A Surgical Method for Oocyte Injection and CRISPR-Cas9 Mutagenesis in Anolis Lizards

Abstract

Squamates, the taxon that comprises lizards and snakes, are a diverse assemblage of reptiles represented by more than 11,000 described species. Studies of gene function in squamates, however, have remained very limited, largely due to the lack of established genetic tools and suitable experimental systems. A major challenge for the development of CRISPR-based gene editing in these reptiles is that the isolation of fertilized oocytes or single-celled embryos is impractical for most species given that fertilization occurs internally, the females of many species can store sperm, and simple methods for detecting ovulation are lacking. To overcome these challenges, we have developed a unique surgical approach in the brown anole lizard, Anolis sagrei. This procedure enables users to access and microinject unfertilized oocytes while they are still maturing within the lizard ovary. We describe methods to anesthetize adult female anoles, access the ovary through a surgical incision into the coelomic cavity, and microinject unfertilized oocytes with CRISPR-Cas9 ribonucleoprotein complexes to generate targeted mutations. This protocol enables the routine production of gene edited lizards.

INTRODUCTION

CRISPR-based gene editing has permitted studies of gene function in a wide-range of species, including vertebrates. The production of gene-edited vertebrates commonly requires access to single celled embryos, and in the case of mice and other mammals, advanced methods for retrieving and injecting fertilized eggs have long been available and enabled the rapid adoption of CRISPR-based editing (Wang et al. 2013). This process, however, is not easily achieved in certain species, including many reptiles (see the accompanying article by Sabin et al. (2025)). To enable CRISPR-based gene editing in anoles reptiles, we have developed a surgical protocol that permits the injection of Cas9-single guide RNA (sgRNA) ribonucleoprotein (RNP) complex into immature oocytes while they are still physically attached to the ovary. This allows for the continued development of oocytes after injection, ultimately leading to the production of gene edited lizards (Rasys et al., 2019).

The protocol involves identification and testing of sgRNA target sites, preparation of equipment for surgery, preoperative preparation of lizards, animal anesthetization, surgery and Cas9-sgRNA RNP microinjection, animal recovery, and identification of gene-edited embryos or hatchlings. As conveyed below, sgRNAs for the gene of interest are designed, complexed with Cas9, and tested by an in vitro cleavage assay prior to performing lizard surgeries. We then describe the steps required to access oocytes via surgery, including the anesthetization of female lizards, the application of analgesics, and the creation of an incision into the coelomic cavity. This allows oocytes to be microinjected with Cas9-sgRNA RNP after which the incision site is closed, and the incision and microinjection is repeated on the other side. Following recovery from surgery, we describe the steps for egg collection, which is done over the course of several weeks, and then outline how to genotype the offspring to detect CRISPR-induced mutations.

Users should obtain the proper permissions and follow relevant guidelines for the use of Anolis lizards before following this protocol.

MATERIALS

It is essential that you consult the appropriate Material Safety Data Sheets and your institution’s Environmental Health and Safety Office for proper handling of equipment and hazardous materials used in this protocol.

Biological Materials and Reagents

Alfaxan Multidose IDX (Alfaxalone, MWI Animal Health, Injectable, 10 mL, 10 mg/mL)

Requires a Drug Enforcement Administration (DEA) license in the United States due to being a class IV controlled substance.

Anolis sagrei lizards

Healthy, wild-caught adult females that are consistently laying fertilized eggs are needed for this protocol. Healthy males are needed to keep the females laying fertilized eggs. For Anolis husbandry, the reader is referred to published protocols (Sanger et al. 2008).

Cas9 enzyme (NEB #M0667 or Synthego SpCas9 2NLS Nuclease) and 10x Cas9 nuclease-free reaction buffer

The buffer contains molecular grade 200 mM HEPES, 1M NaCl, 50 mM MgCl₂, and 1 mM EDTA at pH 6.5.

Chlorhexidine solution (0.02% v/v)

To prepare, dilute 2% chlorhexidine gluconate (Durvet #21281, 1 gallon) in tap water and store at room temperature in the dark until use. We typically make and use within a week, but it can be used until the expiration date on the stock solution.

Dexmedetomidine hydrochloride (Dexmedesed^®, Dechra Pharmaceuticals, Injectable, 10 mL, 0.5 mg/mL)

Ethanol, laboratory grade, 70% (v/v) in deionized water

GeneJET PCR Purification Kit (Thermo Fisher Scientific K0702)

Gene-specific sgRNAs (Synthego, IDT, or similar providers)

Isopropanol, 70% (v/v) in deionized water

Lidocaine (Vet-Med International, Injectable, 50 mL, 2% solution)

Lizard DNA template (0.3μM)

Isolate as described in mouse tail clip isolation protocol (Laird et al. 1991), using lizard tail clips or embryos in place of mouse tail clips.

Meloxicam (Loxicom, Norbrook Laboratories, 10 mL, 5 mg/mL)

If meloxicam is unavailable, carprofen (Rimadyl, Zoetis, Injectable, 10 mL, 50 mg/mL) can be used instead.

Microinjection buffer

7.5mM Tris pH 7.4 and 0.2mM EDTA in nuclease-free water

Nuclease-free water (Thermo Fisher Scientific AM9937, 0.2-μm filtered)

Oligonucleotides for amplification of the test sgRNA target regions (see Step 4)

Phenol red solution

1% (w/v) phenol red solution in 7.5mM Tris pH 7.4 and 0.2mM EDTA.

Plasma-lyte A pH7.4 (VWR 68000–434)

Povidone Iodine Scrub Solution (Dynarex Povidone Iodine Scrub Solution 16 oz #1425)

Povidone Iodine Solution First Aid Antiseptic (Dynarex Povidone Iodine Solution 16 oz #1415)

Proteinase K (20 mg/mL, Thermo Scientific EO0491)

Reagents for genomic DNA extraction, PCR, gel electrophoresis and sequencing (Laird et al. 1991; Green and Sambrook 2018; Green and Sambrook 2019; Corkins et al. 2022)

RNaseZap^™ RNase Decontamination Solution (Invitrogen AM9780)

TE buffer

10 mM Tris-Cl and 1.0 mM EDTA, pH 7.5. Use molecular biology grade reagents.

Topical antibiotic

Over-the-counter triple antibiotic ointment (e.g., bacitracin, neomycin, polymyxin B)

Tris Acetate EDTA (TAE) buffer, 50X, molecular biology grade

2M Tris, 1M acetic acid, and 50 mM EDTA in double distilled water with pH 8.5

Vetbond Tissue Adhesive, 3 mL (3M, 1469SB)

Equipment and Supplies

Absorbent pads

Angled Forceps (FST 45 angled forceps 00649-11)

Box for weighing lizards

To accurately calculate anesthesia dose, each lizard must be weighed. A plastic container, small cage, or empty pipette tip box to put the lizard in, is suggested. For easy transfer, we cut a small “lizard-sized” hole into one end of an empty pipette tip box. We then pre-weigh the box and hold a lizard gently by its body with its snout facing the small hole in the pipette tip box. The natural tendency of the lizard will be to escape from your hold into the small hole. Once the lizard is secured in the box, cover the hole with a flap of lab tape. Weigh the box again to calculate the lizard’s weight.

Butterfly Forceps (Fischer Scientific DR Instruments Featherweight Entomology Forceps S72110)

Conical tubes, 50 mL

Cotton-tipped applicators (6 inch, e.g. Puritan 806-PC)

Equipment for genomic DNA extraction, PCR, and gel electrophoresis (Laird et al. 1991; Green and Sambrook 2018; Green and Sambrook 2019).

Food coloring

This is used for applying a unique color combination to each lizard upon the completion of surgery, for easy identification.

Headrest for the lizard during surgery

This can be made from floral wire (we suggest 0.5–1.0 mm in width) or a similar product.

Heat block

Iris Scissors (Roboz Surgical Instrument Co. Inc. RS 5601, McPherson-Vannas Micro Dissecting Spring Scissors: Curved, Sharp Points; 5 mm Cutting Edge; 0.15 mm Tip Width; 3” Overall Length)

Lab tape

Lights for better visibility during surgery

A ring light can be used, but we prefer photography lighting (e.g., EMART 60 LED Light)

Metal instrument tray (e.g., VWR 10193-162)

Microcentrifuge tubes (1.5 mL)

Microinjection needles, glass

These can either be purchased (e.g. World Precision Instruments TIP05TW1F) or can be pulled from glass capillary tubes (e.g. World Precision Instruments TW100F-4) using a micropipette puller (e.g., Sutter Instrument P-97)

Microinjector

Standard pressure injection system with needle holder and foot pedal. Any microinjector that works for zebrafish or Xenopus eggs should work for lizard oocytes. We have successfully used both older (i.e., Harvard Apparatus Injection System, Model PLI-100) and newer (ASI’s MPPI-3) microinjectors. A nitrogen gas tank or an air compressor (e.g. Tritech Research MINJ-CMPR1) can be used as an air source. A needle holder is required, but a micromanipulator is not needed as follicles are injected by hand.

Microscope with scale bar in eye piece

Nanodrop (e.g. Thermo Fisher ND-2000)

Needles for Hamilton syringe: (VWR, 7803–05. 33-gauge, Small Hub RN NDL. Custom specs: Length 10 mm; Point Style 4; Angle 12)

PCR tubes (0.2 mL)

Scale

To weigh lizards (female lizards’ body weight averages ~3 grams).

Small cage (e.g., Lee’s Kriter Keeper)

Used for solo housing lizards.

Surgical Record sheet

For each lizard, anesthesia and surgery data should be recorded: weight, anesthesia dose, anesthesia start time, respiration before the start of surgery, surgery start time, size of follicles found, presence or absence of fertilized eggs for the right and left oviduct, respiration after completing one side and before starting the other side if additional anesthesia was needed, surgery end time, anesthesia end time, and a unique dye identification. At the end of the surgery, we apply food coloring to the dorsal side of each lizard for identification, since they are group housed. Details are described in Step 41.

Stereomicroscope

The scope should provide 5x magnification, a large field of view, ample working distance, and a boom arm. The boom is required so that the warming platform where surgery will be performed can be placed directly below the nosepiece.

Straight Forceps (FST straight forceps, 00632–11)

Syringe, 10μL (Hamilton Syringe Model 801 Removable Needle)Warming platform

A slide warmer works well.

Weck-Cel spears (optional, see Step 31 or Troubleshooting for Step 21–23 or 23ii)

METHOD

Identification and Testing of sgRNA Target Sites

• 1Retrieve DNA sequence for the gene of interest from the AnoSag2.1 A. sagrei reference genome and identify potential Cas9 sgRNA target sites (Geneva et al. 2022).

The CRISPOR gRNA design tool contains the A. sagrei reference genome in its database (http://crispor.gi.ucsc.edu/) and can be used to identify unique target sites within the genome (Concordet and Haeussler 2018).

• 2Determine the location of known polymorphisms in your sequence of interest using the polymorphisms annotated by Geneva and colleagues (Geneva et al. 2022). Alternatively, extract DNA following a protocol such as that in Laird et al. 1991, and then PCR amplify and Sanger sequence your region of interest from several individuals to identify target sites that lack polymorphisms. Purchase 2–3 promising sgRNA sequences from IDT or Synthego. Once received, resuspend the sgRNAs in TE buffer and store at −20°C until use.

Because of the high level of genetic polymorphism in A. sagrei, care needs to be taken in the selection of target sites.

• 3Prepare 10μM Cas9 RNP by mixing the components described below in a 1.5-mL tube on ice, using all designed sgRNA sequences (two in the example shown below; see note). After the components are mixed, incubate at room temperature for 10 minutes and then store at 4°C until use.

Reagent	Reaction volume
20μM Cas9	15μL
30μM sgRNA A	5μL
30μM sgRNA B	5μL
1% phenol red with 0.01M KCl	1μL
Microinjection buffer	4μL
Total	30μL

If a different number of sgRNAs are being used than in the listed recipe, adjust the reaction volume for each sgRNA accordingly to keep the working molar concentration of Cas9 and sgRNAs equal in the final solution. Approximately 1μL of RNP solution is needed per lizard, so scale as needed.

• 4Create a DNA template for the in vitro testing of the RNPs by amplifying a 300–500 base pair region centered around the sgRNA target site(s). Perform PCR using standard practices (Green and Sambrook 2018) and check size by agarose gel electrophoresis (Green and Sambrook 2019). Using a PCR purification kit, purify the PCR product and measure the concentration using a Nanodrop spectrophotometer. Calculate the molarity of the amplified and purified DNA template using the formula (DNA concentration in ng/μL/660 × amplicon size in bp) ×10⁶ = DNA nM, where 660 is the molecular weight in g/mol for 1 base pair. Dilute the PCR product using TE buffer to a final DNA concentration of 0.3μM.
• 5Dilute 1 μL of the 10 μM RNP from Step 3 with 2.3 μL of microinjection buffer, to make 3 μM RNP.
• 6Set up the following test reaction, and incubate it at 37°C for one hour.

Reagent	Reaction volume
3μM RNP (from Step 5)	1μL
0.3μM DNA (from Step 4)	1μL
10x Cas9 nuclease free buffer	1μL
Nuclease-free water	7μL
Total	10μL

• 7Add 1 μL of 20 mg/mL Proteinase K and incubate at 56°C for an additional 10 minutes.
• 8Add loading dye to the 11-μL product from Step 7 and run this result of the test in vitro reaction on an 1.5% agarose gel with 1× TAE as a running buffer to visualize the digested product. As a control, run 1 μL of the 0.3 μM DNA template (from Step 4) mixed with 9 μL of nuclease-free water and loading dye.

The control lane will show the 300–500 bp product created in Step 4. A successful Cas9 RNP test will show at least a 70% digestion of the DNA template in the digested product lane when compared to the control lane. Often, the uncut band will almost completely disappear from the digested product lane, with smaller DNA fragments appearing.

We recommend the in vitro RNP test to be repeated before performing surgery if it has been more than two weeks since the last in vitro RNP test.

Preparation of lizards, materials, and equipment for surgery

Steps 9–12 should be performed prior to the intended day of surgery

• 9For one day of surgery, select 8–10 healthy females with a consistent history of laying eggs (ideally one egg per female per week). Healthy females are plump-looking and are bright, responsive to stimuli, and active.

The selection of reproductively active females for surgery can be achieved by routinely monitoring the lizards in your colony and recording their egg laying. If females are not laying one egg a week, see Troubleshooting.

The number of lizards needed for a project depends on several variables, but for a new gene, we typically perform 20 surgeries. This typically produces 3–5 mutants, allowing for an initial observation of genotypes and phenotypes.

An ideal weight for mature females is 2.0 g to 2.8 g. Females under 1.8 g are rarely reproductively active.

• 10If pulling glass microinjection needles instead of purchasing pre-pulled, use a micropipette puller and a program/preset for zebrafish embryo microinjection. We use a Sutter Instrument P-97 and follow the Sutter Instrument Pipette Cookbook (2025) directions for zebrafish embryo microinjection. After pulling needles or purchasing pre-pulled, break needles to the desired size. Under a stereomicroscope use a sterile pair of forceps to gently pinch the needle so it breaks and leaves an opening of 10–20 μm.

The precise diameter of the opening is not critical, as the oocytes are large and injection time can be adjusted to produce the desired volume. What is important, however, is that individual needle bores are of a similar size, to ensure consistent performance. The diameter of the needle can be measured using a microscope with a scale in one eyepiece.

The preparation of glass pipettes should be done under RNAse-free conditions. To achieve this, treat the surface of the stereomicroscope and forceps with RNAZap before breaking needles.

• 11Withhold food from selected females the day before surgery, move them into a clean cage, and remove any male.
• 12Sterilize surgical tools by leaving them in a metal instrument tray containing 0.02% (v/v) Chlorhexidine Gluconate overnight. The morning of surgery, remove tools, rinse with sterile water, air dry, and then spray with 70% isopropanol.
• 13On the day of surgery, create a solution of Dexmedetomidine (Dex) and Alfaxalone (A) with 12μL of Dex and 188 μL of A. This can be scaled as needed. Lizards will be given Dilute meloxicam (M) by adding 12μL of M to 188 μL of sterile water. Dilute 2% Lidocaine (L) by adding 20 μL of L to 180 μL of sterile water..

In the United States, alfaxalone is a categorized as a class IV controlled substance and requires a DEA license to purchase.

Carprofen (Rimadyl, 50 mg/mL) can be used for pain management in place of meloxicam. Aliquots of 4 μg/μL of Rimadyl are prepared by diluting stock with sterile water and are stored at 4°C.

Both diluted meloxicam and diluted lidocaine can be kept for 3 months before being replaced.

• 14Turn on the microinjector and air compressor. Use water mixed with a drop of food coloring in a prepared glass microinjection needle to adjust the microinjector until beads with a 100 nL volume are propelled from the end of the glass pipette with each press of the foot pedal. Since the injection time may be changed throughout the surgery depending on the follicles, note the pulse duration setting on your microinjector used for this bead size, smaller follicles will be injected with this volume.

If having trouble optimizing volume and force or if the needle’s contents are ejected upon contact with any surface, see Troubleshooting.

• 15Load approximately 2 μL of tested RNP solution from step 3 into a glass pipette using capillary action.
• 16Connect the Hamilton syringe and needle, then fill a 1.5-mL tube with 70% v/v laboratory-grade ethanol. Sterilize the Hamilton syringe by pulling up and then ejecting ethanol from the 1.5-mL tube 5 times.

When preparing the Hamilton syringe for use, spray both the plunger and the syringe barrel with 70% v/v ethanol to allow the parts to slide together smoothly. Forcing the syringe and plunger together will cause the plunger to bend, which can ruin the syringe.

• 17Take two 50-mL conical tubes and fill one with ~20 mL of Povidone Iodine Surgical Scrub Solution and the other with ~20 mL of 70% v/v ethanol. Place cotton-tipped applicators in each tube; 3 applicators will be needed for each animal undergoing surgery. Open a sterile aliquot of 10 mL of Plasma-Lyte A pH 7.4. Ensure the warming platform is covered with an absorbent pad, is set to 32°C, and the lights are on and focused on the surgery station.
• 18Lay out triple antibiotic ointment, Vet-Bond glue, food coloring, sterile pipette tips (10μL and 1000μL) and the corresponding pipettes, and a lizard headrest.

Animal Anaesthetization

• 19Weigh and tare a small cage or plastic box, then place a female lizard inside to measure its weight. After recording the weight on the surgical record sheet, calculate the anesthesia dose needed (2.8μL of the A/Dex mixture per gram of the lizard) and record this value.

Do not exceed 9.6μL of A/Dex, even if the lizard weighs more than 3.4 g.

• 20Load the sterilized 10-μL Hamilton syringe from Step 16 with the relevant A/Dex mixture calculated in Step 19
• 21After removing the lizard from the box, restrain the lizard by positioning the ventral side down and holding the lizard between your index finger and thumb. Position the needle parallel to the lizard’s spine, with the needle pointed towards the head, and inject, bevel up, between the shoulder blades (Figure 1). Inject at a rate of ~2 μL per second. Record the anesthesia start time. After every injection, sterilize the Hamilton syringe by rinsing 5 times with 70% v/v ethanol before loading again.

Figure 1: Anesthesia and analgesic injection location.

Anolis sagrei females should be injected subcutaneously with alfaxalone and meloxicam using a 10-μL Hamilton syringe. The injection location (indicated by either of the two black arrowheads) is slightly lateral to the spine and anterior to the forelimb. The lizard should be held with the non-dominant hand using the index finger and thumb, along the head and neck. This is to provide stability, as many lizards move during anesthesia injection.

Preoperative Procedures

It is important to monitor respiration throughout the surgery. Respiration is most easily observed in the upper part of the lizard’s thoracic region, slightly posterior to the forelimbs. Heart rate can be seen as fluttering along the neck and occasionally posterior to the forelimbs. The fluttering produced by heart pumping has a faster rate and produces a less obvious motion than the thoracic motion associated with respiration (Figure 2). If the lizard stops breathing, see Troubleshooting.

Figure 2: The location and relative intensity of the respiration and heart rate in an unconscious Anolis sagrei female.

Respiration can be readily monitored by the expansion and contraction of the ribcage posterior to the forelimbs (blue). Heart rate is best viewed just anterior to the forelimbs (orange). While heart rate is faster than respiratory rate, it is more subtle and can only be observed under the microscope.

• 22Place the lizard on the warming platform, and using a cotton-tipped applicator, scrub both sides of the lizard along the trunk with surgical scrub iodine, followed by scrubbing with 70% v/v ethanol. Move anteriorly up to the forelimbs and then posteriorly to the hindlimbs, ensuring the entire lateral surface is covered. Repeat for a total of three times, using a fresh cotton-tipped applicator for each iodine and ethanol scrub.
• 23Using a pipette, apply 15 μL of 0.2% lidocaine topically to both the left and right lateral regions of the lizard along the trunk, between the forelimbs and hindlimbs.
• 24Check the lizard’s respiration. Only proceed to Step 25 if respiration is at least 17 breaths per minute. If respiration is too low, wait for it to rise to 17 breaths per minute. If 10 minutes has gone by and respiration is too low, do not perform surgery on that lizard and allow it to wake up naturally, returning it to the main lizard colony.
• 25Using a sterile 10-μL Hamilton syringe, slowly inject 1 μL/g of the lizard’s body weight of the diluted meloxicam solution near the A/Dex injection location.

Forceps can be used to lift the skin and provide tension to make the injection easier. If respiration has decreased to 16 breaths a minute or lower, do not inject meloxicam. Wait until respiration has improved to at least 17 breaths a minute. See Troubleshooting if lizard stops breathing

• 26Use butterfly forceps to tightly clamp near the base of the tail (Figure 3). A lack of response indicates that the lizard is under a surgical plane of anesthesia.

Figure 3: The position of the tail clamp.

A tail clamp is performed to test if the lizard is at a surgical plane of anesthesia. The illustration indicates the location of the tail clamp with respect to the hind limbs.

Entering a surgical plane of anesthesia averages 3–5 minutes from when A/Dex is given (Step 21), although there is variability from lizard to lizard. See Troubleshooting if the lizard is not becoming unresponsive.

• 27Record respiration by counting breaths for 15 seconds. This provides a baseline for each lizard’s respiration rate under deep anesthesia. The rate of respiration should be at least 16 breaths per minute. If it is lower, wait to begin surgery.
• 28Lay the lizard on its side on the warming platform, place its head on the headrest, and gently tape its tail down with lab tape to maintain the preferred body position, with the upcoming incision location under the stereomicroscope (Figure 4A).

Figure 4: Surgical preparation and location of incisions.

The steps for preparing the flank for surgery and generating incisions to access ovarian follicles are shown. (A) The lizard shown is positioned for surgery after being scrubbed by alternating iodine and 70% v/v ethanol; the discoloration is due to the iodine. The animal should be turned slightly onto its ventral side, so the accessible region is lateral and slightly dorsal. (B) A 5-mm incision through the skin is sufficient for many surgeries and so should be used when possible, as smaller incisions heal more quickly. It should be slightly posterior to midway, between the forelimbs and hindlimbs. (C) Larger incisions will be tolerated, and so, when learning the procedure, an 8-mm incision is recommended. (D) Pushing the skin anteriorly and posteriorly with forceps exposes the ribs in the translucent layer beneath the skin with the dark pigmented layer visible below the ribs. (E) Cutting parallel and between the two ribs shows a subtle change; the darkly pigmented layer is now more visible. (F) Cutting the pigmented layer at the same angle as the rib layer exposes the organs beneath. (G) Using forceps to push the ribs dorsal and ventral shows a large yolky follicle directly underneath the incision location.

Surgery and Cas9 RNP Microinjection

• 29Record the surgery start time on the surgical record sheet. Then, proceed to the initial incision. There are three layers: skin, rib musculature layer, and pigmented layer. Incisions need to be made through these layers to access the lizard’s body cavity. The first incision is through the skin layer. Use iris scissors to cut through the skin slightly posterior to the midpoint between the hindlimbs and forelimbs, in the dorsal to ventral direction on the flank region of the lizard. The cut should not extend past the transition to belly scales on the ventral side or come within a couple of millimeters of the spine on the dorsal side (Figure 4B–C). To avoid accidentally damaging the spinal cord, pay attention to the transparency of the rib musculature layer. There is a noticeable decrease in the transparency of it as it gets close to the spine; this region should not be cut. If there is a bulge in the abdomen, suggesting the lizard is carrying a fertilized egg, shift the incision site approximately 5 mm anteriorly to avoid the egg, as it obstructs visibility and access to the follicles. (The presence of a fertilized egg can be felt while scrubbing or seen by observing the flank of the lizard. This distinct bulge is firmer, slightly smaller, and oval when compared to a large yolky follicle).

We suggest to first do an initial smaller cut of 3 mm through all three layers (skin, rib musculature layer, pigmented layer, see Steps 30–31). Then, once the location of follicles (dorsal or ventral relative to the 3 mm cut), has been identified, the cut can be lengthened in the appropriate direction as needed. A cut of no more than 5 mm (Figure 4B) will provide enough working space for an experienced surgeon to visualize key areas and locate follicles. For newer surgeons, an incision of 8 mm (Figure 4C) is suggested, as this larger incision is still tolerated by the lizard and helps the surgeon’s visualizing and maneuvering.

See Troubleshooting if the initial skin cut is too anterior or posterior.

• 30Make a smaller cut through the rib musculature layer, a translucent layer thicker than the skin, between a pair of ribs (Figure 4E). Make this cut parallel to the ribs and oblique to the initial incision through the skin.

Do not pinch the ribs with the forceps at any point during the surgery as they will break. See Troubleshooting if a rib is pinched and broken.

• 31Make a smaller cut through the pigmented layer (Figure 4F) at the same angle as the cut through the rib musculature directly above. When all the three layers (skin, rib musculature layer, and pigmented layer) are successfully cut, the internal organs in the body cavity are visible. See Troubleshooting if blood begins to hemorrhage.

If the lizard is dry (usually observed by layers of tissue or internal organs sticking to each other during surgery), add a drop or two of Plasma Lyte using a P1000 pipette. This can be repeated as needed throughout the surgical procedure. If too much Plasma Lyte was applied, the excess can be removed with Weck-Cels or a comparative absorbent material.

When cutting layers of tissue during surgery, it is important to avoid nicking the tissue beneath. This can be avoided during the initial incision by lifting the layer away from the tissue beneath using a pair of forceps, making a small cut in the desired location using the iris scissors, and then releasing the tissue. When lengthening the incision, be careful to again lift the tissue slightly before cutting, with the iris scissors angled slightly away from the tissue beneath.

• 32
  Find and position the ovarian follicles for injection.

  • i
    The typical location for the follicles is posterior and dorsal to the incision location. Look for large white or yellow balloon-like structures, as most other organs are pink or darkly pigmented (Figure 4G).

See Troubleshooting if having trouble finding the follicles.

• iiOnce follicles are found, maneuver them with blunt forceps so the cluster of small oocytes and the larger yolky oocyte are accessible for microinjection. Do not grab follicles larger than a couple millimeters, as they will burst. Move them as needed by pushing with blunt forceps or grasping the smallest follicles to manipulate all follicles.

See Troubleshooting if you are having trouble maneuvering follicles without damaging them, or if you only see one large yolky follicle.

• 33Using a glass needle, inject up to 3 oocytes with the Cas9 RNP solution from Step X. Include large yolky oocytes if present. Inject only oocytes with a diameter over 1 millimeter (see note below); note that this may result in only 1 or 2 injected oocytes per side. The successful injection of RNP solution into follicles can be tracked, as the phenol red added to the RNP solution makes injections visible within follicles. Record the presence of any oocytes over 1 mm and make note of which oocytes were injected in the surgical record sheet (i.e. the left side has injected follicles of 8mm, 4mm, and 2mm). Adjust the injection volume per the size of the oocyte (see note below). Do not inject fertilized eggs (see Step 34). See Troubleshooting if the needle clogs.

Oocytes under 1 mm should not be injected, as we have found that injection of small oocytes rarely produces mutants. Follicle diameter is estimated by comparing the follicle size to the diameter of the glass pipette, which is 1 mm (Figure 5)

To adjust the injection volume for different size oocytes, the number of injection pulses and the amount of RNP complex per injection pulse should be modified. Small oocytes (1–3mm) should be injected the same number of times as their diameter in mm (e.g., a 3-mm follicle is injected with 3 pulses of RNP). However, as the diameter of the oocyte increases, the volume increases exponentially. Therefore, the amount of RNP cannot increase linearly without dramatically decreasing the ratio of RNP volume to oocyte volume. To measure these larger volumes needed for larger follicles, the glass pipette holding the RNP can be used. By observing the decreasing volume in the glass pipette as RNP is injected into a follicle, we can track injection volumes in larger follicles. If injecting a moderately large follicle (5–7mm), the liquid level in the glass pipette should decrease by 0.5mm - 1.0mm along the length of the pipette. If injecting a very large follicle (10mm+), the liquid level in the glass pipette should decrease by 1mm to 2mm. In these circumstances, the number of injection pulses are not counted, but rather are continued until the desired decrease in volume in the glass pipette is reached. To more quickly reach these higher injection volumes, the injection time can be increased, allowing the amount of RNP per injection pulse to be doubled or tripled. Spending less time with the glass pipette in the follicles decreases the chances of damaging them. Injection volume can be decreased again once injection of a smaller follicle is desired.

Figure 5: Glass microinjection needles are used to estimate the size of follicles in the lizard.

To inject the ideal amount of RNPs, the size of the ovarian follicles needs to be estimated. This can be done by comparing the follicles to the glass microinjection needles, which are 1 mm in diameter. The ovarian follicles are depicted as circles. The numbers in the circle indicate the estimated diameter of the ovarian follicle in mm. Comparisons with the microinjection needle diameter is depicted underneath each ovarian follicle. The slight yellowish tinge in the follicles with 5 mm+ size indicates the presence of yolk.

• 34Record if a fertilized egg is observed. Eggs are generally located ventrally and posteriorly of follicles. They are more oval in shape and bright white compared to large yolky oocytes of similar sizes. Do not inject fertilized eggs, as injected eggs are quickly laid by lizards and do not develop properly. However, it is necessary to know if they are present, to accurately assess how many eggs laid from post-surgery lizards need to be screened (described in Step 49).
• 35Close the surgical incision by first aligning each cut layer so that both sides touch, with no gaps but do not overlap. Do this first with the pigmented layer, then the rib musculature layer, and finally, the skin. Once all layers are properly aligned, apply VetBond on and around the skin incision location until there is VetBond on the exterior of the skin 3 mm from the incision in every direction (see note). If it is difficult to keep the layers aligned, see Troubleshooting.

Apply VetBond by filling a 10-μL pipette tip by capillary action and then tapping on the wide end until a small droplet of the desired size comes out; using this method provides better control.

• 36Measure the respiration rate while waiting for the VetBond to dry (this typically takes ~1 minute). This information will be used in Step 38. If the VetBond dries almost immediately, see Troubleshooting.
• 37Once the glue is dry, apply triple antibiotic ointment around the glue but not on the glue, as petroleum-based products, like triple antibiotic ointment, can break down the glue.
• 38
  Prepare for the surgical incision on the other side.

  • i
    Perform a tail clamp to test if the lizard remains at a surgical plane of anesthesia.

We have found that lizards remain at a surgical plane of anesthesia for about 30 minutes. Performing surgery on one side takes an experienced surgeon about 5 minutes, meaning that most lizards do not need a second dose of anesthesia.

• iiConsider the respiration rate measured in Step 36. If the respiration rate has increased significantly compared to the initial respiration at the start of the first side (Step 27) and/or the lizard is responsive to a tail clamp, give a second dose of A/Dex. Further, if 20 minutes or more have passed since initial anesthesia (Step 25), then give a full anesthesia dose again. If it has been less than 20 minutes, then give the animal a percentage of the original dose.. See Troubleshooting for details about how much anesthesia can be given. Record the respiration rate and note whether a second dose of A/Dex was needed, as well as its volume, on the surgical record sheet.

• 39Rotate the lizard to allow access to the other side.
• 40Repeat Steps 29–37 on the other side.

Animal Recovery

• 41Lay the lizard on its ventral side on the warming platform. Then, dye the lizard’s head and/or hips with a unique color combination using food coloring for easy identification, as they will often be re-caged with other lizards for a few days after surgery (Step 45). The dye can be used as a marker for about 5 days, although dye remnants will remain for several weeks. Record the combination of dye on the surgical record sheet.

Because the lizard’s skin is naturally hydrophobic, rubbing the dye for a few seconds with a gloved finger can be effective. An alternative option is to apply 70% v/v ethanol to the skin before applying dye.

• 42Once the dye has dried, place the lizard on its dorsal side so its righting response (the ability of an animal to reorient itself in an upright position by moving its body) can be observed. Cover the lizard with the lid of a small cage.

As most lizards will instinctively right themselves when conscious, laying them on their dorsal side and observing the righting response signals them regaining consciousness and muscle control. In our experience, Anolis sagrei will transition from unconscious due to anesthesia to awake and running within a few seconds, so containment is required. The lid provides enough space for an animal to flip from its back to its stomach and move while still being in a confined space. Often, the lizards will climb onto the lid, making transfer into a cage easier.

• 43Once awake, transfer the lizard to a clean cage with other post-surgery females. Feed the lizards 24 hours after surgery and then resume a normal feeding schedule.
• 44Examine all post-surgery lizards twice a day and look for reopening of surgical sites and behavioral abnormalities.

Sometimes, the pressure of laying an egg, poor closure, or skin shedding can cause a surgical site to reopen. This can range from a small gap in the skin appearing where the surgical cut was made, to internal organs or follicles protruding from the side of the lizard. If a very small (1–2 mm) gap has appeared while the rest of the closure is healing well, consider adding a drop of VetBond. This can be less stressful for the lizard than going under anesthesia and closing the surgical site, as this can be done without handling or anesthesia. See Troubleshooting if a surgical site has opened and needs to be closed.

If a lizard is lethargic and/or will not lift its head or torso off the ground, examine carefully and consider euthanizing using approved methods (Conroy et al. 2009); in our experience, these lizards are unlikely to recover.

Screening Time

Embryos or hatchlings from each post-surgery lizard need to be screened, as they are potentially mutant. However, when and for how long those mutant eggs could be laid varies for each lizard, based on the number of follicles injected and presence or absence of fertilized eggs in the reproductive tract during the procedure. As eggs are typically laid once a week, the screening time, i.e. the number of weeks post-surgery that the injected lizard should be monitored for, as it could potentially lay an egg containing a mutant, can be calculated. Often, lizards that are in the same initial cage post-surgery will require different amounts of screening time. Because lizards in this protocol are group housed, it is almost always impossible to tell which egg comes from which lizard in a cage, and so, determining screening time and then re-caging post-surgery females allows us to group lizards by screening time, significantly decreasing the amount of unnecessary screening.

• 45
  If lizards are alert and feeding well with no incisions opening, re-cage the females 3–4 days after surgery, grouping them by screening time. To determine the length of screening time for each lizard, it is important to estimate when all injected follicles will be laid. This is done by creating a “follicle train”, a term that refers to the order that follicles will be laid in each lizard (Rasys et al., 2019). Several assumptions are made when creating the follicle train: One, that lizards will alternate laying eggs between their two ovaries, and two, that larger follicles (and eggs) are developmentally more advanced and will be laid sooner than smaller follicles. As an example, see the illustration in Figure 6.

  • i
    For each lizard, use the surgical record sheet containing notes from Step 33 and Step 34 to create the follicle train and find the screening time. First, compare the largest follicle or egg found on the left and right side. In this example, the left side has an egg, while the right side has a 10+ yolky. The approach assumes that the egg will be laid first, making the left ovary the leading ovary and the right ovary the lagging ovary. Once this is determined, the timing of follicle ovulation is alternated between the ovaries, with the largest remaining follicle being the next laid. So, in the ovaries depicted in Figure 6, one would predict the following laying order: egg (L), 10+(R), 6 (L), 4(R), 3 (L), 2.5 (R), 2.5 (L), 1.5 (R), 0.5 (L).
  • ii
    Find the latest follicle in the follicle train that was injected based on that lizard’s surgical record sheet. For instance, if the latest follicle was the 2.5-mm follicle from the left ovary, it will take 7 weeks for the example lizard to lay the last injected follicle. Assuming that one egg is laid a week, one could potentially obtain mutants from eggs laid until 7 weeks post-surgery for that lizard.

Figure 6: The follicle train.

The illustration depicts the eggs and ovarian follicles found in the left and right ovary of a reproductively active and healthy lizard. The oval shape indicates an egg and the circles indicate ovarian follicles. The numbers associated with the follicles are the size of the respective follicle in mm. The yellow color of the circles shows the presence of yolk in the follicle. In this example, the left ovary has one egg and 4 follicles, while the right has 4 follicles and no egg.

If for any reason no follicles were injected in one of the ovaries (i.e., because follicles were not seen or could not be injected), those weeks are still included in the follicle train.

• iiiAdd an extra 2–3 weeks on top of screening time calculated, in case the lizard slowed egg production after surgery.

• 46Add the breeding male back to the cage 5–8 days after surgery. The male is needed to continue to provide sperm to fertilize the oocytes of the post-surgery females. The decision of when to reintroduce the breeding male into the cage depends upon the health and behavior of all the post-surgery lizards in that cage. Daily health monitoring of the lizards will dictate this time window.
• 47Monitor for two additional days after the male is introduced, as the male can sometimes stress the females. Mating activity might lead to a reopening of surgical incisions in females who are in the early stages of recovery.
• 48Two days after the male has been added, the cage no longer needs additional monitoring and can be treated like the rest of the colony.

Identification of Gene-edited Embryos or Hatchlings

• 49Collect eggs and incubate until hatch or dissect at a desired timepoint of embryonic development. Follow standard protocols for the collection and incubation of Anolis sagrei eggs (Sanger et al. 2008). To determine which embryos or hatchlings are mutants, isolate genomic DNA, PCR amplify the region of interest, and sequence using standard practices (Laird et al. 1991; Green and Sambrook 2018; Corkins et al. 2022).

Most of our analysis is carried out on F0 lizards, without additional breeding. If large numbers of mutants are needed, F0 heterozygous lizards can be raised to adulthood and crossed to produce a stable line following standard Anolis husbandry, which can take close to a year (Sanger et al. 2008). Crossing F1 lizards will yield F2 lizards homozygous for the mutation. However, it is often easier to perform more surgeries to increase mutant numbers rather than creating and maintaining stable lines.

TROUBLESHOOTING

Problem (Step 1): Females are not producing enough eggs for surgery.

Solution: This can occur for several reasons. For females to lay eggs at a high frequency (1 egg/week), they need plentiful access to calcium-dusted food for several weeks; increasing levels of calcium supplementation can also be effective. If adult females have been recently caught (under 2 months), wait to see if egg laying increases. If the lizards are over 1.5 years old, their rate of egg lay will often be lower compared to their younger counterparts. Additionally, lizards typically lay best in the late spring and summer months. Perform surgeries during this time and ensure the light/dark cycles in their colony housing is similar to spring and summer time.

Problem (Step 6): The beads of solution coming from the injection needle are not the correct size or are running up the side of the needle.

Solution: Adjust the injection time to create beads of the desired size. Injection pressure should be as low as possible while the beads of liquid still eject from the glass needle. If they run up the side, the injection pressure needs to be increased.

Problem (Step 6): Liquid ejects from the injection needle upon contact with any surface.

Solution: This is caused by the holding pressure being too high. Decrease the holding pressure in small increments until the problem is resolved. Having the holding pressure set too high will cause liquid to eject upon any contact with a follicle before it can be punctured, and setting the holding pressure too low will lead to an excess of yolk wicking into the needle. Therefore, maintain as high of a holding pressure as possible without fluid ejecting from the needle unprompted by a pedal push.

Problem (Steps 22–33): The lizard crashed and stopped breathing.

Solution: This can happen for a variety of reasons, including injecting the anesthesia too quickly or injecting a dose that is too high for that particular animal. As these animals are not inbred and come from a variety of populations with an array of genetic differences, the exact response of an individual lizard to anesthesia varies. Dosages in this protocol have been optimized but adjust if needed. If the animal stopped breathing within the first 30 seconds of receiving anesthesia and the body went limp almost immediately after the injection, the lizard has most likely died and is not recoverable. Lizards rarely crash more than five minutes after receiving anesthesia unless they were recently given meloxicam while their respiration is low (below 16 breaths per minute). Lizards almost never crash during surgery if they have maintained good respiration after drug administration. Their breathing should remain consistent throughout the surgery until they begin to leave the surgical plane of anesthesia. The most common time for respiration to be disrupted is between 3 and 5 minutes after the administration of anesthesia; many of these lizards recover. The following procedure can aid recovery:

• 1Begin by positioning the lizard under the microscope, dorsal side up, with the forelimbs extending towards the head (Figure 7). Using two pairs of butterfly forceps, grasp the skin posterior to the forelimbs and pull it laterally—as far as it will comfortably go. Hold there for 5 seconds, then push medially as far as it will go without applying too much pressure to the ribs. Repeat this sequence at least 3 times every minute.

Figure 7: Manual respiration support for a lizard in respiratory arrest.

A lizard that has stopped breathing should be positioned on its ventral side with its forelimbs raised towards the head. This is to allow the butterfly forceps easier access to the posterior of the forelimbs, where manual respirations should be performed. The heart rate can be seen anterior and posterior to the forelimbs, usually in some combination of these four positions.

The objective is to push oxygen in and out of the lungs.

• 2Between manual respirations, observe the heart rate. With a stereomicroscope and careful inspection, the heartbeat can be observed as a fluttering motion directly anterior and posterior to the forelimbs (Fig. 2). The faster the heartbeat and the more apparent it is in the anterior and posterior positions, the more likely it is that the animal will recover.
• 3Continue manual respirations until the animal begins to breathe independently again, or until the heartbeat stops. Most animals that begin breathing again do so within about thirty minutes, although it can take an hour or more.
• 4Once the lizard’s respiration has increased to over 16 breaths a minute, proceed to surgery. In our experience, these lizards suffer no obvious side effects post-surgery of having stopped breathing, other than bruising at the site where the forceps were holding the skin to provide manual respiration.

Keep a careful eye on the respiration of these lizards during their surgery, as the length of time they will stay unconscious is less predictable, and they may need additional anesthesia. If additional anesthesia is needed, decrease the original dose that caused them to crash by 10–20%.

Problem (Step 17): The lizard does not become unresponsive after anesthesia is administered.

Solution: If the lizard has not reached a surgical plane of anesthesia within five minutes, a second smaller dose of anesthesia can be given. The amount of anesthesia in this second dose will vary depending on how responsive the lizard is to the tail clamp. The following are guidelines based on our experience but can be modified based on your results: If the lizard can move its whole body: 2/3 the original dose; If the entire body twitches: 1/3 the original dose; If there is only limb or digit twitching: ¼ the original dose.

Problem (Step 20): The skin cut is placed too far anteriorly or posteriorly relative to the follicles.

Solution: Only one skin incision per side should be made along the flank. However, the location of the cut in the underlying rib musculature can be adjusted if the initial skin incision is not ideally placed. Once the skin incision is made, typically 2–3 ribs are visible and so there are a few options of which ribs to cut between. To increase the likelihood of accessing the follicles, cut the rib musculature layer closer to the prospective location of the follicles. Keep in mind that the greater the distance between the incision at the rib musculature layer and the skin incision, the smaller the working space within the lizard.

Problem (Step 21): A rib is broken.

Solution: This can be caused by pinching a rib with forceps or angling the initial skin cut too deeply, causing the iris scissors to cut a rib in addition to the skin layer. Anolis sagrei are remarkably hardy. Usually, surgery can continue as normal, but the female should be solo-housed for the first few days upon completion of surgery, for closer observation.

Problem (Steps 21–23): There is blood hemorrhaging soon after making a cut or maneuvering internal organs or follicles.

Solution: This usually happens for one of two reasons. The first, is that one of the many small blood vessels in the skin or rib musculature was nicked or severed while cutting into the coelomic cavity. This is rare but does happen and is not predictable. To remove the blood, Weck-Cels or a comparable absorbent material can be used to absorb the fluid. Typically, only a small amount of blood seeps out from these injuries, and surgery can proceed normally. The second reason for a hemorrhage during surgery is if an internal organ or follicle is moved too far from its initial position within the body cavity, causing the blood vessels attached to it to break. This can cause a substantial amount of blood to be released into the coelomic cavity. As before, if the blood can be removed, the surgery can be continued; however, the flow of blood into the coelomic cavity often makes it too difficult to find and inject follicles. Surgery can be attempted on the other side although it is not always successful because of poor visibility. Post-surgery, solo-house these lizards for closer observation. We have observed no obvious difference in the survival rate of lizards that have hemorrhaged compared to other post-surgery lizards.

Problem (Step 23i): Follicles cannot be found.

Solution: This can be caused by looking in the wrong location or because the lizard is not producing eggs. If there is a lung (fairly translucent, honeycomb or bubbly-looking tissue) visible, look posteriorly. If there is a visible fat pad (off-white/light yellow, flat, very long tissue with a reflective sheen due to the fat droplets), look dorsally from the ventrally-located fat pad; however, sometimes lizards have very large fat pads that will cover their entire side. While lungs should not be touched, fat pads can be pushed or cut out of the way while searching for follicles. If there is an egg, follow the oviduct from the anterior of the egg to the follicles. Sometimes lizards are not actively laying. These could be recently captured lizards, smaller lizards, or lizards with very large fat pads. Even if a lizard is not actively laying, they should have a few small (1 mm or less) follicles still attached to the oviduct, but their location can vary. If only small follicles are observed, then the incision should be closed without attempting to inject any follicles.

Problem (Step 23ii): Maneuvering follicles is damaging them.

Solution: The cluster of small follicles can be maneuvered by grasping individual follicles (which may burst so, ideally, only hold follicles too small to inject) or the membrane between them. The largest follicle, which often is yellower due to increased yolk content and over 5 mm, should not be grabbed, as it can easily burst open and then greatly reduce visibility. In this scenario, Weck-Cels or a comparative absorbent material can be used to increase visibility.

Problem (Step 23): The small follicles cannot be found, only the large yolky one.

Solution: If only a single, large yolky follicle can be seen, roll it around carefully to find the smaller follicles, as all follicles are attached to each other. Rolling can be done by dragging angled forceps along the side, starting more medial and dragging laterally. As the follicle rolls slightly, look along the newly exposed region for any follicles or change in the pattern of the blood vessels along the surface, as this can be easier to see than small follicles. Once they are visible, carefully grab one of the smallest follicles to hold them in place before injecting as usual. If there is tension, do not force the small follicles further laterally, as this will rip blood vessels and cause hemorrhaging.

Problem (Step 24): The needle becomes clogged when injecting yolky follicles.

Solution: When injecting yolky follicles, yolk can come up into the needle and harden, causing it to clog. The frequency with which this occurs will depend on the holding pressure and needle diameter, but it can usually be avoided by ejecting the yolk in the end of the needle as soon as the needle is removed from a yolky follicle. If the needle becomes clogged, break the clogged portion off by tapping the tip of the needle on the forceps away from the lizard. If the diameter of the needle after breaking the tip exceeds 30 μm, consider changing to a new glass needle, as it becomes difficult to puncture follicles. Be sure to thoroughly clean the glass off the forceps before continuing.

Problem (Step 26): It is difficult to keep the layers of tissue aligned after surgery for closure.

Solution: This is especially difficult with lizards that have eggs and/or large yolky follicles because of the increased tension on the tissue layers. To reduce the tension on the side, raise the lizard’s head so the animal is bent at a 45-degree angle and hold there while the glue is applied and dries. Additionally, a drier surface will allow layers to stick to each other a little bit, so not applying Plasma Lyte unless necessary can help the layers stick in place during closure.

Problem (Step 27): The VetBond dried much faster and much whiter than normal.

Solution: VetBond solidifies when exposed to water, so a particularly wet surface of the lizard will make the glue solidify almost immediately. This is also true if a large yolky follicle was popped; the mixing of this fluid with the VetBond will cause the VetBond to dry almost instantly. We have observed no noticeable difference in the healing of animals that had VetBond that dried quickly versus those where VetBond dried more slowly, so efforts (beyond drying the skin of the lizard) to avoid this phenomenon have not been attempted.

Problem (Step 36): A surgical incision has opened in the days following surgery.

Solution:

• 1If most of the surgical incision has opened, prepare the surgery station as before with the following exceptions: Cotton applications, Povidone-Surgical Scrub Iodine, and 70% v/v ethanol are not needed for scrubbing the lizard. Instead, in a 50-mL sterile conical tube, add a few drops of 10% Povidone Iodine Solution First Aid Antiseptic and ~10 mL of Plasma-Lyte. As there is no microinjection taking place, there is no need for RNPs, needles, the microinjection rig, or the air compressor.
• 2Weigh the lizard and provide anesthesia as done previously (Step 19). The normal wiggling often seen when providing anesthesia to lizards coupled with holding the lizard in place can result in organ/follicles pushing out the surgery incision. Do not stop the anesthesia injection if this occurs. The organs or follicles will be returned to the coelomic cavity later (see below).
• 3Wait until the lizard is unresponsive to a tail clamp. Clean the open surgical incision site by bathing it with at least 3 mL of the iodine and Plasma Lyte solution.

Avoid filling the coelomic cavity with too much fluid as this can be dangerous to lizards.

• 4Inject meloxicam into the shoulder as done previously.
• 5Use forceps to pick off all the dried VetBond from the original closure.
• 6If any healthy tissue and/or follicles are outside the body cavity, push them back into the coelomic cavity using sterile forceps. If anything is sticking to the healthy tissue and/or follicles such as dirt or dried VetBond, remove that as well.

If follicles found outside the body do not look healthy, they can be removed. Use forceps to pinch hard at the junction between the follicles and the surrounding tissue. This helps to crush the blood vessels leading to the follicles and avoid unnecessary blood loss. After this is done, detach the cluster of follicles from the lizard.

• 7Close the lizard as in Steps 35–37. Follow the steps described above under “Animal Recovery” but house the lizard separately in a small cage for 3 days before transferring back to its previous cage of post-surgery females.

DISCUSSION

Using this protocol, we have successfully generated mutant alleles for a dozen loci, including tyr (Rasys et al. 2019; Borteiro et al. 2021) and tfec (Garcia-Elfring et al. 2023). Because CRISPR-Cas9 is used here without a homology donor, the expectation is that error-prone repair mechanisms will yield small insertions and deletions (indels) at targeted sites; that is indeed generally what we observe. Most mutant alleles generated with this method carry indels that are under 30 base pairs, and deletions are more commonly generated than insertions. We have, however, sporadically detected indels of several hundred base pairs, and have also identified a few rearrangements among the mutant alleles we have produced (Garcia-Elfring et al. 2023). Typically, 5% to 15% of screened offspring carry monoallelic or biallelic mutations. We do occasionally see mosaicism with some loci. These mosaic animals are often collected earlier in the screening process, around 3 weeks after surgery. In cases where co-injected sgRNAs showed variable success in disrupting open reading frames, we use only the most effective sgRNA in subsequent rounds of gene editing surgeries. While sgRNAs can vary in their efficiency, we find the main factor impacting our efficiency is the experience and skill of the surgeon. A proficient surgeon is able to quickly perform surgeries, minimizing anesthesia time, and can accurately find and handle follicles, avoiding damage to the lizard.

While this protocol employs Cas9 to generate targeted mutations, we expect that other Cas systems, including base editors, would work with this surgical approach to gene editing. In addition, this method has been modified to create targeted mutations in the Madagascar ground gecko (Abe et al. 2023), and this approach has also been adapted for the production of gene edited corn snakes, showing that the method can work in species that produce larger oocytes and large clutches of eggs (Tzika et al. 2023). Together, the gene editing results in anoles, geckos, and snakes suggest that injection of oocytes through a surgical incision is a general approach that may work in many different reptile species.

Table 1. Drug information for surgeries.

Information about each drug used during surgeries is given. The stock concentration and the needed volume of stock per gram of lizard are listed so drugs can be accurately administered. The storage of each drug is also considered, with the required temperature being listed below while the expiration date is found on the purchased bottle itself.

Drug	Drug Stock Concentration from Manufacturer	Dose of drug used per gram of animal weight	Volume of stock used per gram of animal	Storage Temperature
Dexmedetomidine hydrochloride (Dex)	0.5μg/μL	0.1μg/g	0.2μL/g	RT
Alfaxalone (A)	10μg/μL	30μg/g	3.0μL/g	RT
Meloxicam (M)	5μg/μL	0.3μg/g	0.06μL/g	RT
Carprofen Can be used as an alternative to M	50μg/μL	20μg/g	0.4μL/g	4°C
2% Lidocaine (L)	20μg/μL		1μL/g	RT