Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

1. Generation of Deletion Constructs

1. Generate a plasmid containing the deletion construct (Figure 1) comprising a respective 1 kb upstream flank (UF) and downstream flank (DF) each and the appropriate resistance cassette flanked by blunt cutting restriction sites.
   NOTE: Any cloning strategy can be used (for cloning see reference ²²) we recommend Golden Gate cloning for such plasmids ⁹.
2. Excise the deletion construct from the plasmid using a blunt cutter to obtain 1 µg DNA of the deletion construct ⁹. Purify the deletion construct to eliminate enzyme and buffer ²² e.g. by using commercial reagents and follow the manufacturer's instructions.
   NOTE: The vector backbone can remain in the transformation mixture.

2. Preparation of Protoplasts

NOTE: Keep sterile conditions during all times of the experiment. The cell wall is a strong protective barrier that limits access of molecules to the plasma membrane. To allow uptake of the DNA containing the deletion construct, the cell wall needs to be removed by cell wall degrading enzymes in a protoplasting reaction. Critical steps in protoplast preparation are 1) osmotic stabilization of the medium and 2) avoiding mechanical stress on the protoplasts.

1. Mark 2 ml tubes with a round bottom to freeze the protoplasts and cool them down at -20 °C. Start a pre-culture in 3 ml YEPS-Light from a fresh plate of the strain with the desired genetic background. Incubate on a rotating wheel for 24 hr at 28 °C.
   NOTE: The strain can be the solopathogenic strain SG200 ¹⁰, wildtype strains, tester strains such as AB33 ²³, or any mutant background for generation of double mutants. Make sure the strain is free of the desired resistance.
2. Dilute the culture in 50 ml YEPS-Light in a baffled flask and incubate on an orbital shaker at 28 °C with 200 rpm.
   NOTE: The doubling time is about 2 hr for wildtype strains. Allow three duplications minimum.
3. Grow until exponential phase. Ensure that the optical density, measured at a wavelength of 600 nm (OD₆₀₀), is around 0.8 (0.6 to 1.0 is acceptable). An OD₆₀₀ of 1.0 corresponds to about 1 to 2 x 10⁷U. maydis cells per ml.
4. Check the cells for contamination under a microscope. Use 40x magnification. Pellet the cells of the complete 50 ml culture for 5 min, 1,500 x g and discard the supernatant. Resuspend the pellet in 25 ml sodium citrate, sorbitol solution (SCS), centrifuge 5 min, 1,500 x g, and discard supernatant.
   NOTE: Bacterial contaminations can be identified as small and often moving cells. Fungal contaminations can be identified based on a different cell shape or size compared to the cigar shaped cells of U. maydis.
5. During centrifugation prepare protoplasting solution (12.5 mg/ml Trichoderma lysing enzymes, in SCS; prepare 3 ml per cell pellet; filter sterilize through a 22 µm filter; solution has to be fresh for optimal enzymatic activity). SCS is an osmotic stabilizer to prevent rupture of protoplast upon removal of the cell wall.
6. Resuspend the pellet in 2 ml protoplasting solution. Incubate for 5 to 20 min at RT and check under the microscope until 30 to 40% of the cells are round or like pinheads (Figure 2).
   NOTE: Perform all steps on ice from now on and handle protoplasts with care!
7. Wash 3x in 10 ml cold (4 °C) SCS, centrifuge at 1,000 x g for 5 min.
8. Resuspend the pellet in 10 ml cold sorbitol, TrisHCl, CaCl₂ solution (STC), spin at 1,000 x g for 5 min, discard supernatant. Resuspend the pellet in 1 ml cold STC, make 100 µl aliquots in the cooled tubes. Freeze the aliquots at -80 °C until further use.

3. Transformation of U. maydis

NOTE: The transformation of U. maydis protoplasts relies on the polyethylene glycol (PEG)-mediated transformation method, which is technically simple and opposed to electroporation or biolistic transformation does not require specialized equipment.

1. Prepare the two-layered selection plates (2 plates per transformation reaction). The bottom layer contains the antibiotic in a doubled concentration. Pipet 12 ml of RegLight containing 400 µg/ml hygromycin (or 300 µg/ ml nourseothricin or 4 µg/ml carboxin or 1 mg/ml G418) into a Petri dish. Avoid bubbles.
   NOTE: Bottom layer plates may be stored a couple of days at 4 °C, but prepare the top layer freshly during transformation (see below).
2. Thaw protoplasts on ice (1 tube/transformation). Add 1 µl heparin (15 mg/ml). Add 1 µg DNA (linearized construct) or 10 µl H₂O (water control). Incubate for 10 min on ice.
3. During this time, for the upper layer spread 12 ml liquefied RegLight (boil and cool to 60 °C) onto the RegLight bottom plate.
   NOTE: This layer does not contain antibiotics.
4. Add 500 µl STC/PEG (polyethylene glycol) to the transformation tube (3. 2) and mix carefully by inverting the tube. Incubate 15 min on ice.
5. Distribute each transformation reaction on two of the two-layered RegLight plates. Spread carefully and slowly with a glass pipet. Incubate plates upright at 28 °C for 5 to 7 days until transformants grow as colonies. Obtain about 100-200 colonies per plate.
   NOTE: Lower numbers might be caused by "bad" protoplasts that either contain too much cell wall and therefore cannot take up DNA or cannot regenerate. The former can be tested by transformation with a self-replicating plasmid, the latter by testing regeneration on plates without antibiotics.
6. Prepare YEPS-Light plates containing the appropriate antibiotic at 200 µg/ml hygromycin (or 150 µg/ ml nourseothricin or 2 µg/ml carboxin or 500 µg/ml G418; these concentrations equal half of the one used for bottom plates). Re-purify at least 24 putative transformant colonies on these plates. Single colonies should be obtained. At the same time streak out the original strain on non-selective media.
   NOTE: Plates can be stored at 4 °C for 1-2 weeks.

4. Verification of Correct Deletion Events

NOTE: The mutations are verified in a three-step procedure (Figure 1). First, candidates that contain the wildtype copy of the gene are excluded by PCR. Second, the integration of the resistance cassette into the locus is verified by PCR. Third, the integration of the resistance cassette only into the desired locus is verified by Southern blot analysis. Careful strain verification is essential, so that mutant phenotypes truly correlate with the deletion.

1. First Diagnostic PCR ²⁰
   1. Design primers pairing in the gene to be deleted that result in a product of 250-600 bp (Figure 1). Such small products are easy to amplify in a colony PCR, and long enough for detection in standard gel electrophoresis.
   2. For each candidate and the original strain as a positive control, resuspend a little bit (amount of a pinhead) of cell material of a single colony with a toothpick in 20 µl 0.02 M NaOH. Incubate at RT for 30 min.
      NOTE: Use fresh colonies. If the plates are older than 1 week re-streak to obtain a fresh colony.
   3. Use 1 µl of the cell suspension for the PCR immediately. Run a standard PCR reaction and analyze resulting fragments.
      NOTE: These samples cannot be stored.
      NOTE: This PCR tests for presence of the wildtype gene and thereby allows rapid identification of negative candidates in which the gene of interest has not been replaced. Clones yielding bands in this PCR reaction will be discarded. One such negative clone should anyway be carried along as an additional negative control. Candidates without a PCR product have to be analyzed further. For genes without detrimental effects approximately half of the candidates should contain the wildtype gene, while the other half should not result in a band. This would correspond to a homologous recombination rate of 50%.
2. Second Diagnostic PCR
   1. Preparation of genomic DNA.
      NOTE: Genomic DNA (gDNA) can be prepared using two different protocols. The first involves toxic reagents like phenol and chloroform and hence, should be handled only by experienced researchers (described in 4.2.1.1 to 4.2.1.5) while the second can also be handled by less experienced students (described in 4.2.1.6 to 4.2.1.11). Both protocols are working reliably. Step 4.2.1.1 is identical for both protocols.
      1. Inoculate 5 ml YEPS-Light with each candidate and incubate on a rotating wheel for 24 hr at 28 °C. Drop 2 µl of the culture on a CM plate. Keep the culture sterile.
         NOTE: Following is a standard protocol (CAUTION: toxic steps).
      2. Put 1 scoop (~200 µl) of HCl-washed glass beads to 2 ml tubes. Add 2 ml of the U. maydis culture. Spin 12,000 x g for 5 min. Discard supernatant (pellet can be frozen at this stage). Add 500 µl Phenol/Chloroform/Isoamylalcohol (25:24:1) to the pellet. CAUTION! PHENOL IS TOXIC!
      3. Add 500 µl Usti-lysis-buffer 1. Shake for 6 to 10 min on a vibrax at 1,000 rpm. Verify that the cell pellet is completely resuspended, but avoid extended shaking to prevent shearing of the gDNA. Spin at 12,000 x g for 15 min. Transfer 400 µl of the aqueous phase into the new reaction tube (do not touch the interphase).
      4. Add 1 ml 100 % (v/v) ethanol. Invert the tube 3x to mix, observe the cloud of DNA that shortly appears. Spin at 12,000 x g for 5 min. Remove supernatant and wash with 200 µl 70% EtOH. Do not resuspend the pellet in this stage.
      5. Remove supernatant completely (do not let the pellet dry completely). Add 50 µl TE/RNase. Dissolve gDNA by shaking at 400 rpm at 50°C for 10 min. Analyze 2 µl of the gDNA on a 0.8 % agarose gel ²⁰.
         NOTE: The steps below comprise an alternative non-toxic protocol.
   2. Alternate preparation of genomic DNA
      1. Put 1 scoop (~200 µl) of HCl washed glass beads to 2 ml tubes. Add 2 ml of the U. maydis culture and spin at 12,000 x g for 5 min. Discard supernatant (pellets can be frozen at this stage).
      2. Add 500 µl Usti Lysis buffer 2 to the pellet. Shake for 5 to 15 min on a vibrax at 1,000 rpm. Verify that the cell pellet is completely resuspended.
      3. Incubate the tubes at 65 °C for 15 to 20 min. Using appropriate incubators designed to host 2 ml tubes is critical. Then, place it on ice for 5 min.
      4. Add 100 µl 8 M potassium acetate, vortex or invert 8 to 10 times. Spin 12,000 x g for 15 min at RT.
      5. Transfer 500 µl of the supernatant to a fresh 1.5 ml tube and add 400 µl isopropanol. Vortex or invert 8 to 10 times. Spin 12,000 x g for 15 min. Remove supernatant and wash with 500 µl 70% EtOH. Spin 12,000 x g for 5 min.
      6. Soak off supernatant completely! Eventually, add a short spin (10 sec) to remove residual liquid. Let the DNA pellet dry for 3 to 5 min. Add 50 µl TE/RNAase and incubate at 50 °C for 10 to 15 min at 400 rpm.
   3. PCR
      1. Design two primer pairs with one of them binding outside the flanks and the corresponding one binding within the resistance cassette (UF-f, RC-r; RC-f, DF-r, Figure 1) ⁹.
      2. Use 1 µl of a 1:10 dilution of the gDNA of every candidate as a template in 2 PCR reactions verifying the correct insertion upstream and downstream ⁹.
      3. Use standard PCR reactions for each candidate and analyze resulting fragments ²².
         NOTE: These PCRs test for the insertion of the resistance cassette in the correct genomic locus. If products are obtained for both sides, the candidate is likely a correct insertion. However, also candidates where only one side (upstream or downstream) could be confirmed should be carried along for further analysis in case that PCR reactions simply failed for one flank.
3. Verification of Correct Genomic Insertion by Southern Blot Analysis ²²
   1. Digest 15 µl of gDNA (4.2.1) of the mutant candidates and the corresponding wildtype/progenitor strain with an appropriate enzyme that cuts outside the locus/construct and in addition, cuts either in the gene or in the resistance cassette leading to clearly distinguishable patterns (Figure 1) ⁸.
      NOTE: None of the expected bands should be larger than 10 kb, so that they are clearly distinguishable from undigested DNA.
   2. Use the upstream and the downstream flank as probes. To label them for example use the PCR DIG labelling Kit or any similar standard method. Mix the labelled probes in a 1:1 molecular ratio and carry out the Southern blot. Keep at least two independent correct transformants.
      NOTE: The wildtype pattern should be clearly detectable in the original strain and the correct clones should contain only the expected bands. Additional bands only present in the candidates indicate other insertions of the construct in a wrong locus. Such clones should be discarded. The negative candidate identified in the first diagnostic PCR should contain the wildtype bands and the bands (unpredictable in size) of the wrongly integrated deletion construct.
4. Glycerol Stocks
   NOTE: Glycerol stocks serve to maintain the clones for years in strain collections. At least two independent transformants should be kept for each mutant. This allows comparing phenotypes that should be identical.
   1. Grow 5 ml cultures of the confirmed mutant strains in YEPS-Light on a rotating wheel for 24 hr at 28 °C Mix 500 µl of each culture with 500 µl NSY-Glycerol-medium. Invert tube until culture and medium are well mixed. Glycerol in the medium prevents the formation of ice crystals.
   2. Freeze at -80 °C. Re-streak a part of the frozen culture to test if the stock is functional.

5. Microscopic Phenotypes

1. Grow 5 ml cultures of wildtype and mutant strains in YEPS-Light on a rotating wheel for 12 hr at 28 °C.
2. Next day, dilute the culture to an OD₆₀₀ of about 0.1 and grow it till an OD₆₀₀  between 0.5 and 0.7. An OD₆₀₀ of 1.0 corresponds to about 1 to 2 x 10⁷ U. maydis cells per ml.
3. Inspect the cell morphology microscopically.
   NOTE: Healthy cells appear cigar shaped (Figure 4, WT). Pronounced vacuoles or filament formation indicate that the cells are stressed.
   NOTE: Depending on the expected mutant phenotype, the analysis can be adapted, e.g., if reporter strains are used the appropriate assay can be carried out at this stage. If cell morphology is changed statistical analysis should be conducted, e.g., to determine the mean cell length.

6. Infection Assay

NOTE: The seedling infection assay has been visualized previously in this journal ¹¹. It can either be carried out using a haploid, solopathogenic strain background such as SG200 ¹⁰ or by mixing compatible mating partners which both carry the mutation.

1. Grow at least 40 maize seedlings per strain for 7 days at a light cycle of 16 hr, 200 µE, 28 °C/22 °C (day/night). They should be at the 3-leaf stage and 10-15 cm tall at the day of infection.
2. Two days before infection, start a pre-culture of the strains in 5 ml YEPS-Light on a rotating wheel for 24 hr at 28 °C. Grow a main culture in 50 ml YEPS-Light up to OD₆₀₀ = 1.0 (cell division rate = 2 hr, allow 3 divisions minimum, can be done O/N).
3. Spin down the cells of 50 ml culture at 1,500 x g for 5 min and discard the supernatant. Wash 3x with sterile H₂O. Resuspend the cells in sterile H₂O to an OD₆₀₀ of 3.0. If necessary, mix the mating partners.
4. Inject 250 – 500 µl of the cell suspensions into the stem of 7 day old maize seedlings using a small syringe. Use sterile H₂O as a control. Keep infected seedlings for an additional 7-14 days in the same light and temperature conditions. Score the phenotype of each plant according to healthy, chlorosis, anthocyanin formation, small tumors, large tumors, dead plants ¹⁰.
   NOTE: Scoring can be done as early as 7 days and repeated at 14 days to follow infection over time.