Generating Nonmosaic Mutants in Xenopus Using CRISPR–Cas in Oocytes
- ↵1Correspondence: cha{at}ucmo.edu
Abstract
In CRISPR–Cas9 genome editing, double-strand DNA breaks (DSBs) primarily undergo repair through nonhomologous end joining (NHEJ), which produces insertion or deletion of random nucleotides within the targeted region (indels). As a result, frameshift mutation-mediated loss-of-function mutants are frequently produced. An alternative repair mechanism, homology-directed repair (HDR), can be used to fix DSBs at relatively low frequency. By injecting a DNA-homology repair construct with the CRISPR–Cas components, specific nucleotide sequences can be introduced within the target region by HDR. We have taken advantage of the fact that Xenopus oocytes have much higher levels of HDR than eggs to increase the effectiveness of creating precise mutations. We introduced the oocyte host transfer technique, well established for knockdown of maternal mRNA for loss-of-function experiments, to CRISPR–Cas9-mediated genome editing. The host-transfer technique is based on the ability of Xenopus oocytes to be isolated, injected with CRISPR–Cas components, and cultured in vitro for up to 5 d before fertilization. During these 5 d, CRISPR–Cas components degrade, preventing further alterations to the paternal or maternal genomes after fertilization and resulting in heterozygous, nonmosaic embryos. Treatment of oocytes with a DNA ligase IV inhibitor, which blocks the NHEJ repair pathway, before fertilization further improves the efficiency of HDR. This method allows straightforward generation of either nonmosaic F0 heterozygous indel mutant Xenopus or Xenopus with efficient, targeted insertion of small DNA fragments (73–104 nt). The germline transmission of mutations in these animals allows homozygous mutants to be obtained one generation (F1) sooner than previously reported.
MATERIALS
Reagents
Agarose
BSA (0.1% in 0.1× MMR)
Collagenase VII, cell culture grade (Sigma-Aldrich C2799-15KU) (optional; see Step 13)
Dejellying solution (2% cysteine [Sigma-Aldrich] in 0.1× MMR)
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Prepare fresh; adjust the pH to 7.6–7.8 with NaOH.
DNA ligase IV inhibitor (Selleck Chemical SCR-7)
Ethanol (100%)
Ethyl 3-aminobenzoate methanesulfonate salt (tricaine, MS-222; 2%)
Forward and reverse polymerase chain reaction (PCR) primers to make single-guide RNA (sgRNA) templates (see Step 2)
Gel electrophoresis buffer
Genomic PCR primers (see Step 33)
Human chorionic gonadotropin (HCG; Chorulon)
Marc's modified Ringer's (MMR; 10×; pH 7.6)
MegaShortscript Kit (Ambion)
Nested PCR primer (optional; see Step 36)
Nuclease-free water
Phusion High-Fidelity PCR Kit (NEB)
Progesterone (e.g., Sigma-Aldrich; 1 mm in 100% ethanol)
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Store at −20°C.
QIAquick PCR purification Kit (QIAGEN)
Recombinant SpCas9 protein (PNA Bio)
Repair oligo (Ultramer, Integrated DNA Technologies)
TOPO TA Cloning Kit (Life Technologies)
Xenopus laevis or tropicalis female frogs
Xenopus laevis or tropicalis testis (obtained as in Protocol: Obtaining Xenopus laevis Embryos [Shaidani et al. 2021] or Protocol: Obtaining Xenopus tropicalis Embryos by In Vitro Fertilization [Lane and Khokha 2021])
Equipment
Catheters (BD Insyte Autoguard 381457 [for X. laevis] or 330003 [for X. tropicalis]) (optional; see Step 22)
Gel electrophoresis apparatus
Glass bead sterilizer
Low-temperature incubators
Microinjector with stereomicroscope
Microscope slides
Pasteur pipettes
PCR machine
Petri dishes (90- and 100-mm)
Plastic spoon (perforated)
Plastic tissue grinder
Platform rocker
Surgical instruments (scissors, surgical blades, needle holders, and forceps)
Sutures (4-0 silk black braided c-17, with 12-mm 3/8 circle needle; Surgical Specialties 785B)
METHOD
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This procedure is based on Tadjuidje and Cha (2018).
Preparation of CRISPR–Cas9 Component
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1. Identify potential CRISPR–Cas9 target sites(N18–20) by finding sites adjacent to protospacer-adjacent motif (PAM) sequences within the genomic sequence of the gene of interest. Identify sequences using CHOPCHOP v3 (https://chopchop.cbu.uib.no) for both X. laevis and X. tropicalis.
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Off-target screening is automatically performed in the algorithm (Labun et al. 2019; Montague et al. 2014).
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2. To make sgRNA templates, design two long PCR primers as described below.
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i. Design a unique forward primer for each target site that contains the T7 promoter sequence followed by an initiator guanine for better transcriptional activity, followed by the targeting sequence(N18–20) without the PAM sequence, and then a portion of the sgRNA backbone: 5′-GAAATTAATACGACTCACTATAGGN18–20GTTTAAGAGCTATGCTGGAAACAGCATAGCA-3′.
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ii. Use a universal reverse primer, with the sequence 5′-AAAAGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCTTATTTAAACTTGCTATGCTGTTTC-3′, for all sgRNA templates.
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A region of partial complementary to the forward primer is indicated in italics. We use the sgRNA(F+E) backbone to improve the specificity and stability (Chen et al. 2013).
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When mixed, these two primers base pair to one another. Each primer is then extended to generate a double-stranded DNA template for sgRNA transcription.
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3. Prepare a PCR reaction to generate the sgRNA templates using a high-fidelity DNA polymerase (e.g., NEB Phusion HF).
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i. Prepare a total reaction mixture of 100 µL according to the manufacturer's instructions.
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As an example, a typical reaction might contain 10 µL of 10× buffer, 1.2 µL of 25 mm dNTP mix, 2 µL of 50 mm MgSO4, 2 µL each primer at 100 pmol/µL, 1 µL of DNA polymerase, and nuclease-free H2O to 100 µL.
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ii. Carry out PCR with the following conditions: 5 min at 94°C, 20 cycles of 20 sec at 94°C, 20 sec at 58°C, and 15 sec at 68°C, followed by a final extension of 5 min at 68°C.
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iii. Analyze 1 µL of the reaction by agarose gel electrophoresis to confirm the synthesis of the desired product.
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iv. Purify sgRNA templates with a column-based PCR cleanup kit.
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4. Synthesize sgRNA using the MegaShortscript Kit (Ambion) according to the manufacturer's instructions.
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A minimum of 300 pg of the templates is required.
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5. Purify the sgRNA with the RNeasy Kit (QIAGEN).
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6. If the aim of genome editing is to introduce or change DNA sequence, design the repair oligo, a ∼200 base, single-stranded oligonucleotide, for the desired purpose.
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For example, if adding the sequence for a carboxy-terminus epitope tag to a gene, the oligonucleotide would typically include 30–50 base homology at either end spanning the targeted locus. An epitope tag sequence would be added just before the endogenous stop codon. To prevent repaired sequences from being cleaved by Cas9 after successful recombination, silent mutations are introduced into the sgRNA targeting sequence. Also, the efficiency of the HDR mediated knock-in can be increased by including phosphorothioate modifications in the final two bases on both ends and by having asymmetric homology arms in the donor oligo (Renaud et al. 2016; Richardson et al. 2016) including in Xenopus (Nakayama et al. 2020). To confirm the incorporation of the desired sequence, introduce a new restriction enzyme site in the oligonucleotide to facilitate the screening process. When introducing an epitope tag, the epitope sequence is usually large enough to create a second band on 2% agarose gel of a short amplicon across the target region.
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Oocyte Preparation for Microinjection
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Perform oocyte collection following the method described by Mir and Heasman (2008) and Olson et al. (2012) with the modifications described below.
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7. Perform frog surgery under aseptic conditions. Sterilize all surgical instruments except single-use surgical blades for 5 min at 250°C in a glass bead sterilizer.
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8. Submerge females that will act as ovary donors in 0.2% Tricaine (MS-222) for 5–8 min at room temperature. Closely monitor frogs until they become unresponsive to a gentle touch on the lower jaw (also known as the swallowing reflex).
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9. Place the frog on its back. Use a surgical blade to make a small incision in the skin on one side of the lower abdomen. Use sterile curved scissors to extend the incision to ∼1.0 cm in length. Lift the abdominal wall away from underlying organs and make a similar incision through the muscle layer. Pull part of the ovary out of the abdominal cavity. Cut out a portion of the ovary with scissors. Place the ovary fragment in OCM and evaluate under a dissecting microscope.
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10. Suture the incision, beginning with the muscle layer (three stitches) and followed by the skin (three stiches).
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11. Rinse the frog briefly in running tap water to remove any residual MS-222 from her skin and return her to paper towel soaked in fresh water to ensure she neither dries out nor drowns during recovery. Monitor her until she is fully awake and alert then return her to a tank alone to prevent the stitches from being inadvertently caught on other frogs’ claws. Return to a normal tank after 2–3 d.
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12. Cut the ovary into 1- to 2-cm3-sized pieces using surgical scissors. Place the pieces in OCM with approximately eight to 12 pieces per 90-mm dish. To keep oocytes healthy, change the medium every day.
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13. Leave the pieces of ovary in OCM overnight at 16°C to soften the surrounding follicle tissues and make the defolliculation easier. Using a pair of forceps, manually defolliculate both X. laevis and X. tropicalis oocytes in OCM under aseptic conditions. Do this by holding the transparent stalk linking the oocyte to the ovary with forceps and using the other pair to gently pull away the oocyte, without touching the oocyte itself. Alternatively, for X. tropicalis, gently shake small pieces of ovary in 0.1 KU/mL collagenase VII in OCM for ∼2 h at room temperature to release the individual, full-grown oocytes from the ovarian tissue. Sort and thoroughly rinse them in fresh OCM. Store sorted oocytes in fresh OCM at room temperature while you test for maturation.
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14. Before beginning a full experiment, test the maturation of each batch of defolliculated oocytes. Do this by treating a dozen defolliculated oocytes with 2 µm progesterone in OCM for 5 to 8 h (for X. laevis) or 30 min to 1 h (for X. tropicalis) at room temperature. Calculate the maturation rate as the % of oocytes undergoing germinal vesicle breakdown (GVBD). Judge GVBD by the appearance of a white spot at the animal pole of the oocyte. If the rate of GVBD is <80%, then discard the current batch and use ovaries from other donor females.
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We routinely use mixed oocytes from at least two different donor females, which can increase the overall success rate.
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sgRNA Injection and Oocyte Host Transfer
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15. Using standard techniques, inject oocytes with sgRNA as follows:
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For indel mutations, inject 300 pg of sgRNA and 600 pg of Cas9 protein for X. laevis in up to a 10-nL injection volume or 200 pg of sgRNA and 300 pg of Cas9 protein for X. tropicalis in up to a 4-nL injection volume.
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For knock-in, HDR experiments, include the repair oligo (200 pg for X. laevis, 30 pg for X. tropicalis) in the injection solution, staying within the maximum injectable volume.
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16. For indel mutations keep the injected oocytes in OCM for 72 h at 18°C (for X. laevis) or 23°C (for X. tropicalis). Change the culture medium daily and remove any damaged oocytes from the culture. For knock-in, HDR experiments, include 5 µM SCR-7, the DNA ligase IV inhibitor, in the OCM, including when the culture medium is changed daily.
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17. Prepare host X. laevis females and inject with 1000 units of HCG into the dorsal lymph sac. Keep injected frogs for 12–14 h at 18°C before the host transfer surgery. Prepare three female frogs for each host needed to ensure appropriately timed egg laying.
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X. laevis hosts are used for X. laevis oocytes as well as X. tropicalis oocytes.
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18. Following the 72-h incubation in Step 16, culture oocytes in 10 mL of 2 µm progesterone in OCM for 12 h at 18°C (for X. laevis) or 3 h at 23°C (for X. tropicalis). Omit SCR-7 during this incubation.
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19. On the day of surgery, stain oocytes to distinguish among experimental groups. Add vital dye stock solutions to the dish containing the oocytes (in 10 mL OCM) in the amounts listed below. Stain for 10 min at room temperature with gentle rocking.
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Red: 100 µL of Neutral Red
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Brown: 100 µL of Bismarck Brown
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Blue: 100 µL of Nile Blue A
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Mauve: 75 µL of Neutral Red + 100 µL of Nile Blue A
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Green: 100 µL of Bismarck Brown alone for 5 min + 100 µL of Nile Blue A for 10 min
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20. While staining the oocytes, submerge a host female that has just begun laying good-quality eggs in 0.2% Tricaine at room temperature. Monitor the frog until it just becomes unresponsive. Once the frog becomes unresponsive, remove it from the tricaine and lay it on a water-soaked paper towel to prevent overdosing.
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21. Transfer the stained oocytes to a 100-mm dish containing fresh OCM to prevent overstaining. If using more than two colors, place the eggs in separate piles in the dish until the last moment to avoid unnecessary contact between two different colored eggs.
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22. Make a small incision in the frog similar to that made when removing ovaries but make the body wall incision just large enough to accommodate the tip of a glass pipette. To prevent the loss of experimental oocytes, hold one side of the body wall up until the first stitch is made after oocyte implantation.
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Alternatively, lay the anesthetized frog on its back and insert the needle of a catheter into the lower abdomen at an angle almost parallel to the surgery bench. Face the catheter tip toward the frog's midline to avoid puncturing an internal organ. Pipette the oocytes through the barrel of catheter with minimum amount of OCM. Once the eggs are transferred, tilt the frog to help the transferred oocytes spread into the abdominal cavity. Slowly remove the catheter and place the frog in 2 L of high-salt solution. Sutures are not required.
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23. Load the treated oocytes into an OCM-coated Pasteur pipette. Insert the pipette tip into the opening of the body wall. Spread the oocytes in between the ovary and the body wall.
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24. Suture the body wall and skin. Allow the frog to recover from anesthesia as before. When the frog is fully awake and alert (within 30 min), transfer her into 2 L of high-salt solution.
Egg Collection and Fertilization
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25. To ensure the eggs’ competency, allow the host frog to lay eggs into the high-salt solution. Two hours after colored eggs first appear, gently squeeze the frog in high-salt solution at intervals of 30 min or more for a maximum of two more hours to collect additional colored eggs. If colored eggs are not released after 5 h postsurgery, squeeze the frog anyway.
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26. Use a pipette to place only the colored eggs into a new dish. Remove excess high-salt solution by decanting and using a pipette.
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27. Place a large piece of species-appropriate testis into a 1.5-mL tube containing 200 µL of 0.1× MMR, 0.1% BSA at room temperature. Crush it with a disposable plastic tissue grinder and immediately spread the sperm around the colored eggs with a plastic transfer pipette. Wait 5 min, and then flood the dish with 0.1× MMR. Keep the embryos in jelly until the blastula stage to improve the survival rate of host transfer embryos unless you are performing early analysis. Before dejellying, keep the embryos in 0.1× MMR at 18°C for X. laevis and 23°C for X. tropicalis.
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28. When the embryos reach the desired stage, remove the 0.1× MMR and add dejellying solution to the dish. Gently swirl the embryos for 2–3 min at room temperature until the jelly coats are removed. Rinse thoroughly with 0.1× MMR and keep embryos in 0.1× MMR at desired temperature, which is generally 18°C for X. laevis and 23°C for X. tropicalis.
Genotyping
Genomic DNA Extraction and PCR from Tadpoles
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29. Transfer each tadpole (NF stage 42–45) to a 0.0025% Tricaine solution at room temperature and wait for 30 sec. Once it has stopped moving, take it out with a perforated plastic spoon and place it on top of a microscope slide. Cut off one-fourth of the tail with a surgical blade and place the fragment in lysis buffer (150 µL per tail for X. laevis; 100 µL per tail for X. tropicalis).
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30. Return the amputated tadpole to a small volume of 0.1× MMR. From this point on, assign an identification code for each tadpole. Make sure to clean the blade and forceps between the tadpoles to minimize cross-contamination.
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31. Incubate the tail in lysis buffer for 6 h (up to overnight) at 56°C.
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32. To denature the proteinase K in the lysis buffer, incubate the lysate for 10 min at 95°C. Cool to room temperature. Dilute the lysate 1:5 dilution in nuclease-free water to prepare the template for the PCR reaction and store at −20°C.
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33. Use the Phusion High-fidelity Kit to carry out PCR according to the manufacturer's instructions. We use the Primer3 website (http://bioinfo.ut.ee/primer3) to design primers for genomic PCR and sequencing. Use 1–4 µL of diluted lysate from Step 32 as a template in each sample.
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34. Perform PCR as follows: 5 min at 96°C followed by 40 cycles of 20 sec at 96°C, 20 sec at 64°C, 20 sec at 72°C, followed by a final extension of 5 min at 72°C.
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35. Analyze a fraction of the PCR reaction on a 2% agarose gel to confirm successful amplification. If further analysis is necessary, purify the PCR product with a QIAquick purification kit.
Sequencing
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36. Submit the purified PCR product for direct Sanger sequencing using one of the PCR primers or one nested primer. Mixed peaks will occur in the sequencing results after the predicted sgRNA targeting site or for the insertion.
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37. If your mutation was an indel, once the result has come back, use TIDE or CRISP-ID on the web to reveal the frequency and the nature of mutations (Brinkman et al. 2014; Dehairs et al. 2016). TIDE analysis will reveal the number of inserted or deleted bases and the proportion of mutations found in the PCR products. CRISP-ID analysis will differentiate between unmixed sequences and mixed peaks for up to three sequences.
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38. If your mutation was a knock-in using HDR, use the TIDER algorithm to quantitate the frequency of the templated mutation and the spectrum of nontemplated indels (Brinkman et al. 2018)
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39. To get the precise sequence information within the embryo, clone the PCR products with a TA Cloning Kit according to the manufacturer's instructions. Sequence at least 20 randomly selected clones for both type and frequency.
DISCUSSION
CRISPR–Cas9 genome editing is based on sequence recognition between a guiding RNA and the target DNA, combined with the catalytic ability of the Cas9 enzyme to make double-strand DNA breaks within the target sequences (Cong et al. 2013; Mali et al. 2013; Qi et al. 2013; Wang et al. 2013; Nakayama et al. 2014). These double-strand breaks are mainly repaired by nonhomologous end joining (NHEJ), which produces insertion or deletion of random nucleotides within the targeted region (indels). As a result, frameshift mutation–mediated loss-of-function mutants are easily produced (Blitz et al. 2013; Nakayama et al. 2013; Xue et al. 2014; Kotani et al. 2015; Wang et al. 2015). An alternative repair mechanism, homology-directed repair (HDR) can be used to fix the double-stranded break at relatively low frequency. By providing a DNA-homology repair construct together with the CRISPR–Cas components, specific nucleotide sequences can be introduced within the target region by HDR (Gratz et al. 2014; Miyaoka et al. 2014; Yu et al. 2014; Aslan et al. 2017).
To increase the frequency of HDR and hence the effectiveness of creating precise mutations in Xenopus, we have taken advantage of the fact that oocytes have much higher levels of HDR than eggs (Hagmann et al. 1996). The oocyte host transfer technique is well established for knockdown of maternal mRNA for loss of function experiments (see Protocol: Oocyte Host-Transfer and Maternal mRNA Depletion Experiments in Xenopus [Houston 2018]; Mir and Heasman 2008). We have introduced this technique to CRISPR–Cas9-mediated genome editing. The method takes advantage of the fact that Xenopus oocytes can be isolated, injected with CRISPR–Cas components, and cultured in vitro for up to 5 d before they are fertilized using the host-transfer technique. During these 5 d, we have shown that the CRISPR–Cas components degrade, preventing further alterations to the paternal or maternal genomes after fertilization and resulting in heterozygous, nonmosaic embryos. The efficiency of HDR is further improved by treating the oocytes with a DNA ligase IV inhibitor (which inhibits the NHEJ repair pathway) before fertilization (Chu et al. 2015; Maruyama et al. 2015; Ratzan et al. 2017). The method described is a straightforward approach to generating either nonmosaic F0 heterozygous indel mutant Xenopus or those with efficient, targeted insertion of small DNA fragments (73–104 nt) (Aslan et al. 2017). These have germline transmission, which offers the chance to obtain homozygous mutants one generation (F1) earlier than previously reported (Nakayama et al. 2013; Yu et al. 2014; Ratzan et al. 2017).
