Transforming, Genome Editing and Phenotyping the Nitrogen-fixing Tropical Cannabaceae Tree Parasponia andersonii

Parasponia is a tropical tree from the Cannabis family that is able to establish a nitrogen-fixing symbiosis with rhizobium. Comparative studies between Parasponia and legumes could provide insight into the core genetic networks underlying rhizobium symbiosis. With this protocol, Parasponia andersonii stable transgenic mutant lines can be generated in a period of two to three months.

These lines can be efficiently propagated through in vitro propagation. This protocol provides an experimental platform to study rhizobial symbiosis as well as other aspects of the biology of this tropical tree. Basic skills in tissue culture and molecular cloning are sufficient to implement this protocol.

This allows any laboratory to adapt Parasponia as research model. Demonstrating the procedure will be PhD students Titis Wardhani and Yuda Roswanjaya together with Marijke Hartog, a technician in our lab. To grow Parasponia andersonii trees, begin by germinating the seeds.

Remove the seeds from Parasponia berries by rubbing against the inside of a tea sieve or squashing them on a piece of tissue paper. Disinfect seeds using 4%hypochlorite bleach for 15 to 20 minutes, and then wash the seeds six times with sterilized water. Transfer the seeds to sterile 200 microliter PCR tubes, and fill the tubes with sterilized water to submerge the seeds.

Incubate the tubes for 10 days in a thermocycler according to manuscript directions. After the 10-day incubation, transfer the seeds to SH-0 plates, and close them with two layers of elastic sealing foil to prevent drying. Incubate at 28 degrees Celsius with a 16-hour-day, eight-hour-night cycle for three to four weeks.

Once the seedlings develop their first set of true leaves, transfer them to pots filled with commercial potting soil, and cover them with a translucent plastic cup to prevent desiccation. Place the pots in a 28-degree Celsius, 85%relative humidity climate room or greenhouse under a 16-hour-day, eight-hour-night regimen. After one week, remove the plastic cup.

Regularly water the pots, and supplement with fertilizer to sustain tree growth. Prepare for transformation by harvesting young five-to eight-centimeter branches from the greenhouse-grown trees, making sure to use only healthy, noninfected branches. Cut off the leaves, leaving one centimeter squared of leaf tissue at the end of each petiole.

Disinfect the tissue for 15 minutes with 2%hypochlorite bleach containing a few drops of polysorbate 20, and then rinse it eight times with autoclaved water. Re-suspend Agrobacterium cells in 25 milliliters of infiltration medium to an optical density of about five. Cut the stem and the petiole tissue into one-centimeter pieces inside the cell suspension.

Remove the leaf tissues and incubate the stem and petiole pieces in the cell suspension for 10 to 30 minutes. Dry the tissues pieces on a sterile piece of filter paper and place them on rooting medium prepared according to manuscript directions. Incubate the plates in the dark at 21 degrees Celsius for two days.

After the two days, inspect the plates for fungal or bacterial contamination and discard contaminated plates. Transfer the tissue pieces to 10 milliliters of SH-10 medium prepared according to manuscript directions. Leave the tissue in the medium for at least 10 minutes, and gently agitate every two to three minutes to wash.

Prepare rooting medium with cefotaxime and kanamycin according the manuscript directions, and pour plates. Dry the tissues on sterile pieces of filter paper, and transfer them to the plates. Incubate plates for seven days at 28 degrees Celsius under a 16-hour-day, eight-hour-night regimen.

Check plates for fungal or bacterial contamination every two days. If contamination occurs, transfer the noninfected tissue pieces to a fresh plate. After seven days, transfer the tissue pieces to propagation medium prepared according to manuscript directions and incubate at 28 degrees Celsius under a 16-hour-day, eight-hour-night regimen.

Refresh the plates once a week until transgenic shoots develop, making sure to only transfer noninfected tissue pieces to new plates. Once the putative transgenic shoots are at least one centimeter long, cut the shoots and culture them independently in propagation medium with antibiotics. To ensure that the shoots represent independent transformants, take only a single shoot from each side of an explant.

Propagate the tissue by placing about 10 shoots on a fresh plate of propagation medium and closing the plate with two layers of elastic sealing foil. Incubate the plates at 28 degrees Celsius under a 16-hour-day, eight-hour-night regimen, and repeat this step every four weeks. When the shoots reach one centimeter in length, cut them at their base and place them on the rooting medium.

Position the shoots upright by inserting the basal tip of the shoot into the medium. About 10 shoots can be placed on a single plate. Start by preparing rhizobium inoculum.

Inoculate 10 milliliters of liquid YEM medium from a single colony of Mesorhizobium plurifarium BOR2, and incubate at 28 degrees Celsius for two days. Use this culture to inoculate a larger volume of liquid YEM medium. Once grown, centrifuge the bacterial culture for 10 minutes at 3, 500 times g to harvest the cells.

Re-suspend the bacterial pellet in liquid EKM medium, and determine the optical density. Prepare three liters of EKM medium, which is enough for about 20 pots, and inoculate it with the rhizobial suspension. Mix the medium with 1.25 kilograms of perlite, and add 210 grams of this mixture to sterile translucent polypropylene pots.

Plant one to three Parasponia andersonii plantlets to each pot, and prepare several pots with plantlets transformed with the control construct. Weigh several of the pots, and cover the bottom of each pot to shield the roots from light exposure. Incubate the pots in a climatized growth room at 28 degrees Celsius under a 16-hour-day, eight-hour-night regimen for four to six weeks.

Once a week, weigh several pots to determine water loss. If the water loss exceeds 10 milliliters, supplement with ultra-pure water to compensate for the loss. After the incubation, clean the roots from perlite, and determine nodule numbers using a binocular.

This technique has been used to successfully transform petioles and segments of Parasponia andersonii stems with Agrobacterium strain AGL1. First, the tissue explants are co-cultivated with Agrobacterium for two days. Prolonged co-cultivation results in bacterial over-colonization and should be prevented.

A few weeks later, small green micro-calli are observed along the original wound surface. These calli continue to grow and develop one or more putatively-transformed shoots at six to eight weeks after the start of transformation. Transformation efficiencies vary from 10 to 30%for tissue explants from mature branches to 65 to 75%for tissue explants from young and rapidly growing branches.

Transgenic shoots can be multiplied through in vitro propagation, which gives rise to tens of shoots within a month. These shoots can be placed on rooting medium that will induce root formation in about two weeks and yield plants that can be used for experimentation. When starting a stable transformation experiment, it is important to select healthy branches as source material.

Parasponia T0 transgenic lines are propagated in vitro. Therefore, it is essential to thoroughly genotype Parasponia transgenic lines. The establishment of Parasponia as an experimental model allows comparative analysis between two distantly related nodulating lineages, Parasponia and legumes.

This could identify conserve genetic networks underlying rhizobium symbiosis. Parasponia can also be a valuable model to study other research topics such as wood formation, the development of bisexual flowers, or the biosynthesis of Cannabaceae-specific secondary metabolites.