Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

This study was carried out in line with the guidelines in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocol was approved by the University of Texas at Austin Animal Care and Use Committee (AUP-2021-00254).

1. Designing the sgRNA for CRISPR mutagenesis

1. Obtain the exon sequence containing the targeted loci.
2. Design a synthetic guide RNA (sgRNA) with a protospacer adjacent motif (PAM) site that is specific to the Cas endonuclease.
   1. If co-injecting a donor construct, design the sgRNA to have a predicted cut site as close to the desired edits as possible.
      NOTE: APE is a free software with a tool to find Cas9 sgRNA sites¹¹. Alternatively, tools such as CHOPCHOP can be used, which offers an online interface for designing sgRNAs that considers the different PAM sites of Cas9 and other Cas endonucleases¹².
3. Design forward and reverse primers to amplify the region surrounding the designed CRISPR/Cas cut site. The PCR product should be approximately 200-400 base pairs (bp) in length to be able to resolve small indels on a gel later in the protocol.
   NOTE: CHOPCHOP¹¹ automatically designs primers for every sgRNA it outputs.
4. Optional: Design a DNA donor construct with the desired edit(s).
   1. Obtain the genomic sequence of the exon containing the loci of the desired mutation.
      NOTE: Intronic sequences can vary between individuals, so it is best to avoid the inclusion of intronic sequences in the donor construct as this could interfere with the homology-dependent repair pathway.
   2. If a specific amino acid change is desired, alter the specific codon accordingly.
      NOTE: When making these changes, be aware of the codon frequency. If possible, choose a codon with high usage.
   3. To prevent CRISPR/Cas digestion of the donor construct, make additional synonymous mutations that (i) disrupt the PAM and/or (ii) disrupt the sgRNA sequence, with higher preference being placed on making non-synonymous changes closer to the PAM site.
   4. Use a restriction site finder (APE¹¹ also has this feature) to find unique restriction sites (those that only occur once) in the predicted PCR product of the wild-type sequence.
      1. Repeat this process for the predicted PCR product that reflects the successful integration of the donor construct.
      2. Compare the list of restriction sites between the wild-type and donor-integrated sequences to find a restriction site that is present only in the edited sequence.
      3. If no unique restriction site is found, continue making synonymous changes within the donor construct until this is achieved.
         NOTE: If possible, continue making synonymous mutations that would disrupt the sgRNA binding to the donor construct.
   5. Trim the donor sequence to a manageable size (50-100 bp) with at least 20 bp of homology surrounding the predicted CRISPR/Cas cut site and edits, if possible.
5. Obtain the designed sgRNA and/or DNA donor oligonucleotides, as well as any primers required for genotyping.
   NOTE: To prevent the digestion of the DNA donor oligonucleotides by nucleases, it is recommended to modify the first three phosphate bonds on the 5' and 3' ends of the DNA oligonucleotides with phosphorothioate.
6. Inject 1 nL of the injection mix into one-cell stage embryos. Typically, a standard injection mix contains the following:
   1 µL of 25 µM locus-specific sgRNA
   1 µL of 25 µM Cas9 endonuclease
   ​Optional: 1 µL of 3 µM DNA donor construct
   1. Bring the total volume to 5 µL with 0.1 M RNase-free KCl.
7. Allow the F0-injected individuals to develop into sexually mature adults. The remaining protocol focuses on screening the F0-injected males for germline edits.

2. Setting up the breeding tanks

1. Prime F0 males for mating by setting up breeding tanks with dividers between the F0 males and females.
   1. To minimize the number of breeding tanks required, set up three males across from two females in each tank.
      1. Set up as many F0 males as can effectively be screened and housed as individuals after sperm collection. The genotype of the females is not important since their gametes will not be collected during this procedure.
   2. Set up one breeding tank of wild-type males across from females. Wild-type sperm will serve as a control for the interpretation of the results later in the procedure.
2. Incubate the prepared breeding tanks overnight.

3. Preparing the materials for the sperm collection procedure

1. Prepare enough anesthetizing solution (0.016% tricaine-HCl in fish water) to fill a small glass bowl.
2. Dispense 40 µL of 50 mM NaOH into 0.2 mL PCR tubes. Prepare one tube for each sperm sample that will be collected. Store on ice.
3. Prepare individual 0.8 L tanks for each fish that will be sampled during the procedure. These tanks will house the sampled fish until the genotyping is complete.
   NOTE: This is the primary factor that limits the number of fish that can be effectively screened in this protocol. Ensure adequate space for housing the sampled individuals until the genotyping is complete.
4. Moisten a 1-inch x 1-inch sponge (cut with a shallow, oval-shaped divot to hold a male zebrafish ventral side up during the procedure) in fish water and squeeze out excess liquid.
   1. Position the sponge in the field of view of a stereomicroscope at low magnification (Figure 1A).
   2. Position oblique overhead lighting for optimal viewing.

4. Anesthetizing the male fish and collecting the sperm

1. Prepare a clean capillary tube.
   1. Gently shake the tube container to release a single capillary tube.
   2. Place the capillary tube into the provided bulb, and set it aside.
2. Transfer a male fish into the prepared anesthetizing solution (0.016% tricaine-HCl in fish water).
   1. Gently stir the solution with a slotted spoon to speed up the process of anesthesia.
   2. Once the operculum (gill) movement slows down (approximately 1-2 min), proceed with the sperm collection.
3. Use the slotted spoon to transfer the fish onto a clean stack of paper towels.
   1. Use the slotted spoon to gently roll the fish to remove excess water.
   2. Gently blot the fish with clean, folded tissue paper to remove any remaining water.
4. Use the slotted spoon to transfer the fish to the prepared sponge (Figure 1B).
   1. Position the fish ventral side up with its head nearest the dominant hand of the person performing the procedure.
   2. Gently blot the area around the anal fins with clean, folded tissue paper.
5. Collect the sperm.
   1. Using the end of the capillary tube, gently move the pelvic fins laterally away from the midline to expose the cloaca (Figure 1C).
   2. Position the end of the capillary tube near the cloaca.
   3. Using the forceps, gently squeeze the sides of the fish beginning just below the gills and ending at the cloaca (Figure 1C').
   4. Collect sperm into the capillary tube by capillary action. Approximately 1 µL is sufficient for this procedure (Figure 1D).
      NOTE: Some labs report using p10 pipet tips instead of capillary tubes for sperm collection.
   5. If sperm is not released with gentle pressure, return the fish to the system water.
      1. Monitor for recovery from the anesthesia.
      2. Do not squeeze the fish hard to expel sperm, as this can injure the fish.
      3. Rest the fish for 2 weeks before attempting to collect sperm again.
   6. After collection, place the fish into a clean isolation tank with system water for individual housing.
      1. Monitor the fish for recovery from the anesthesia.
      2. Rest the fish for 2 weeks before attempting to collect sperm again.
   7. Place the capillary tube with the collected sperm into the prepared PCR tube (with 40 µL of 50 mM NaOH).
   8. Squeeze the rubber bulb to expel the collected sample.
   9. Dispose of the used capillary tube into an approved glass waste container.
   10. Incubate the samples on ice until all the males have been squeezed.
6. Repeat steps 4.1-4.5 for each individual male.
7. Label the isolation tanks and sperm samples such that the final sperm genotyping results can be traced back to the corresponding individual with putative mutations.
8. Monitor all the individuals, and make sure they are upright and exploring their tanks before putting the tanks back on the system.
9. Put the females used for priming back into their respective tanks on the system.
10. If any fish does not recover from the anesthesia after 10 min, follow the euthanasia procedure approved by the institution associated with the lab.
    1. Example: Rapid chill the individual in an ice bath with fish water that is 2-4 °C.
    2. Dispose of the euthanized fish in an appropriate animal carcass waste container according to the procedure approved by the institution associated with the lab.

5. Extracting DNA from the sperm samples

1. Briefly spin down all the sperm samples in a minicentrifuge, and place PCR tubes into a thermal cycler.
2. Close the lid of the thermal cycler.
3. Run the following settings in the thermal cycler:
   1. Heat the samples for 40 min at 95 °C.
   2. Cool the samples to 25 °C.
4. Remove the samples from the thermal cycler.
5. With a clean pipette tip for each sample, neutralize the pH by adding 10 µL of 1 M Tris-HCl (buffered to pH 8).
6. Mix well by pipetting up and down.
7. Briefly spin down the samples in a minicentrifuge.
8. Genomic DNA samples can be stored for several days at 4 °C or placed at −20 °C for up to 6 months.

6. PCR amplification (and/or restriction digest) of the desired locus

1. Amplify the desired locus using a standard PCR protocol.
   1. Preheat the thermal cycler to the initial denaturation temperature in the PCR reaction below.
   2. Prepare a 25 µL reaction mixture for each sperm sample in PCR tubes:
      12.5 µL of 2x Taq Polymerase Master Mix
      1.5 µL of 10 µM forward primer
      1.5 µL of 10 µM reverse primer
      4.5 µL of nuclease-free water
      5 µL of neutralized sperm DNA sample
   3. Mix well by pipetting up and down.
   4. Briefly spin down the samples in a minicentrifuge.
   5. Place the samples in the pre-heated thermal cycler with the following settings:
      Initial denaturation: 95 °C for 3 min
      35 cycles of denaturation (95 °C for 30 s), annealing (55 °C for 30 s), and extension (72 °C for 30 s)
      Final extension: 72 °C for 5 min
      NOTE: It may be necessary to adjust the extension temperature and/or time depending on the specific Taq polymerase and the expected length of the PCR amplification product.
   6. Remove the PCR samples from the thermal cycler.
2. Optional: Perform the donor-specific restriction digest on a small volume of the PCR product, leaving at least 5-10 µL of undigested PCR product for gel electrophoresis.
   1. Prepare a 30 µL reaction in 0.2 mL PCR tubes:
      10 µL of unpurified PCR product
      15 µL of nuclease-free water
      3 µL of 10x restriction enzyme buffer
      2 µL of restriction enzyme (BstNI is used in the representative results for analyzing the dnah10 knock-in allele)
      ​NOTE: Many restriction enzymes are still active in a typical PCR reaction buffer; however, the addition of restriction digest buffer can help improve the results. Refer to specific manufacturer instructions for troubleshooting if any problems occur.
   2. Mix well by pipetting up and down.
   3. Briefly spin down the samples in a minicentrifuge.
   4. Place the samples in a thermal cycler with the following settings:
      1. Cut the PCR amplicon at 60 °C for 1 h.
         NOTE: The cut temperature and incubation time may need to be adjusted for the specific restriction enzyme being used.
      2. Inactivate the restriction enzyme with the required temperature and incubation time, if indicated by the manufacturer.
         ​NOTE: The BstNI enzyme does not require heat inactivation.

7. Performing gel electrophoresis to separate PCR amplicons of varying sizes

1. Prepare 500 mL of 0.5x TBE running buffer: add 50 mL of 10x TBE buffer concentrate to 450 mL of deionized water.
2. Prepare 100 mL of 4% agarose gel in 0.5x TBE running buffer: add 4 g of agarose with 10 µL of 10,000x gel stain to 100 mL of 0.5x TBE running buffer. Microwave until the agarose powder is fully dissolved.
   1. Pour the gel solution into an appropriately sized gel casting frame, and insert the comb on one side of the gel.
      NOTE: A larger gel size (recommended: 15 cm x 15 cm) is advantageous for this procedure, as it provides the amplicons with more space to resolve on the gel.
      1. If a restriction digest product is to be analyzed, it is best to run it on the same gel as the PCR product so that the results can be compared. Adjust the comb well size as necessary to fit all the samples on the gel.
   2. Leave the gel at room temperature until solidified. Carefully remove the comb. Carefully remove the gel from the casting frame.
3. Set up the gel electrophoresis rig.
   1. Position the gel in the electrophoresis rig such that the wells are nearest the negative electrode. Pour 0.5x TBE running buffer into the electrophoresis rig until the wells are fully submerged.
4. Load the gel.
   1. Load 5 µL of DNA ladder into the first well (choose a DNA ladder that contains DNA fragments that are similar in size to the PCR amplicon).
   2. Using a new pipette tip for each sample (including the wild-type control sample), load 5-10 µL of the PCR product into the remaining wells.
   3. Optional: If a restriction digest product is to be analyzed, load these samples onto the gel as well.
      NOTE: It is best to load equal amounts of DNA between the PCR and the restriction digest product. For example, in the 30 µL restriction digest reaction mixture, 10 µL of PCR product was added. Therefore, the DNA concentration of the 15 µL of digest product was comparable to the 5 µL of unpurified PCR product.
5. Run the gel.
   1. Connect the electrodes to the gel electrophoresis rig, and apply 150 V for at least 1 h to ensure adequate separation of the amplicons.
   2. Image the gel to view the DNA bands.
      ​NOTE: Strongly mutagenized CRISPR-targeted loci will run as multiple band sizes representing a combination of insertion/deletion alleles with the wild-type amplicons.
      1. If the gel bands are not resolved after 1 h, apply 150 V to the gel for 15 min intervals until the bands are sufficiently separated on the gel.
      2. Optional: If screening for the insertion of a precise edit such as an SNP, epitope tag, or fluorescent tag, screen the sperm DNA with PCR methods that are specific to the desired edit. For example, when identifying a donor-specific restriction site, an additional band will be present in the restriction digest product compared to the PCR product of the sperm sample.

8. Isolating germline stable alleles

1. After 2 weeks of rest from the sperm squeezing procedure, outcross the F0 individuals whose sperm genomic DNA contained the desired edits.
   NOTE: Sperm generation from individual spermatogonia clones may be cyclical in zebrafish; thus, the desired allele might not be producing sperm at the time of the outcross. For this reason, additional crossing may have to be performed more than once to isolate the allele implicated by the initial screening.
2. Once the F1 outcrossed fish are sexually mature, proceed with standard genotyping methods (for example, fin-clip DNA) using the same PCR and gel electrophoresis protocol performed during the screening of the sperm genomic DNA.
   1. Sanger sequence the PCR products to identify the F1 individuals with the same desired edit.
   2. Optional: If the desired edit introduces or disrupts a restriction site, screen with an appropriate restriction digest protocol.
3. In-cross F1 heterozygotes with the same desired edit.
4. Screen the F2 progeny for the expected phenotype and/or genomic edits.