Peanut Nodule Trapping Protocol

Made by Dr. George Colin diCenzo and lab

Step 1: Prepare seedlings.

• Prepare petri dishes with autoclaved 1X water agar (15 g/L of agar in ddH2O).
• Add the required amount of seeds a 50 mL conical tube, or larger container as required.
  • Prepare extra seeds in case not all seeds germinate.
• If using a 50 mL conical tube, add 40 mL (or other appropriate amount) of 95% ethanol to the seeds and gently shake by inverting the tube by hand for 30 seconds.
  • From this point forward, the tube containing the seeds should only be opened in a sterile environment.
  • The length of this treatment can be adjusted based on the plant type. The notes from my student say 30 seconds for Phaseolus vulgaris. My protocol for Medicago sativa (alfalfa) says 5 minutes.
• Pour out the ethanol.
• Add 40 mL (or appropriate volume) of autoclaved ddH2O to rinse.
• Pour out the autoclaved ddH2O.
• Repeat the previous two steps.
• Add 40 mL of 1% sodium hypochlorite (bleach, often sold as 6.5% hypochlorite) and invert by hand for 3.
  • The concentration and length can be adjusted as required. Navy bean was sensitive to bleach so could only tolerate 2 to 3 minutes. On the other hand, my protocol for alfalfa say 2.5% sodium hypochlorite for 20 minutes.
  • Note, if using seeds that need to be scarified with sulfuric acid, perform the scarification prior to the bleach treatment; the ethanol treatment may also not be required in this case.
• Pour out the sodium hypochlorite.
• Wash with sterile ddH2O and pour out the water. Repeat until bleach odor is undetectable.
• Using sterile tweezers, spread seeds on water agar plates (either 10 or 15 g/L of agar in ddH2O).
• Wrap the plates with aluminium foil to maintain darkness and leave at room temperature to germinate.
  • For Phaseolus vulgaris, this is ~3 days.

Step 2: Grow plants in the presence of soil or soil suspensions.

• Plant one germinated seedlings either directly in a pot containing the soil of interest, or in sterile vermiculite in Leonard assemblies (see details below).
  • For nodule trapping of bean rhizobia, we used vermiculite in Leonard assemblies.
  • Using Leonard assemblies is not necessary; it is how I have always grown my plants and there are advantages to using them, but if you do not have the required Magenta jars, it should be fine using other types of pots
• Place pots in the greenhouse and water as required.
• If seedlings are planted in vermiculite, after two to three days, inoculate the pots with a soil resuspension.
  • For bean, we mixed 10 g of soil with 50 mL of sterile ddH2O, then mixed 5 mL of this soil suspension with 20 mL of sterile ddH2O (25 mL total), and then add all 25 mL to one pot, pouring on and around the seedling.
• Grow plants for three to four weeks, at which point they should be harvested to collect nodules.

Step 3: Harvest nodules and isolate rhizobia.

• Uproot the plants and wash the roots to remove soil/vermiculite.
• Use tweezers to remove nodules from the roots of the plant (being careful not to squish the nodules), and place each nodule in a separate, sterile 1.5 mL tube.
• Wash nodules twice with 1 mL of ddH2O, and then discard the liquid.
• Add 1 mL of 1% sodium hypochlorite (bleach, often sold as 6.5% sodium hypochlorite) and incubate for 15 minutes with occasional inversion, to surface sterilize the nodules. Then discard the bleach.
• Add 1 mL of sterile YEM (or modified YM or TY + 300 mM sucrose; compositions of media are below), invert a few times to mix, and discard the YEM; repeat at least twice.
  • For Rhizobium and Sinorhizobium, any should work. For Bradyrhizobium, use YEM or modified YM. If unsure what you are isolating, YEM or modified YM is the safer choice.
• Add 100 to 200 µL of YEM (or modified YM or TY + 300 mM sucrose) and squish nodules using sterile pestles.
• Pipette 10 µL of cell suspension onto an agar plate, and then use sticks and dilution streak the cell suspension to get individual colonies.
  • Can use either TY or YMA plates (compositions of media are below)
  • For Rhizobium and Sinorhizobium, either work. For Bradyrhizobium, use YMA. If unsure what you are isolating, YMA is the safer choice.
• Incubate at 28˚C until colonies form.
• From each plate, pick a single colony and dilution streak on a new plate. Incubate at 28˚C until colonies form. From this new plate, pick a single colony and dilution streak on another fresh plate, and incubate. Repeat a third time.
• Grow a liquid culture of each strain starting from a single colony. Prepare -80˚C frozen stocks as either 15% glycerol stocks or 7% DMSO stocks.
  • E.g., mix equal volumes of the overnight culture with YEM (or TY) containing 30% glycerol.

Preparation of Leonard assemblies

• Prepare Leonard Assemblies by connecting two Magenta Jars with a cotton wick extending from the top jar into the bottom jar.
• Prepare a 1:1 (w/w) mixture of vermiculite and sand. Practically speaking, to a large autoclave bin add 16 L of vermiculite and 800 mL of sand, wet with ddH2O, and mix.
• Fill the top jar of each Leonard Assembly with the vermiculite/sand mixture until fill.
• Prepare 13 L of Jensen’s medium.
• Add 250 mL of Jensen’s medium to the top jar of each Leonard Assembly. Pour slowly to prevent overflowing.
• Add lids to each Leonard Assembly.
• Autoclave all Leonard Assemblies.

Jensen’s medium (1X)

Per 1 L of ddH2O, add:

• 1 g CaHPO4 (7.35 mM)
• 0.2 g K2HPO4 (1.15 mM)
• 0.2 g MgSO4 heptahydrate (0.811 mM)
• 0.2 g NaCl (3.42 mM)
• 0.17 g FeCl3 hexahydrate (0.616 mM)
• 1 mL of 1000X trace minerals solution
• pH adjust to 7.0

Jensen’s medium (4X)

When preparing large quantities of Jensen’s medium, it is often easier to prepare a 4X solution and dilute to 1X. To prepare a 4X solution, add the following to 4 L of ddH2O:

• 16 g CaHPO4
• 3.2 g K2HPO4
• 3.2 g MgSO4 heptahydrate
• 3.2 g NaCl
• 2.72 g FeCl3 hexahydrate or 1.63 g FeCl3 anhydrous Fe
• pH adjust to 7.0
• Add 1000X trace minerals solution after dilution to 1X (1 mL per 1 L of 1X solution)

1000X Trace Nutrients

Per 1 L of ddH2O, add:

• 1 g H3BO3 (16.2 mM)
• 1 g ZnSO4 heptahydrate (3.48 mM)
• 0.5 g CuSO4 pentahydrate (2 mM)
• 0.5 g MnCl2 tetrahydrate (2.53 mM)
• 1 g Na2MoO4 dihydrate (4.13 mM)
• 10 g Na2EDTA (26.9 mM)
• 2 g NaFeEDTA (5.45 mM)
• 0.4 g Biotin (1.64 mM)
• (My protocol doesn’t say this, but I believe we autoclave the 1000X trace nutrients)

YEM medium

• 10 g/L mannitol
• 1 g/L yeast extract
• 1.71 mM NaCl
• 1.87 mM K2HPO4
• 0.81 mM MgSO4
• Autoclave

Modified YM medium

• 5 g/L mannitol
• 0.5 g/L K2HPO4
• 0.1 g/L NaCl
• 0.2 g/L MgSO4 heptahydrate
• 0.4 g/L yeast extract
• 6.8-7 pH
• Autoclave

YMA composition

• 5 g/L mannitol
• 0.5 g/L K2HPO4
• 0.1 g/L NaCl
• 0.2 g/L MgSO4 heptahydrate
• 0.4 g/L yeast extract
• 6.8-7 pH
• 15 g/L agar
• Autoclave
• Prior to use, and after autoclaving, optionally add 2.5 mL of 1% Congo Red dye (filter sterilized and stored in the dark in the fridge). Congo Red is an indicator dye and helps distinguish bradyrhizobia from other rhizobia.

TY composition

• 5 g/L tryptone
• 2.5 g/L yeast extract
• Autoclave
• Prior to use, and after autoclaving, add CaCl2 to a final concentration of 10 mM

TY agar composition

• 5 g/L tryptone
• 2.5 g/L yeast extract
• 15 g/L agar
• Autoclave
• Prior to use, and after autoclaving, add CaCl2 to a final concentration of 10 mM