A 56 44 79

RR412A 147 102 69

J1acontrol 399 247 62

RR4control 143 108 76

2402 1558 65

Total 18868 11099 59


Table 2. (Part 1) Greenhouse survival of rooted clonal microshoots – 2012 #Rooted #Survived %Survival

PDSParadox

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

WNxW10.05b 309 280 91

CLRVTolerant

WIP3 57 28 49

English Chandler 131 28 21

J.microcarpa

JMOP2 56 40 71

3S14 201 184 92

257 224 87J.microcarpaxSerr

JMS3 74 69 93

JMS4 127 113 89

JMS5 99 91 92

JMS5A 114 98 86

JMS7 124 112 90

JMS9 144 95 66

JMS11 132 114 86

JMS11A 170 149 88

JMS12 101 81 80

JMS13 90 70 78

JMS15 77 65 84

JMS18 103 83 81

JMS19 116 104 90

JMS20 145 122 84


Table 2. (cont.) Greenhouse survival of rooted clonal microshoots – 2012 #Rooted #Survived %Survival

JMS21 114 77 68

JMS24 90 74 82

29JM1 229 208 91

29JM2 89 77 87

29JM3 177 154 87

29JM4 101 85 84

29JM5 232 132 57

29JM8 145 121 83

29JM10 97 71 73

29JM11 81 73 90

29JM12 97 82 85

29JM17 218 173 79

29JM22 80 63 79

STJM4 105 96 91

STJM6 111 99 89

STJM7 90 66 73

STJM11 151 116 77

3823 3133 82J.cathayensisxSerr

JCS1 20 17 85

JCS2 15 9 60

JCS3 100 79 79

JCS6 25 21 84

3S3 1 1 100

3S5 42 36 86

3S8 3 3 100

3S9 3 3 100

3S13 1 1 100

3S16 2 2 100

3S17 66 50 76

3S18 41 32 78

3S19 1 1 100

3SH2 19 18 95

3SH3 1 1 100

3SH4 4 3 75

3SH5 1 1 100

3SH6 8 6 75

3SH7 25 23 92

3SH8 8 8 100

386 315 82


Table 2. (cont.) Greenhouse survival of rooted clonal microshoots – 2012 #Rooted #Survived %Survival

SerrxJ.cathayensis#21

SC1 7 1 14

SC5 16 13 81

SBB7 62 50 81

SM1 39 24 62

SM5 27 18 67

S1 28 20 71

S2 1 1 100

S8 25 18 72

S10 25 18 72

S13 12 7 58

S15 21 19 90

S17 3 3 100

S21 11 6 55

S25 4 2 50

S32 5 2 40

S35 20 11 55

S38 13 7 54

S39 27 13 48

S40 17 5 29

S42 8 8 100

S45 4 4 100

S49 11 3 27

S54 5 3 60

S58 6 3 50

S59 11 7 64

S61 26 16 62

434 282 6595‐007‐13xJ.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


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

J11A 93 58 62

J119A 160 125 78

J120A 105 77 73

RR41A 123 99 80

RR48A 39 28 72

RR410A 73 49 67

RR411A 21 18 86

RR412A 75 44 59

J1acontrol 123 94 76

RR4control 65 51 78

1427 1065 75

Total 9628 7378 77


Table 3 (part 1). Allocation and distribution in 2011-2012 of clonal walnut plants produced for rootstock pathogen resistance testing.

Genotype

BrownePhytophthora

Tests

McKenryNematode

Tests

KluepfelCrown

GallTests Controls

AX1 125

PX1 108

RX1 152 16 20

Vlach 0 16 20

VX211 27 16 20

W17black 94

Wingnut10.05 100

Nematodeselection

RX032 35

Transgenicsandcontrols

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

J11A

J13A

J119A 5

J120A 5

J1acontrol 17

RR41A

RR44A 4

RR46C

RR410A

RR411A 5

RR412A

RR4control 5


Table 3 (cont.). Allocation and distribution in 2011-2012 of clonal walnut plants produced for

rootstock pathogen resistance testing.

Genotype

BrownePhytophthora

Tests

McKenryNematode

Tests

KluepfelCrownGall

Tests J.microcarpaxSerr

JMS3 63 8 12

JMS4 25 8 12

JMS5 57 8 12

JMS5A 67 8 12

JMS7 76 8 12

JMS9 50 8 12

JMS11 50 8 12

JMS11A 50 8 12

JMS12 54 8 12

JMS13 53 8 12

JMS15 51 8 12

JMS18 13 8 12

JMS19 53 8 12

JMS20 50 8 12

JMS21 50 8 12

JMS24 70 8 12

29JM1 34 8 12

29JM2 60 8 12

29JM3 50 8 12

29JM4 77 8 12

29JM5 25 8 12

29JM8 50 8 12

29JM10 79 8 12

29JM11 54 8 12

29JM12 48 8 12

29JM17 50 8 12

29JM22 47 8 12

STJM4 53 8 12

STJM6 80 8 12

STJM7 30 8 12

STJM11 73 8 12


Table 3 (cont.). Allocation and distribution in 2011-2012 of clonal walnut plants produced for rootstock pathogen resistance testing.

Genotype

BrownePhytophthora

Tests

McKenryNematode

Tests

KluepfelCrownGall

Tests J,cathayensisxSerr

JCS1 8 12

JCS2 7

JCS3 9 8 12

JCS6 8 12

JCS7 4 6

3S3

3S5 8 8

3S8

3S9 3

3S10 1

3S12 6

3S13 1

3S16 1

3S17 8 8

3S18 8 8

3S19 1

3SH2 8 11

3SH3 1

3SH4 3

3SH5 1

3SH6 6

3SH7 8 7

3SH8 4 SerrxJ.cathayensis#21

SC1 1

SC5 7

SBB7 8 8

SM1 8 8

SM5 8 4

S1 8

S2

S8 8

S10 3


Table 3 (cont.). Allocation and distribution in 2011-2012 of clonal walnut plants produced for rootstock pathogen resistance testing.

Genotype

BrownePhytophthora

Tests

McKenryNematode

Tests

KluepfelCrownGall

TestsS15 8

S17 2

S21 6

S35 6

S38 1

S39 9

S40 4

S42 6

S45 3

S49 3

S54 2

S58 1

S59 2

SC5 5 12

SBB7 8 2

SM1 3 11

SM5 8

95‐007‐13xJ.cathayensis#21

7‐1 6

7‐2 8

7‐3 5

7‐4 4

7‐9 6

7‐10 8

7‐11 3

7‐12 1

7‐13 6

7‐14 6

J.microcarpa

3S14 35 8 12

JMOP2 29 8 12

JMOP3 3

K3 8

K5 8

TotalPlantsAllocated 2321 714 706


Budding Experiment Results

Table 5: Estimated Average Water Use: (ml/per day/ plant)

Shade Bench Coldmls 69 108 44

Table 9: Rootstock Caliper take not total %BelowMedian 86 64 150 57Abovemedian 92 58 150 61

Table 11: Scion Take Scion Take not total %CGR 124 57 181 68.5TOP 15 16 31 48.4GH 39 49 88 44.3

Table 12: Bud Position Budposition take not total %

Top 63 49 112 56

Bottom 115 73 188 61

Table 6: Pot

Pottype take not total %Tall 158 95 253 63

Round 26 21 47 55

Table 7: Scion Caliper and Maturity

ScionAvg

CaliperAvg

MaturityAvg

SlippingCGR 6.5 2.7 2.8GH 5.5 3.6 3.5TOP 6.0 3.9 3.9

Table 4: Temperature/Water Treatment Treatment take not total %Bench/dry 20 13 33 61Bench/mid 20 14 34 59Bench/wet 28 6 34 82Cold/dry 16 22 38 42Cold/mid 18 17 35 51Cold/wet 11 15 26 42Shade/dry 20 15 35 57Shade/mid 23 13 36 64Shade/wet 21 8 29 72

Total 177 123 300 59

Table 8: Rootstock CharacteristicsRootstock score take not total %Slipping 1 0 1 1 0 2 5 21 26 19 3 70 42 112 63 4 101 63 164 62 5 2 0 2 100Maturity 1 0 0 0 NA 2 15 25 40 38 3 129 70 199 65 4 42 27 69 70 5 2 0 2 100

Table 10: Scion Characteristics Scion score take not total %Slipping 1 0 0 0 NA 2 45 20 65 69 3 83 49 132 63 4 50 52 102 49 5 0 1 1 0Maturity 1 1 0 1 100 2 48 15 63 76 3 85 58 143 59 4 38 48 86 44 5 6 1 7 86


Table13.Effectofmicronutrientmixornickelonwalnutmouseear/littleleafsymptoms Treatment %withsymptomsControl 47.8Micromax 26.1Micromax+Ni 0Nickel 0ANOVAforheightSource DF SumofSquares FRatio Prob>FClone 2 1975.9713 30.7323 <.0001Treatment 3 310.8858 3.2235 0.0274Clone*Treatment 6 361.6608 1.875 0.0964

TreatmentHeight(cm)

Micromax+Ni 14.4aMicromax 13.6abControl 11.2abNi 9.5a ANOVAfordiameterSource DF SumofSquares FRatio Prob>FClone 2 1.117515 0.2825 0.7547Treatment 3 17.852227 3.0081 0.0356Clone*Treatment 6 12.650368 1.0658 0.3908

TreatmentDiameter(mm)

Micromax+Ni 5.55aControl 5.31abMicromax 5.10abNi 4.28b

Clone

Height(cm)

RX1 18.3aVlach 11.7bVX211 6.9c


Table 14. Interaction of clone and chemical amendment on plant height


Table 15. Galling response of seedling of wingnut genotype DPTE 1.09 and a Paradox control following stab inoculations with Agrobacterium strain EC1 in 2011 and 2012.

GenotypeSeedlingNumber

YearEval.

No.ofStabs

MeanGallScore

MeanGall

volume

Gallvol./stemdiameterratio

Wingnut 1 2011 4 1.0 0 0DPTE1.09 2012 4 1.0 0 0

3 2011 4 1.0 0 0 2012 4 1.0 0 0

4 2011 4 1.0 0 0 2012 4 1.1 9 0

5 2011 4 1.0 0 0 2012 4 1.0 0 0

13 2011 4 1.4 0 0 2012 4 1.1 1 0

2 2011 4 3.0 1813 215 2012 4 1.8 1745 114

14 2011 4 3.5 3648 264 2012 4 1.6 500 34

8 2011 4 2.8 22569 904 2012 4 1.9 823 54

6 2011 4 3.0 25480 1004 6 2012 4 1.9 454 28

ParadoxRR4 2011 13 2.4 524 120Control 2012 4 4.0 11017 1360


Table 16. Current Clonal Rootstock Field Trials.

County Grower Genotypes Date

Established Comments Tehama

H. Crain RX1, VX211, Vlach, Paradox sdlg.

2009 Budded Sept 2009 New orchard Scion: Howard

Butte

Deseret RX1, AZ2

2006 New orchard Scion: Chandler

Lake Valadez VX211, Vlach, Paradox seedlings

2011 Nematode Scion: VX211 and Vlach are grafted to Chandler; seedlings will be grafted to Chandler in 2012

Lake Valadez (Suchan)

VX211, Vlach, Paradox seedlings

2011 Unfumigated new planting Nematode site Scion: black walnut scion for nursery trees, so will be Chandler interstems on the VX211 and Vlach

Lake VX211, Vlach, Paradox sdlg.

2012 No fumigation, walnut to walnut; will be budded to Chandler in 2013

Lake VX211, Vlach, Paradox sdlg.

2012 No fumigation, pear, fallowed, then to walnut; will be budded to Chandler in 2013

Glenn Anderson Vlach, Paradox sdlg 2006 Grafted to Howard Sutter/Yuba

Conant RX1,VX211, Serr, Vlach 2007 New orchard Spot fumigated old Hartley site

Sutter/Yuba

Conant RX1, VX211, Vlach 2009 Power-line planting Scions: Ivanhoe, Howard, Sexton, 91-077-40 New orchard

Sutter/Yuba

Double Nut

VX211, AZ2, NZ1, JX2 2005 Phytophthora Replants DONE

Sutter Ruzich RX1 bare root and containerized, VX211 bare root, Paradox sdlg.

2011 Phytophthora Replicated trial in a P. cinnamomi site To be grafted

Solano

Cilker RX1, VX211, Burbank, Vlach, Paradox sdlg.

2009 New orchard Budded fall 2009 Scion: Tulare

Yolo Turkovich RX1, VX211 2011 New Orchard Grafted: 2011 Scion: Forde,

Contra Costa

Tennant WIP2, WIP3, clonal Sunland and Vina, Paradox sdlg.

2005 Blackline tolerant New orchard Scion: Vina

Contra Costa

Maggiore WIP3, clonal Chandler, Paradox sdlg., WIP1, WIP2

2011 Blackline tolerant New orchard Scion: Chandler

San Joaquin Dondero RX1, VX211, AZ2, 2005 Phytophthora


County Grower Genotypes Date

Established Comments

NZ1, JX2 Replants DONE

San Joaquin

Taylor WIP3, WIP5, WIP6 2005 Blackline tolerant New orchard Scion: Chandler

San Joaquin

Lagorio (Concar)

RX1, VX211, WIP3, AZ025, Vlach June-bud, Vlach-grafted, Chandler own-rooted, Paradox sdlg.

2007 New orchard Scion: Chandler

San Joaquin

Chiappi RX1, Paradox seedlings 2010 Phytophthora plot Replanted block Scion: Serr

Calavaras

Gotelli RX1, VX211 2007 Phytophthora, water table Replants DONE

Stanislaus

MJC Clonal Vina and Chandler, Vlach, 84-121, Sunland sdlg., Px sdlg.

1999 Planted 1999 Grafted 2000 New orchard

Kings

Verboon VX211, Px sdlg, with Vlach buffer

2008 Fumigation trial treatments New orchard Scion: Tulare


Appendix 1. Laboratories Licensed to Produce Liners of UC Clonal Rootstock Selections

Acemi Nursery

License: RX1, VX211

Agromillora – Gridley

License: RX1, VX211

California Seed and Plant LabTest agreement: Px1, WIP3

Duarte Nursery

License: RX1, VX211

Test agreement: WIP3

North American Plants

License: RX1, VX211

Test agreement: WIP2, WIP3, WIP6

ProTree License: VX211, RX1

Tissue Grown Corporation

License: VX211, RX1

V-Tree

License: VX211, RX1

Test Agreement: WIP3

VitroTech

License: RX1, VX211

Test Agreement: WIP2, WIP3


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