Hop Virus Elimination Protocol

This protocol is based on a published method using thermotherapy and apical meristem culture (Postman, DeNoma, and Reed 2005). In trialing this method, our aim was to record a detailed procedure to make the method more accessible to producers who may not have a fully equipped tissue culture laboratory. Additionally, we wished to test the success rate for virus elimination using cuttings larger than the 0.5 mm size used in the published protocol, since larger cuttings have a greater survival rate and are less technically challenging to take.

Stock plants: Virus-infected plants for hop cultivars Brewers Gold, Centennial, Crystal, Golding, Mt. Hood and Cascade were maintained as stock plants for virus elimination experiments. Plants were grown in a greenhouse with a 16 hour photoperiod at 25 C (77 F). Potting medium was PRO-Mix HP (Premier Tech Horticulture), with up to 20% extra perlite added. Plants were fertilized weekly with Peters 20-10-20 (ICL Fertilizers) and supplemented with iron HEDTA (Jay-Mar, Inc) as needed. Leaf tissue for each variety was tested for Apple Mosaic Virus (ApMV) by ELISA (Agdia), and carlaviruses (Hop mosaic virus, Hop latent virus, American hop latent virus) by PCR (Plant Disease Diagnostic Clinic, University of Wisconsin-Madison). Test results are shown in Table 1.

Table 1: Testing results for hop cultivars used as stock plants for virus eradication experiments

The infected stock plants were clonally propagated as follows. Single node cuttings were taken from established bines. The stem end closer to the root was dipped in rooting hormone (0.1% IBA, e.g. Bontone Rooting Powder, Bonide), and inserted into potting medium in plug trays. Tools were decontaminated between cuttings by soaking for 5 minutes in a 1% Alconox/1% SDS solution and then rinsed in water. Cuttings were transplanted into three inch pots after 3-4 weeks and were maintained in greenhouse conditions for at least another 4 weeks (more usually 6-8 weeks), and up to 16 months. We noted that older clones with well-established roots, a woody base to the bines, and bushy bine growth were more resilient to thermotherapy treatment than younger clones. Several days prior to thermotherapy, excessive bine growth was trimmed to leave four to five nodes per bine. At least 24 hours prior to thermotherapy, a soil drench of Safari (Dinotefuran, Valent) was applied to control fungal gnats, since these pests would otherwise thrive under thermotherapy conditions with detrimental effects on the hop plants.

Thermotherapy chamber design: Thermotherapy growth chambers were constructed from plywood and insulated with one inch expanded polystyrene (EPS) foam with plastic facing for easy cleaning. Internal dimensions were 22 inches by 10 inches by 20 inches tall, to allow for a single standard 1020 nursery flat. Chambers had Plexiglass tops under light fittings for two fluorescent bulbs (Cool White) and were equipped with a heating cable under a plastic grating on which the plants were placed. A thermostat and timer were used to control the heat and light cycles. Note that the placement of the heating cable at the base of the chambers resulted in a temperature gradient, and an alternative placement to reduce the temperature gradient through the chamber may be desirable. Additionally, ventilation of chambers may be desirable to control humidity and reduce the potential for mold growth.

Thermotherapy and apical cutting culture: Plants were subjected to four hour cycles of light at 38 C (100.4 F) alternating with darkness at 30 C (86 F), for 14-16 days. At the end of the thermotherapy period, a clean scalpel was used to remove a 1-inch section from the end of each bine. Scalpels were decontaminated in 70% ethanol between cuttings, to avoid transferring virus particles that could be present in bines. Cuttings were surface sterilized as follows: up to five cuttings from the same plant were placed into a labeled tube containing 70% ethanol, shaken briefly and allowed to sit for five minutes. Following steps were conducted in a sterile laminar flow hood, with tools such as forceps, dissecting needles and tweezers sterilized in 70% ethanol between uses. Using long forceps, cuttings were transferred to a labeled tube containing a solution of bleach-Tween20 (see Media and Solutions for preparation details), shaken briefly and allowed to sit for at least 5 and no more than 10 minutes. Cuttings were transferred to three successive washes in sterile water.

Apical cuttings were dissected under a dissecting microscope (10x-40x) using 18G needles and small tweezers (such as pointed eyelash tweezers). The dissecting microscope was wiped down with 70% ethanol before being placed in the sterile laminar flow hood. To provide a size reference, a small piece of graph paper was taped to the microscope stage. Autoclaved glass petri dishes were used to provide a sterile work surface for dissection. Apical cuttings were transferred from the final sterile water wash to the sterile petri dish. Leaf primordia were removed using an 18G needle mounted to a handle, and a sterile 18G needle was used to detach a 2-5 mm explant from the tip of the cutting. Dishes were sprayed with 70% ethanol as required to avoid dehydration of small explants; brief exposure to 70% ethanol did not negatively impact explant survival. Explants were transferred to SM media containing 2% glucose and 0.1% BAP (see Media and Solutions) in 50 mm plates; multiple explants from the same originating plant were placed on the same plate. To avoid crushing the explants, they were placed into small slits in the media surface made using the tweezer points.

Explants were placed in growth chambers at 25 C (77 F) with a 14 hour photoperiod. After 1-4 weeks, explants showing shoot growth were transferred to individual baby food jars containing SM media with 2% glucose, 0.1% BAP, and 0.1% IBA. After one to four weeks, plants that showed root development and shoot growth were transferred to potting media. For the first 2-4 days, new transplants were kept under a paper cup and sprayed with water daily to reduce transplant shock. Survival rates are shown in Table 2; note that most mortality occurred in the first 10 days after explant excision.

Retesting for virus infection: Once plants had 2-3 fully expanded leaves, leaf tissue from each plant was tested for Apple Mosaic Virus (ApMV) by ELISA (Agdia), and carlaviruses (Hop latent carlavirus, American hop latent carlavirus) by PCR (Plant Disease Diagnostic Clinic, University of Wisconsin-Madison). Test results are shown in Table 2.

Conclusions: The limiting step in virus elimination was the low survival rate of explants, averaging 10% across varieties. Surviving plants had a high rate of virus eradication: 100% for Apple Mosiac Virus, and 95% for carlaviruses. Larger explants are expected to have a higher survival rate, but are more likely to contain cells still infected with viruses after thermotherapy. It may be worthwhile to trial explants larger than the 2-5 mm range used in this study to find a balance between explant survival and successful virus eradication.

Media, solutions and other materials

Bleach-Tween-20 Solution: Prepare a 0.1% Tween-20 solution by adding 0.1 ml Tween-20 to 99.9 ml of distilled water. Autoclave for 20 minutes to sterilize. Using aseptic technique in a laminar flow hood, dispense 9.5 ml amounts to sterile tubes. One tube will be required for each set of 5 cuttings from a single plant. On the day that the sterilization process will take place, add 1 ml of bleach to each tube. (Alternatively, add 10.5 ml bleach to 100 ml of 0.1% Tween-20 and dispense approximately 10 ml amounts into sterile tubes.) To ensure full hypochlorite activity, do not add bleach to the Tween-20 solution ahead of time, and ensure the bleach bottle was first opened less than 3 months previously.

SM media: Dissolve 4.33 g Murashige-Skoog Salts (Phytotech Labs), 1 ml 1000x MS vitamins (Phytotech Labs), and 20 g glucose in 950 ml distilled water. Adjust pH to 5.6 with potassium hydroxide, and adjust volume to 1000 ml. Add Phytoblend Agar (Caisson Laboratories) at a rate of 7 g per liter for plates and 9 g per liter for baby food jars. Autoclave for 20 minutes and allow to cool to 50 C. Add 20 µl BAP stock and (for baby food jars only) 20 µl (IBA) 5 mg/ml stock. Using aseptic technique in a sterile laminar flow hood, immediately dispense media into 50 mm plates (8-10 ml per plate) or baby food jars (55 ml per jar).

Plant growth regulators:

BAP 5 mg/ml stock – dissolve 50 mg 6-benzylaminopurine (Phytotech Labs) in 10 ml of 1 M sodium hydroxide. Filter sterilize and store at -20C.

IBA 5 mg/ml stock – dissolve 50 ml of indole-3-butyric acid (Phytotech Labs) in 1 ml of ethanol; once dissolved, adjust volume to 10 ml with 1 M sodium hydroxide. Filter sterilize and store at -20C.

 General notes on tools and preparation of sterile materials

If an autoclave is not available, a standard pressure cooker is an effective substitute (Anon. 1984; “Sterilizing Liquids” n.d.). Use all normal precautions to avoid injury. Glass test tubes are useful for preparing surface sterilization solutions for apical cuttings. Solutions can be dispensed into glass tubes for autoclaving, or empty glass tubes can be sterilized by autoclaving and sterile solutions can be dispensed into them using aseptic technique in a sterile laminar flow hood. Glass test tubes, caps and petri dishes can be obtained from many scientific supply companies and scientific surplus outlets. Straight and curved fine “eyelash” tweezers can be considerably less expensive than scientific forceps and are well suited to handling small explants.

References

Anon. 1984. “Using a Pressure Cooker as an Autoclave.” EPI Newsletter 6 (6): 5–8.

Postman, J.D., J.S. DeNoma, and B.M. Reed. 2005. “DETECTION AND ELIMINATION OF VIRUSES IN USDA HOP (HUMULUS LUPULUS) GERMPLASM COLLECTION.” Acta Horticulturae, no. 668 (February): 143–48. https://doi.org/10.17660/ActaHortic.2005.668.18.

“Sterilizing Liquids.” n.d. Accessed October 7, 2020. https://teach.genetics.utah.edu/content/microbiology/liquids/.