Determining the Role of Maternally-Expressed Genes in Early Development with Maternal Crispants

In studies leading to the development of this protocol, all zebrafish housing and experiments were approved by the University of Wisconsin-Madison Institutional Animal Care and Use Committee (IACUC-M005268-R2).

1. Synthesis of Guide RNAs

NOTE: Zygotic crispants have been created using a single guide RNA or multiplexing multiple guide RNAs to a single target^5,6,7,8,9,10. The multiplexing of guide RNAs increases the percentage of embryos showing a zygotic crispant phenotype¹⁰. Due to this increased frequency of embryos exhibiting a phenotype, maternal crispants are created by multiplexing four guide RNAs to a single gene. A more detailed protocol on using CHOPCHOP to design guide RNAs and an annealing method to synthesize guide RNAs for zebrafish can be found elsewhere^{16,17,18,19,20}.

1. To identify a maternally-expressed gene to target, ascertain the mRNA transcript levels during development via an RNA-sequence database that provides transcriptome information from zygote to 5 days²¹. In general, maternal-specific genes are highly expressed in the early embryo and are degraded after the zygotic genome is activated²².
2. Once a maternally-expressed target gene has been identified, determine the first predicted protein domain using the "domains and features" section available on the Ensembl genome browser²³. Use this domain as the target region for the four guide RNAs.
3. Use the guide RNA selection program CHOPCHOP to identify four guide RNA target sites in the first active domain. Design gene-specific oligonucleotides, as shown below for each target site. In the gene-specific oligonucleotide, the N²⁰ section corresponds to the target sequence minus the PAM site (NGG) from CHOPCHOP. Order these gene-specific oligonucleotides and the constant oligonucleotide using standard desalt purification (see Table of Materials).
   Gene-specific oligonucleotide:
   5' TAATACGACTCACTATA- N²⁰ -GTTTTAGAGCTAGAAATAGCAAG 3'
4. To create a guide RNA template for each gene-specific oligonucleotide, anneal it to the constant oligonucleotide and fill in the overhangs with T4-DNA polymerase as previously described¹⁶. After the four guide RNA templates are assembled, purify and concentrate them together using a DNA clean-up and concentrator kit according to the manufacturer's instructions (see Table of Materials).
5. Synthesize the sgRNA mixture from the pooled guide RNA template using an in-vitro T7 transcription kit (see Table of Materials). Perform the in-vitro transcription according to the manufacturer's instructions. Using half-reactions of the T7 Transcription kit can decrease the cost per reaction.
6. After RNA synthesis, purify the resulting pool of sgRNAs using an ethanol/ammonium acetate protocol as previously described^16,20,24. After the RNA has been isolated, resuspend it in 20 µL of nuclease-free water. If half-reactions of the T7 Transcription kit were used to transcribe the pool of sgRNAs, resuspend the purified RNA into 10-15 µL of nuclease-free water.
7. Quantify the amount of pooled sgRNAs that were created using a spectrophotometer. Dilute the pool of sgRNAs in nuclease-free water to a dilution of 1500 ng/µL ± 500 ng/µL. Typically, the final volume of the working dilution ranges from 30-50 µL.
8. After determining the concentration of the pool of sgRNAs, verify the integrity of the sgRNAs on a 1% agarose gel.
   1. Cast a 1% agarose/0.5 µg/mL ethidium bromide/TBE gel. Once the gel has solidified, place it in TBE running buffer.
   2. Mix 1 µL of the sgRNA mixture and 1 µL of RNA gel loading buffer (see Table of Materials). Load this sample in the gel and run the gel at 100 V for 5 min.
9. Visualize the bands using ultraviolet (UV) light. The pool of sgRNAs should appear as a single band. If a smear is observed, RNA degradation has likely occurred.
10. Store the pool of sgRNAs in single-use 1 µL aliquots in nuclease-free PCR strip tubes in the -80 °C freezer. For large volumes of sgRNAs mixture (30 µL or more), aliquot half of the volume into the nuclease-free PCR strip tubes and store the other half as a larger volume in a nuclease-free microcentrifuge tube. Thaw out and aliquot when needed.
11. To prevent RNA degradation, ensure that the samples in the microcentrifuge tube undergo no more than two freeze-thaw cycles.

2. Preparing reagents and materials for microinjection

NOTE: In zebrafish, the injection of Cas9 mRNA can create zygotic crispants. However, studies have shown that Cas9 protein is more efficient in creating INDELs in injected embryos^16,25. This protocol uses Cas9 protein to generate maternal crispants because this protein does not experience the same lag in activity as injected Cas9 mRNA. In theory, this should increase the probability of a biallelic mutation early in development resulting in an increased chance of a more extensive section of the germline being affected. Other protocols and resources detailing how to prepare for microinjections can be found elsewhere^24,26.

1. Purchase or generate Cas9 protein (see Table of Materials). Resuspend the Cas9 protein in nuclease-free water to make a 2 mg/mL solution and aliquot 1 µL into RNase-free polypropylene microcentrifuge tubes. Store these as single-use tubes at -80 °C.
2. The afternoon before the injection, use a micropipette puller to pull a glass capillary and create an injection needle. Store the unbroken needle in an enclosed needle holder until the morning of microinjections.
3. To create an injection plate, pour 20 mL of 1.5% agarose/sterile H₂O to fill half of a 100 mm X 15 mm Petri dish and wait for it to solidify. Once the agarose solution is set, add 20 mL of 1.5% agarose/sterile H₂O to the Petri dish and place the plastic mold (see Table of Materials) into the liquid agarose and allow it to harden.
4. After the agarose has hardened, remove the plastic mold and store the injection plate upside down in a refrigerator until the morning of injections. A single plate can be used for multiple injections as long as the wells maintain their integrity.

3. Microinjection of CRISPR-Cas9 cocktail into a one-cell zebrafish embryo to generate maternal crispants

NOTE: More resources for microinjection into zebrafish embryos can be found elsewhere^24,26,27. Injecting the CRISPR-Cas9 mixture into the developing blastodisc of one-cell embryos may increase the probability of creating maternal crispants. The mixture can also be injected into the yolk sac up to the 2-cell stage. However, mixtures injected into the yolk depend on ooplasmic streaming to reach the blastodisc, so CRISPR-Cas9 injected into the yolk could decrease the cutting efficiency of the CRISPR-Cas9²⁸.

1. The afternoon before microinjections, set up wild-type crosses in zebrafish mating boxes. Keep both the male and female fish in the same tank but separate them with a mating box divider or place the female inside an egg-laying insert.
2. On the morning of the experiment, take out one 2 mg/mL aliquot of Cas9 protein and one aliquot of the pool of sgRNAs. In the RNase-free polypropylene microcentrifuge tube that contains the Cas9 protein, assemble a 5 µL injection mixture that includes the pool of sgRNAs, 1 µL of 0.5% phenol red solution, and nuclease-free water. Aim for a final concentration of 400 ng/µL Cas9 protein and 200 ng/µL of the pooled sgRNAs in RNase-free water or a 2:1 ratio of Cas9 protein to sgRNAs in the injected embryo. This injection mixture can be stored on ice for the morning of the injection.
3. Remove an injection plate from the refrigerator and let it warm up to room temperature (RT) for at least 20 min.
4. After the injection cocktail is assembled, allow the male and female to mate, e.g., by removing the mating box divider or by placing the male in the same egg-laying insert as the female, as appropriate.
5. After the fish have laid but before the embryos have been collected, cut the tip of an unbroken needle using a new razor blade or fine forceps to create a needle that has a bore small enough to avoid embryo damage but is wide enough so that it will not become clogged with injection mixture. After the needle has been cut, load the needle with the injection mixture using a microloader pipette tip inserted into the back end of the capillary (see Tables of Materials).
6. After the needle is filled, incubate the needle for 5 min at RT to assemble Cas9-sgRNA complexes.
7. Turn the microinjector on and insert the needle into the micromanipulator. Place a drop of mineral oil onto a micrometer slide and calibrate the needle by adjusting the injection pressure until the needle ejects a 1 nL bolus into the mineral oil.
   NOTE: When injecting into the mineral oil, a 1 nL bolus will have a diameter of approximately 0.125 mm (or radius of 0.062 mm) as measured with the micrometer slide.
8. To synchronize the embryos, collect them after 10 min using a plastic strainer and rinse them into a Petri dish using 1x E3 media (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl₂, 0.33 mM MgSO₄, and add 20 µL of 0.03 M Methylene Blue per 1 L of 1x E3). Remove 10-15 embryos and place them into a separate Petri dish to be kept as uninjected controls.
9. Transfer the rest of the developing embryos into the wells of the injection plate.
10. Inject 1 nL of solution (a total of 400 pg of Cas9 protein and 200 pg of sgRNAs) into the developing blastodisc of a one-cell embryo. If the tip of the needle becomes clogged, use forceps to break the tip back and recalibrate the needle to eject a 1 nL bolus. Ensure to inject all embryos during the first 40 min of development after fertilization.
11. After the injection is completed, return the injected embryos into a labeled Petri dish that contains 1x E3 media and allow them to develop throughout the day. Remove any embryos that are unfertilized or are not developing normally according to the zebrafish staging series²⁹.

4. Screening for somatic INDELs in F0 injected embryos

NOTE: Other methods for identifying INDELs, such as T7 endonuclease I assay or high-resolution melting analysis, can be used when determining if the embryos contain somatic INDELs ³⁰.

1. The next day after injections, remove defective and lysed embryos from the Petri dish and replace the 1x E3 media to maintain embryo health.
2. After cleaning out the dish, collect six healthy injected embryos and two control embryos from the uninjected plate. Place each embryo individually into a single well of a PCR strip tube and label the top of the tubes.
3. To extract the genomic DNA of individual embryos, remove the excess E3 media from the wells of the strip tube and add 100 µL of 50 mM NaOH per well.
4. Incubate the embryos at 95 °C for 20 min. Then cool the samples down to 4 °C, add 10 µL of 1 M Tris HCL (pH 7.5), and vortex them for 5 to 10 s. This extracted DNA can be stored at -20 °C for at least 6 months without significant DNA degradation.
5. Design unique screening primers for each guide site to amplify a 100-110 bp DNA fragment that includes the CRISPR-Cas9 target site. If possible, place the target site in the middle of the amplified fragment, allowing for the identifications of larger deletions.
6. For each of the four guide target sites, set up eight 25 µL PCR reactions using 5 µL of the prepared single-embryo genomic DNA, PCR mix, and the guide-specific screening primers to identify somatic mutations in the target site (Table 1).
7. Cast a 2.5% agarose/0.5 µg/mL ethidium bromide/TBE gel using combs that create approximately 0.625 cm wide wells. This wide comb allows for better resolution when detecting size changes to the genomic sequence. After the gel has solidified, place it into an electrophoresis chamber that contains TBE running buffer.
8. Add 5 µL of 6x loading dye to the PCR product and load 25 µL of this mixture into the gel. Make sure that the injected and control samples are run on the same row of the gel. After all the samples are loaded, add 5 µL of ethidium bromide per 1 L of TBE running buffer to the positive end of the gel box.
9. Run the gel at 120 V until the DNA bands resolve or the DNA has approached the end of the lane. If the Cas9 created INDELs in the target site, a smear is typically observed in injected samples but not the controls.
10. If smears are observed in a minimum of three out of the four guide sites in embryos injected with four guide RNAs, grow up the sibling injected embryos.
11. Whenever the injected samples do not contain smears in the required number of guide sites, design new guide RNAs to replace those that did not work and create a new pool of guide RNAs that includes the ones that worked and the newly designed ones. Inject and test the new pool for somatic INDELs as described above.

5. Identification of maternal-effect phenotypes in maternal crispant embryos

NOTE: Once the injected F0 females have reached sexual maturity, their germline cells have the potential to generate a mixture of maternal crispant and wild-type embryos. Even though this mixture allows for internal controls for fertilization and developmental timing, it is still beneficial to set up a wild-type incross as an external control in case a clutch from F0 female contains only maternal crispant embryos.

1. The afternoon before the experiment, set up the F0 injected females against wild-type males and control wild-type crosses in standard zebrafish mating boxes. Place both the male and female fish in the same tank but separate them with a mating box divider or place the female inside an egg-laying insert.
2. On the morning of the experiment, allow the male and female to start mating, e.g., by removing the mating box divider or placing the male in the same egg-laying insert as the female.
3. Collect the embryos every 10 min by moving the egg-laying insert into a new mating tank bottom that contains fresh system water and label the tank with a tag identifying the individual F0 female. Take the old mating tank and pour the water through a tea strainer to collect the embryos from one individual 10-min clutch.
4. Once the embryos from a single 10-min clutch have been collected in the strainer, transfer them to a Petri dish containing 1x E3 media. Label the Petri dish with the time of collection and the fish information.
5. Under a dissecting microscope with a transmitted light source, observe the embryos undergoing development every hour for the first 6-8 h and daily for the next 5 days.
6. Identify potential maternal crispant embryos by gross morphological changes in their development compared to time-matched wild-type controls²⁹.
7. Move the potential maternal crispant embryos to a Petri dish that contains 1x E3 media and assay for morphological phenotype at 24 hpf and viability (e.g., swim bladder inflation) at 5 days post fertilization.

6. Sequencing alleles in maternal crispant haploids

NOTE: Maternal crispant haploids contain a single allele in the targeted locus, allowing for the identification of INDELs in the target gene via Sanger sequencing. Maternal crispant haploids embryos can also be analyzed using next-generation sequencing assays. Embryos that show a maternal crispant phenotype are expected to carry a lesion in at least one of the four target sites (See Discussion).

1. The afternoon before the experiment, set up mating pairs of F0 females known to produce maternal crispant embryos crossed to wild-type males. Keep the wild-type males physically separated from the females using a mating box divider or place the female in the egg-laying insert.
2. On the morning of the experiment, remove the physical partition or place both the males and females within the egg-laying insert to initiate mating. At the first sign of egg-laying, interrupt breeding by separating the male and F0 females. Keep each separated F0 female in individual mating boxes.
3. Prepare UV-treated sperm solution using testes from one wild-type male for every 100 µL of Hank's solution (Table 2), sufficient to fertilize extruded eggs from one female, as previously described³¹.
4. Manually extrude mature eggs from the pre-selected F0 females and perform in vitro fertilization (IVF) using the UV-treated sperm³¹.
5. After in vitro fertilization, allow the haploid embryos to develop until the maternal crispant phenotype is observed and place those embryos into a different Petri dish.
6. Once the maternal crispant haploid embryos have been identified, allow them to develop for at least 6 h post-fertilization.
7. To extract the genomic DNA from at least ten maternal crispant haploid embryos, place a single haploid embryo into an individual well of a PCR strip tube, remove excess E3 media from the well and add 50 µL of 50 mM NaOH.
8. Incubate the embryos at 95 °C for 20 min. Then cool the samples down to 4 °C, add 5 µL of 1 M Tris HCL (pH 7.5), and vortex for 5-10 s. The extracted DNA can be stored at -20 °C for up to 6 months.
9. To identify which guide sites contain INDELs, design sequencing primers to amplify a DNA fragment that includes all four CRISPR-Cas9 target sites. These sequencing primers allow for the identification of INDELs that span multiple guide sites.
10. Set up two 25 µL PCR reactions per embryo using 5 µL of the prepared genomic DNA and the sequencing primers.
11. After the PCR is finished, purify and concentrate the two samples using a DNA clean-up and concentrator kit (see Table of Materials). Then submit the DNA fragment to Sanger sequencing using both the forward and reverse sequencing primers.
12. After the haploid maternal crispant fragment has been sequenced, align it to the wild-type sequence and identify INDELs in the target sites using a sequence alignment program.