1. Plant growth conditions

1. Grow Arabidopsis thaliana plants in a short-day light regime with 9 h of light at 22 °C and 15 h of dark at 20 °C and an irradiance of 120 µmol·m⁻²s⁻¹ (measured as the photosynthetically active radiation, PAR at the canopy level, see Table of Materials).

2. Preparation of spore inoculum

NOTE: For details, see Fuchs et al.¹⁴.

1. Homogenize two to three frozen galls from Chinese cabbage plants (Brassica rapa var. pekinensis cultivar "Granaat") in a blender containing 300 mL of autoclaved distilled water and filter through four layers of sterile gauze.
2. Centrifuge the filtrate (at 6,000 x g, for 5 min, and at 4 °C) and mechanically remove the starch layer from the spore pellet using a spatula. Repeat this process until most of the starch is removed.
3. Determine the spore concentration using a hemocytometer¹³ by counting the number of spores in 20 different field areas.
4. Ensure that the final concentration of spore suspension used for inoculating is 1 x 10⁶ spores·mL⁻¹ for Arabidopsis thaliana. Inoculate each plant 20 days after germination with 2 mL of calibrated spore suspension.

3. Tissue preparation and fixation

1. Prepare the plant tissue.
   1. Carefully remove soil from the root systems of clubroot-infected and non-infected plants. Clean thoroughly with water and collect hypocotyls and galls (length between 0.5-2 cm) into microcentrifuge tubes.
2. Perform PFA fixation (4% paraformaldehyde, PFA in 1x PBS + 0.01% Triton X-100) following the steps below.
   1. Add 4 g of PFA powder (see Table of Materials) to 100 mL of PBS (phosphate-buffered saline, pH 7.4, Table 1) by stirring at 65 °C (do not boil). Use KOH (10 M and 1M) dropwise to obtain a clear solution with a pH of around 11.
   2. Adjust the pH with H₂SO₄ to bring it down to 6.9 and add 0.01% (v/v) Triton X-100 to improve fixation. Aliquot the solution to store at −20 °C (do not refreeze) or store at 4 °C.
3. Perform tissue fixation.
   1. Fix the samples in 200-500 µL of PFA fixative for 1 h at room temperature by applying a constant vacuum (700 mbar) using a vacuum pump. Ensure that the object is completely immersed in the PFA solution.
      ​NOTE: Samples can be stored for a few weeks at 4 °C (fridge) and used later for sectioning.

4. Tissue embedding and sectioning

1. Use 4% w/v agarose for agarose embedding of non-infected hypocotyls and infected galls. Boil the solution to dissolve agarose and pour it when it cools down while it is still viscous.
2. Dry the object slightly on a tissue for a few seconds to remove excess PFA.
3. Use toothpicks/forceps to carefully embed and orient the plant object in agarose. To prevent oblique sections, ensure that the object is oriented and molded correctly so that its axis is perpendicular to the plane of the vibratome blade (for radial sections).
4. To accelerate agarose solidifying, place the Petri or multi-culture plate at 4 °C for 10 min.
   ​NOTE: If the object is too large (as in the case of larger galls of Arabidopsis at 21 days post-inoculation [DPI]), the object can be sectioned even without embedding in agarose.

5. Vibratome sectioning

NOTE: The vibratome is equipped with a vibrating razor blade to cut through plant organs/tissue. The vibration speed, amplitude, angle of the blade, and section thickness are all parameters that can be adjusted (see Table of Materials).

1. Using a blade, carefully cut and mold the object within an agarose block, noting the preferred orientation for sectioning.
2. Stick the agarose block/large gall to properly mount it on the specimen holder using cyanoacrylate/instant glue and masking tape (see Table of Materials).
3. For Arabidopsis hypocotyls and galls (early stages), keep the thickness of sections between 30-40 µm for obtaining good quality images.
   NOTE: For analyzing the later stages of gall progression, the thickness of sections can be between 50-80 µm.
4. Refrain from cutting thinner sections as it can destroy the gall sample due to the vibrations of the moving blade.
5. Adjust the thickness for sections, speed, and vibration amplitude according to the gall's thickness and size (40 µm or 60 µm).
6. Add distilled water to the water bath and begin with sectioning.
7. Carefully collect sections using forceps or brush and transfer them into a microcentrifuge tube containing 1mL of 1x PBS buffer (pH 7.4).
   ​NOTE: Typically, the higher the vibration amplitude, the better the sectioning quality. For non-infected hypocotyls or galls that do not contain mature resting spores, the vibration amplitude can be kept at 1.2 mm with 0.60 mm·s⁻¹ speed. In case of large galls with partial loss of cellular integrity and visible signs of spore maturation, the vibration amplitude should be decreased to 0.55 mm with 0.45 mm·s⁻¹ speed.

6. Specimen clearing

1. Remove 1x PBS from the microcentrifuge tube and add 200-500 µL of clearing solution (Table 1).
2. From now on, store the samples at room temperature in the dark. Ensure that the objects are always submerged in the clearing solution at all times.
3. Replace the clearing solution with the new one if its color changes after some time during sample processing.
   ​NOTE: The color of the clearing solution can turn yellowish due to sample processing. In such a case, replacing the solution is recommended to improve the clearing of tissues.

7. Staining procedure

1. For Calcofluor white staining for the cell walls (for outlaying plant cells), prepare 5% v/v of Calcofluor white stain (see Table of Materials) in the clearing solution. Stain for at least 5 min in the dark.
2. For staining lipids with Nile Red (for staining resting spores and oil droplets in infected cells), prepare 1 mg/mL of the stock (see Table of Materials) in clearing solution. Dilute it further to obtain 1:99. Stain for at least 10 min in the dark.
3. For Basic Fuchsin lignin staining, prepare 0.2% w/v of the stain (see Table of Materials) in clearing solution. Stain for at least 10 min in the dark.
   ​NOTE: During double staining, samples are first stained with Nile Red for 10 min before Calcofluor white application. The excess staining solutions were removed after each step. If the object seems overstained, remove the excess stain by washing with the clearing solution. Dilute stains using the clearing solution, if required. Always stain the sample with Nile Red/Basic Fuchsin before Calcofluor staining. Staining first with Calcofluor can prevent proper staining of the spores by Nile Red.

8. Microscopy

1. Mount cleared sections on the microscopy slide and observe under an epifluorescence or a confocal microscope (see Table of Materials). Use clearing solution as the mounting medium to prevent drying of the sample.
2. Use multiple acquisition modes for imaging more than one fluorescence spectrum simultaneously.
   NOTE: The excitation/emission spectra used in the presented protocol are as follows: for Nile Red 553/636 nm, for xylem autofluorescence 380/475 nm, for Basic Fuchsin 561/650 nm, for Calcofluor white 405/475 nm, for erRFP 585/608 nm, and for GFP 488/509 nm (Table 2).