The Golden Apple Snail Pomacea canaliculata: From Zygotes to Stable Mutant Lines

Summary

This study developed a workflow for the generation and maintenance of stable mutant lines for the freshwater apple snail, Pomacea canaliculata. The protocols include the collection of zygotes, their microinjection with the CRISPR/Cas9 system components to induce genome editing, ex ovo culture of embryos, and genotyping after controlled crosses.

Abstract

The freshwater golden apple snail (Pomacea canaliculata) is emerging as a powerful model organism for regenerative biology, particularly for studying the molecular mechanisms of sensory organ regeneration. They can fully regenerate their camera-type eyes, which have vertebrate-like anatomical features such as retina, cornea, and lens organized in a closed chamber. Coupled with a relatively small diploid genome, genetic tractability, and ease of laboratory maintenance, P. canaliculata offers unique advantages for studies beyond developmental and regenerative biology. This paper presents a comprehensive workflow for generating and maintaining apple snail mutant lines, making these animals one of the very few genetically tractable mollusks available. The possibility of knocking out genes of interest opens the door to understanding the genetic and molecular basis of sensory organ development and regeneration, as well as other aspects of molluscan biology. Here, we provide detailed protocols for the effective maintenance of a large apple snail colony, collection and microinjection of their zygotes, the use of perivitelline fluid for successful ex ovo culture, and techniques for genotyping and maintaining stable mutant lines. By enabling targeted genetic manipulation in this novel model and disseminating methods for functional genomics in mollusks, we aim to encourage the community to grow and adopt apple snails for further studies across biological fields.

Introduction

The golden apple snail, Pomacea canaliculata, is emerging as a valuable model organism in regenerative biology due to its remarkable ability to regenerate complex organs, including its camera-type eyes, after complete amputation¹ (Figure 1A). P. canaliculata’s camera-type eyes, comprising a retina with photoreceptors, a cornea, and a lens organized in a large single chamber, closely resemble vertebrate eyes, making it an ideal and almost unique system for investigating the molecular underpinnings of this complex sensory organ, complete regeneration^1,2,3,4,5,6. In addition to these incredible biological features, its diploid genome, genetic tractability, direct development (lack of free-living larval stage and metamorphosis), number of embryos available daily, and ease of laboratory maintenance enhance its suitability for molecular and cellular research^6,7,8,9,10 (Figure 1B,C).

Stable mutant lines offer a powerful means to probe gene function. The CRISPR/Cas9 system enables easier genetic manipulation of new species of interest^{11,12,13,14,15,16,17}. Some major challenges remain when applying CRISPR/Cas9 mutagenesis to a new species, including obtaining one-cell stage embryos, performing injections without compromising their viability, and raising the injected animals to sexual maturity. All these methods in P. canaliculata have been recently developed from scratch, significantly increasing the potential of this new system to study the genetic basis of camera-type eye development, regeneration, and evolution^2,3,6,18. Genome editing and germline transmission are the first steps to the development of transgenic reporter lines and conditional mutants¹⁹. Such protocols are both timely and essential, given the relative novelty of applying these techniques to mollusks and the interesting biological features of P. canaliculata. These tools will provide essential resources for researchers studying P. canaliculata and for those aiming to apply similar methodologies to species with similar reproductive or developmental traits.

P. canaliculata populations consist of distinct males and females^20,21 (Figure 1D,E). After mating, the females store the sperm to internally fertilize the oocytes shortly before laying the zygotes²⁰. Apple snails lay large clutches, each containing about 100–300 embryos^20,21 (Figure 1B). Each embryo is housed in a capsule that has a hard outer casing and is filled with pink perivitelline fluid (PVF)²². The PVF provides key nutrients and protection to the developing embryo, while the outer layer prevents dehydration²². Capsules are laid outside the water, and they go through important changes during the first 24 h; freshly laid eggs transition from a soft external membrane and a dense and opaque PVF to a harder casing and a less dense, more transparent PVF²⁰ (Figure 1B and Figure 2). Zygotes go through the first cell division approximately 6 h post fertilization (hpf), providing ample time for the microinjections⁶.

This study presents all the methods needed to generate stable mutant lines in P. canaliculata. The pipeline includes maintenance of an apple snail colony, efficient collection of healthy zygotes, and their microinjection using an inverted microscope equipped with micromanipulators and a MICRO-ePORE Pinpoint Cell Penetrator. This work further describes an innovative procedure, embryo ex ovo culture, for culturing embryos outside their capsules. Central to this is the isolation of the PVF, the natural environment for apple snail embryonic development, and its quality control for successful ex ovo embryo culture. Finally, methods for growing and genotyping apple snails, as well as a guideline for designing the crossing for long-term maintenance of the mutant lines, are provided.

By establishing and detailing these protocols, the groundwork is laid for generating stable mutant lines in apple snails, opening avenues for genetic studies in a new, yet highly promising, invertebrate system. At present, the application of genetic manipulation to this model stands to significantly advance the understanding of sensory system development and regeneration. Considering this is one of the very few mollusks where stable mutant lines can be established to date^{11,14,15,16,17}, the vision is that this system will be adopted in other biological fields to study their physiology and specializations. 

Protocol

All the protocols described in this manuscript follow the guidelines of the UC Davis Ethics Committee and Animal Care and Use Committee. The reagents and the equipment used are listed in the Table of Materials.

1. Apple snail P. canaliculata colony maintenance

1. Colony culture
   1. House the snails in 26 °C freshwater in tanks with a 14:10 h light-dark cycle. Maintain the osmolarity of the water at 900 µS with Instant Ocean Sea Salt:Calcion-P:Magnesion-P in a ratio of 6:10:1 in distilled water (dH₂O), and the pH at 7.5–8.0 using Rift Lake dKH. If a self-dosing pump is not used, the salts can be diluted to a final concentration of 153.8 mg/L Instant Ocean Sea Salt, 256.4 mg/L Calcion-P, 25.6 mg/L Magnesion-P, and 128.2 mg/L Rift Lake dKH.
      ​NOTE: At UC Davis, the apple snail colony is maintained in a custom-made water recirculation system built by Iwaki Aquatic (Figure 1C). Housing too many snails in a limited volume can cause reduced growth and animal stress. While hatchlings and juveniles can tolerate higher densities, keep adults at a maximum of 4 snails/L (Table 1). If static water is used, keep snails at a lower density (2 snails/L) and replace a higher percentage of water (30–40%) daily. P. canaliculata is native to South America and is listed as an invasive species in many countries. Permits for importing and maintaining them might be needed, and the space where the snails will be housed might have to be inspected.
   2. Replace 5–10% of the water daily and siphon the bottom of the tanks the day after feeding to maintain high-quality water.
   3. Feed the snails three times a week with organic lettuce. Feed hatchlings and juveniles ad libitum. Feed the adult snails 1 large leaf (about 8 × 8 in) for every 10 snails, and the old snails 1 large leaf for every 20 snails (Table 1).
      ​NOTE: High-quality water and regular feeding give a higher chance of getting frequent and healthy clutches. Hydroponic lettuce seems to reduce egg laying.
   4. Collect the egg clutches daily and place them in Petri dishes or dispose of them. The clutches are laid on the walls and lids of the tanks, and they are easier to detach in the first 24 h after they are laid (Figure 1B,C, and Figure 2A).
      ​NOTE: For optimal development, carefully transfer freshly laid clutches from the tank walls (high-humidity environment) to a place where they can fully dry. At 27 °C, hatching occurs approximately 10–­13 days post fertilization (dpf). During this period, the eggs gradually lose their vibrant color, transitioning from bright pink to a pale gray, which shows the depletion of PVF as it is utilized by the developing embryos (Figure 1B).
   5. Once the clutches are mostly gray in color, gently crush them in between fingers immersed in system water to manually hatch them. The outer casing floats while the released hatchlings sink.
2. Individual sex determination
   1. Place 1.5–2.5-month-old snails in ice for 10 min to relax their muscles. At this age, the snails are already showing sexual dimorphism, but they have not started mating yet.
   2. Using a hemostat locking scissor clamp, pull the operculum anteriorly, ventrally, and then posteriorly to expose the head of the snail.
   3. Observe the internal lining of the shell, on the snail’s right margin of the mantle. Use a bright light or a stereomicroscope to better observe the anatomy of that region.
      ​NOTE: In that region, males possess a penis-sheath, visible as a white triangular and posteriorly elongated structure²³. At the base of the penis-sheath, there are a small bright pink round structure (penial bulb) and a larger pale orange one (penis pouch)²³ (Figure 1D). On the right side of females, the ends of two tubular structures can be observed — a small one (the oviduct) and a slightly larger one (the rectum). The oviduct is the only female-specific structure visible in this region (Figure 1E).
   4. Once the snails are sexed, place them back in the system water, separating them if needed. Alternatively, use nail polish or non-toxic glue to mark their shells.
      ​NOTE: If the mantle detaches from the shell during the operation, do not stretch the snail further. Quickly complete the procedure if possible and place them back in the system water (Figure 1F). The mantle detachment appears to be a sign of stress for the snails and increases the possibility of death.

Stage	Age	Food provided	Notes
Embryos	0 - Hatch (12-13 dpf or 0 dph)	None	Embryos are inside the capsules. Keep the capsules dry.
Hatchlings	0 - 3 dph	None	-
Hatchlings	3 - 15 dph	Lettuce ad libitum	3-7 clutches hatched together to reduce inbreeding
Juveniles	15 dph - 3 mph	Lettuce 3 times/week	Population density of 15 snails/liter
Adults	3 - 12 mph	Lettuce 3 times/week	Population density of 2 - 4 snails/liter
Adults	12 - 24 mph	Lettuce 3 times/week	Significant decrease in offspring
Old snails	> 2 years	Lettuce 3 times/week	No offspring

Table 1: Husbandry conditions based on P. canaliculata life stage.

Figure 1: Morphology and sexual dimorphism in P. canaliculata. (A) Lateral view of an adult P. canaliculata. (B) P. canaliculata egg clutches. These clutches change colors from bright pink to gray as the developing embryo consumes PVF (black arrowheads). Clutches with crushed and drying regions are unlikely to yield healthy embryos. Sterile capsules are more fragile and dry more easily, causing many of them to be crushed. (C) Schematic representation of the water recirculation system used to house a large P. canaliculata colony in the laboratory, with a limited number of hours dedicated to its maintenance. (D,E) Sexual dimorphism between adult female and male snails. Differences in the gonads can be observed through the shell, and differences in the external reproductive organs can be observed after anesthesia. (D) The male digestive gland is covered by a large and thick gonad (white dashed line), resulting in a shell spiral lighter in color. In anesthetized males, on their right side, close to the margin of the mantle, a bright pink penial bulb, an orange penis pouch, and a penis-sheath that is usually stored posteriorly can be observed. All these organs are highlighted by white dashed lines. (E) Females show a dark color in the shell spiral mainly due to the very thin gonad (white dashed line) located above the black digestive gland. In anesthetized females, a smooth margin of the mantle on their right side can be observed, where there are only the ends of the oviduct and the rectum. (F) If the operculum is pulled too strongly, the mantle might detach from the shell. e = eye; m = mouth; f = foot; t = tentacle. Please click here to view a larger version of this figure.

Figure 2: Progressive morphological changes in freshly laid P. canaliculata egg clutches. (A) Freshly laid clutch still covered in mucous and with soft capsules attached to the wall of a tank. (B) The capsules from a freshly laid clutch are uniform in their opaque pink color. As the embryo develops, a transparent pink layer forms and grows from the periphery inwards. At (C) 4 and (D) 12 hpf, two distinct areas are visible: the initial opaque pink layer in the center and the transparent pink layer increasing in size in the external region. Please click here to view a larger version of this figure.

2. Testing the Perivitelline Fluid Extract (ePVF)

1. ePVF collection
   1. Clean the workspace and the tweezers with 70% ethanol and ensure that consumables (e.g., Petri dishes and tubes) are sterile.
      ​NOTE: Since P. canaliculata embryos cannot complete their development in PBS or other commonly used media, collect ePVF aliquots from the capsules to artificially recreate an environment that can support the apple snail embryonic development. Due to the rich PVF organic content, sterility is critical to avoid contamination²⁴. Use pipette tips with filters. If available, a laminar flow hood can be used.
   2. Transfer 1–2 dpf clutches (1 dpf is preferred) into a 50 mL tube containing 40 mL of 0.8% sodium hypochlorite (NaOCl, or commercial bleach) for 3 min. Always prepare the sodium hypochlorite solution fresh using autoclaved dH₂O. Do not excessively shake the tube. Lay the tube horizontally during the incubation.
      ​NOTE: Sodium hypochlorite is considered a hazardous chemical that can cause severe skin burns and eye damage. Wear protective gloves, protective clothing, face protection, and eye protection such as safety glasses. Dispose of contents in accordance with all applicable federal, state, and local regulations. Do not allow products to enter storm drains, lakes, or streams.
   3. Quickly rinse the clutches twice with autoclaved dH₂O and then incubate in fresh autoclaved dH₂O for 2 min to make sure to remove all sodium hypochlorite residuals. Lay the tube horizontally during the incubations.
   4. Remove the autoclaved dH₂O from the 50 mL tube and move the dry clutches to a Petri dish using forceps.
      ​NOTE: Keep each clutch separate. If one clutch is of bad quality, that will decrease the overall ePVF quality. The quality of the ePVF is going to be determined in step 2.2. Contaminated ePVF and ePVF where embryos do not develop properly are considered “bad quality”.
   5. Using tweezers, crush all the capsules of the clutch (Figure 3A).
   6. Transfer the crushed material into 2 mL centrifuge tubes using curved forceps (Figure 3B). The consistency of this material makes it easier to pick it up with forceps than pipette it.
   7. Centrifuge the tubes at 21,000 × g for 40 min at 4 °C.
      ​NOTE: Three layers should be evident after centrifugation: (a) capsule debris in the bottom, (b) an opaque and fuzzy pink layer in the middle, and (c) a transparent and clear pink layer on the top (Figure 3C).
   8. Transfer the top layer (c) into a fresh 1.5 mL tube without combining ePVF obtained from different clutches (Figure 3D,E). In an attempt to collect the entire top layer, part of the middle layer may be collected too. This will not hamper the quality of the ePVF.
   9. Store the ePVF aliquots at −20 °C until ready to use them.
      ​NOTE: The ePVF can be frozen and thawed up to five times without losing its efficiency. A higher number of times was not tested.
2. Testing the collected ePVF with wild-type embryos
   ​NOTE: Test the quality of the ePVF aliquots on wild-type embryos before using them to culture injected embryos, to reduce the growth of bacterial or fungal contamination and increase the success of proper embryo development.
   1. Clean the working area and sterilize the clutches following the steps 2.1.1–2.1.5 on 2 dpf clutches to collect wild-type embryos for ePVF testing.
      ​NOTE: Clutch separation is required only during ePVF collection. Clutches do not need to be separated here since only the embryos are isolated.
   2. Add 40 mL of P. canaliculata Embryo Medium (Pc-EM) (filtered 6 mM KCl, 6.6 mM CaCl₂, 3.3 mM MgCl₂, 1 M HEPES, 33.4 mM NaCl in dH₂O) to the Petri dish and use tweezers to mix the broken capsules and dislodge the embryos from the PVF (Figure 4A–C).
      ​NOTE: HEPES is known to be mildly hazardous and causes respiratory irritation if inhaled. Avoid breathing dust and use only outdoors or in a well-ventilated area. Wear appropriate protective eyeglasses, a lab coat, and gloves as described by OSHA's eye and face protection regulations. HEPES should not be disposed of in the environment. Dispose of contents/container in an approved waste disposal area.
   3. Under a stereomicroscope with transmitted light, collect embryos released from the capsules and move them to a Petri dish containing Pc-EM using a P20 pipette. With transmitted light, the embryos look round and dark and are easier to spot in the middle of capsule debris and PVF (Figure 4D).
   4. Move the embryos to a clean Petri dish (or in a 4-well plate) pre-filled with 5% fetal bovine serum (FBS) in Pc-EM. Swirl the embryos to clean them of debris and PVF. Repeat wash two more times (Figure 4E).
      ​NOTE: P. canaliculata embryos sink to the bottom of the Petri dish at all points of development. They stick to glass and plastic up to 3 dpf. FBS in the media reduces the stickiness. It is good practice to pipette the 5% FBS in Pc-EM up and down before collecting embryos to reduce stickiness inside and to the tip.
   5. Place the lid of a 35-mm Petri dish facing up inside a 60-mm Petri dish, after labeling it with a permanent marker as a reference for the different ePVF aliquots (Figure 4F).
   6. Pipette two drops 3 mm in diameter (about 60 µL) from each aliquot of ePVF into the 35-mm lid using a P200 pipette (Figure 4G).
   7. Using a P20 pipette, place 3–4 healthy embryos in each ePVF drop, minimizing as much as possible the volume of Pc-EM carried over and avoiding bubble formation (Figure 4E,H).
      ​NOTE: If the ePVF drops are too small or if too much Pc-EM is carried over with the embryos, the culture success decreases. Avoid as much as possible to dilute the ePVF.
   8. Gently cover the drops with 3 mL of paraffin oil to prevent evaporation and make sure that all the drops are fully covered (Figure 4I). Avoid drops combining together by pouring oil in between adjacent drops initially, and then covering all drops.
   9. Place the lid on top of the 60-mm dish and incubate at 27 °C in the dark.
   10. Check daily for embryo development, air bubbles forming on top of the ePVF drops, and presence of contaminations (Figure 5). In case of large bubbles or several small bubbles, remove them gently with a P20 pipette (Figure 5C,D). This will prevent the drying of the ePVF drop (Figure 5E).
   11. After 8–10 days, check the ex ovo culture (Figure 5B). In case of contamination or abnormal embryonic development (Figure 5A,F,G), dispose of the corresponding ePVF aliquots. In case of full embryo growth and lack of contamination, save the ePVF aliquot for future culturing of injected embryos.

Figure 3: Collection of ePVF for ex ovo culture. (A) Partially crushed clutch showing intact (top) and broken (bottom) capsules. (B) Crushed 1 dpf clutch transferred to a 2 mL microcentrifuge tube prior to centrifugation. (C) Post-centrifugation separation into three distinct layers: a. capsule casing and debris, b. intermediate opaque pink layer, and c. clear ePVF supernatant. (D) Left over in the microcentrifuge tube after pipetting the supernatant out using a P1000 pipette. (E) ePVF collected in a microcentrifuge tube and ready to be tested or frozen. Please click here to view a larger version of this figure.

Figure 4: Ex ovo culture setup for ePVF quality assessment. For testing the quality of the ePVF, (A) a 2 dpf clutch is placed in a Petri dish with Pc-EM and (B) crushed. (C) The broken capsules are then stirred in the Pc-EM to separate the capsule casing, the PVF and the 2 dpf embryos. (D) Isolated embryos sitting at the bottom of the Petri dish among crushed capsule debris. (E) Healthy and damaged 2 dpf embryos. The damaged embryos have extruding cells. (F) Labeled 35-mm Petri dish lid inside a 60-mm Petri dish for setting up ex ovo embryo culture. (G) ePVF droplets arranged in quadrants with optimal volume and spacing. (H) Successful transfer of three healthy 2 dpf embryos into the ePVF droplets. (I) Completed ex ovo culture setup with paraffin oil overlay to avoid ePVF evaporation. Please click here to view a larger version of this figure.

Figure 5: Assessment of the ex ovo embryo culture. (A) ePVF droplet with a healthy embryo (5 dpf) and an embryo that is not developing properly. Due to intrinsic factors, these two events can sometimes be found in the same droplet. (B) Fully developed embryos at 9 dpf. The embryos at this stage pool together debris present in the droplet and potential contamination. Droplet with one (C) and multiple (D) air bubbles. Air bubbles larger or more numerous than those illustrated will cause the droplet to dry (E). A low level of paraffin oil can also lead to drying of ePVF. A solidified and concentrated center is the first sign of drying ePVF, followed by a thinner and more transparent periphery of the droplet. (F) Example highlighting the difference between an early-stage embryo (3 dpf) and an early sign of contamination. Embryos from these droplets can still be recovered. (G) Fully contaminated ePVF droplet. Embryos from these droplets cannot be recovered. High-quality ePVF is defined when most of the embryos are developing properly, and there is no contamination emerging. Please click here to view a larger version of this figure.

3. Microinjection of P. canaliculata zygotes

1. Preparation of injection materials
   1. To obtain needles, pull 10 cm borosilicate glass capillaries with filament using a horizontal needle puller to obtain needles with a 6 mm long tip.
   2. To obtain holding pipettes, pull 10 cm borosilicate glass capillaries without filament using a horizontal needle puller.
      ​NOTE: For both the needles and the holding pipettes, the heat value is the same value obtained from each batch doing the ramp test on the needle puller.
   3. Cut the tip of the capillary obtained in step 3.1.2 and fire-polish the holding pipette with a microforge. The outside diameter should be 50–60 µm and the internal diameter 15–20 µm.
      ​NOTE: Microscopy images of the needle and holding pipette shape and size are available in the supplementary information of Accorsi et al.⁶.
2. Zygote collection
   1. Collect freshly laid clutches when the capsules are still coated with a heavy layer of mucus and the pink color is homogeneous (Figure 2A,B, and Figure 6A). Place them in a 50 mL tube filled with freshly prepared L-cysteine solution (15 g/L L-cysteine in dH₂O, pH 7.5 with NaOH) and incubate for 3 min.
      ​NOTE: Clutches are usually laid at the end of the night and early in the morning. The oocytes are fertilized right before the capsules are laid, and the first cell division happens about 6 hpf. Freshly laid capsules are soft and malleable, appear uniformly pale pink, and are encased in a soft membrane and a transparent mucus coating. In a few hours, the capsules become increasingly heterogeneous in appearance, exhibiting a brighter and transparent pink layer on the outside that grows with time and an opaque, fuzzier pink layer in the center that shrinks with time. At the same time, the mucus dries, and the external membrane surrounding the capsules hardens (Figure 1B and Figure 2D).
   2. Move 20 mL of the L-cysteine solution and the capsules to a 100-mm Petri dish and add 20 mL of Pc-EM to it.
   3. Using tweezers and a stereomicroscope at magnification 10x, open the capsules one by one, tearing the external membrane and exposing the PVF to the L-cysteine solution. L-cysteine reacts with the PVF, helping to free the embryos and keeping the solution clear to see them at the bottom of the Petri dish (Figure 6B–D).
   4. Very gently, swirl the contents of the Petri dish and let everything settle down for 2 min.
   5. Using a stereomicroscope with transmitted light, scan the dish, and collect the embryos that were released from the PVF with a P20 pipette. Then move them to a separate Petri dish pre-filled with 5% FBS in Pc-EM (Figure 6E,G).
   6. Keep repeating the steps 3.2.4 and 3.2.5 until the desired number of embryos has been collected.
      ​NOTE: Healthy embryos are spherical and dark (Figure 6G). While the PVF sticks to the bottom of the Petri dish, the embryos do not. Do not try to collect embryos still embedded in the PVF because it will damage the embryo (Figure 6F,H). Repeating multiple rounds of embryo collections, alternating it with the PVF mixing, helps to gently release the embryos.
   7. Evenly split the number of embryos into two fresh Petri dishes, both pre-filled with 5% FBS in Pc-EM. This will allow the alternation of the embryos between 4 °C and room temperature (RT) during the microinjections.
   8. To slow down the embryonic development and delay the first cell division, store the embryos at 4 °C until the microinjection station and the injection solution are ready. Avoid placing the embryos directly on ice because it decreases their survival rate (Figure 6I).
      ​NOTE: The embryos can survive for up to 1 h at 4 °C. Longer times have not been tested.
3. Microinjection of zygotes
   1. Turn ON the inverted microscope equipped with a 20x Differential Interphase Contrast (DIC) objective, two micromanipulators, a microinjector, a pinpoint cell penetrator, and a chiller connected to a cold plate and set up at 12 °C.
   2. Prepare 30 µL of injection mix (2 mg/mL Dextran Red, 0.05% Phenol Red, 10 ng/µL gRNA designed using https://chopchop.cbu.uib.no/ or https://www.crisprscan.org/ and following the developer instructions, 10 ng/µL Cas9 in nuclease-free H₂O). Centrifuge it at max speed for 5 min at 4 °C and then move the top 15 µL to a clean tube.
      ​NOTE: The centrifugation step will help to remove small particulates that could clog the needle.
   3. Load the needle with the injection mix using the tapered tip microloader or back-loading by capillary action.
   4. Mount the holding pipette and the needle on the micromanipulator at a 15° angle. Ensure that the electrode of the pinpoint cell penetrator is in contact with the injection mix in the needle. Use the 4x objective to bring everything in focus, at the center of the field of view, and close to the depression slide surface.
   5. Open the needle using the holding pipette to break the tip. Aim for about 5 µm diameter opening with a slight asymmetry in the break.
   6. Place one of the dishes prepared in step 3.2.7 and 3.2.8 at RT and the other one at 4 °C to slow down development.
   7. Add a large drop of 5% FBS in Pc-EM on the depression slide. Then fill part of the holding pipette with the CellTram Air, place the second electrode of the pinpoint cell penetrator in the drop, and move 10–20 embryos from the dish at RT on the depression slide.
   8. After pipetting the first group of 10–20 embryos on the slide on the inverted microscope, move the dish with the leftover embryos to 4 °C. At the same time, move the dish that was at 4 °C from step 3.2.8 to RT, so that it will warm up before the next round of injection.
      ​NOTE: Pipetting embryos while they are in cold media can irreversibly damage them (Figure 6I). Additionally, prolonged exposure to cold or excessive chilling can lead to developmental defects. It is important to keep alternating one dish at RT and one dish at 4 °C. While the first dish warms up for pipetting the second group of embryos on the slide, the second dish chills to delay embryonic development and increase the time available for injections.
   9. Secure one embryo at a time with the holding pipette and make sure the needle and the embryo membrane are on the same focal plane (Figure 6J).
   10. Inject the embryos (Figure 6K) using compensation pressure = 20–30 hPa and injection pressure = 90–100 hPa on the microinjector, and frequency = 520 Hz and amplitude = 0.480 V on the pinpoint cell penetrator. Once the needle is in contact with the embryo, alternate an injection and a pulse until the needle gets into the embryo.
       ​NOTE: The embryos are injected with about 20 pL, the equivalent of about 1/10 of their volume. This is a rough estimate since the needles are not calibrated, and the embryos are too opaque to see the injection mix going in. At the moment of injection, it is possible to see the membrane of the embryo stretched because of the increased internal pressure. If the needle is too dull, the embryo will die after the needle gets in (Figure 6L); if the needle is too sharp, the embryo will lyse as soon as the needle touches the membrane (Figure 6M).
   11. Withdraw the needle firmly without stalling inside the embryo and move to the next embryo injection. To keep track of the embryos that have been injected, move them to a different area of the slide with the holding pipette. Once all the embryos on the slide have been injected, move them into a Petri dish with fresh 5% FBS in Pc-EM at RT.
   12. Replace the media and the needle every two rounds of injections (20–40 embryos). The stickiness of the system increases throughout injections and can compromise the success rate of the microinjections.
       ​NOTE: All the embryos should be injected while they are still at the 1-cell stage. Embryos injected after the first cell division do not survive.
4. Growing F0 generation
   1. Incubate the injected embryos in 5% FBS in Pc-EM at RT in the dark for at least 3–4 h to let the embryos recover and go through the first two cell divisions.
   2. Monitor for cell division and successful injections, looking at the embryos under a fluorescent stereomicroscope with transmitted white light and a UV light source (Figure 7A).
   3. Before the end of the day, culture ex ovo the embryos that are both dividing and fluorescent using the ePVF that passed the quality control following the steps 2.2.6–2.2.11. If embryos were injected with different mixes or using different conditions, label the 35-mm Petri dish lid accordingly. Incubate the ex ovo culture in the dark at 27 °C. If ePVF drops are getting contaminated, use a pipette with a large opening to recover embryos, rinse them in 5% FBS in Pc-EM, and culture them again in a new ePVF drop.
   4. After about 11–13 days, when the embryos become too large for the drops, pipette them out of the ex ovo culture and move them to the Pc-Hatchling Media (Pc-HM) (1 mg/mL Galactose and 1 mM CaCl₂ in Pc-EM).
      ​NOTE: The longer the embryos stay in the ex ovo culture, the more they can grow and the higher survival rate they will have.
   5. After about 3–5 days, move hatchlings into Pc-Juvenile Media (Pc-JM) (1 mM CaCl₂ in Pc-EM) and feed them small pieces of lettuce (2 × 2 mm of the leafy green part).
      ​NOTE: When juveniles eat lettuce and defecate, it is a sign of good health.
   6. After about 7–10 days, move juveniles into system water.

Figure 6: Microinjection workflow and embryo viability assessment in P. canaliculata. (A) Freshly laid clutch submerged in L-cysteine solution, suitable for isolation of embryos younger than 24 hpf. (B) Two egg capsules, one intact (left) and another whose membrane has been opened to expose the PVF (right). (C) Clutch where all the capsules have been opened. (D) As the capsules are stirred with tweezers and the embryos are collected, the individual capsules are not visible anymore, the bottom of the Petri dish is more homogeneous, and the turbidity of the solution increases. (E) A zygote freed from the PVF and ready to be collected. (F) A zygote still embedded within the PVF. Trying to collect this embryo will result in the stretching of its membrane and death. Gently stirring the solution again with tweezers will help in releasing the embryo. (G) Healthy zygote. (H) Zygote irreversibly damaged with stretched membrane. This zygote will not be able to develop. (I) A zygote with an oval shape caused by cold-induced stress. The Petri dish was placed directly on ice (a thin layer of cardboard should be placed between the ice and the embryos). (J) Microinjection needle approaching a zygote held in place by a holding pipette. (K) Penetration of the zygote with an injection needle. Some bending of the cell membrane is expected. The needle successfully penetrates the zygotes when the membrane bounces back. (L) Excessive membrane bending caused by the microinjection needle being too dull. In this case, it is recommended to replace the needle. (M) Embryo lysis following injection with an overly sharp needle. In this case, it is recommended to break further the tip of the needle. Please click here to view a larger version of this figure.

4. Obtaining F1 and F2 generations through crossing

1. Testing the gRNA efficiency
   1. Design primers on the two sides of the gRNA cut site, preferably on the exons, since intron sequences are more variable.
   2. Take some embryos from step 3.4.4 and place them in 50 µL of Lysis Buffer (0.02 M Tris pH 8, 0.1 M KCl, 0.6% Tween-20, 0.6% NP-40) using a PCR tube (one embryo/tube).
   3. Place the tissue in the Lysis Buffer and add 0.5 µg/µL Proteinase K. Treat at 55 °C for 3 h and at 98 °C for 10 min in a thermal cycler.
   4. Vortex the tubes and use the extracted DNA as a template for the next reactions.
   5. Set up a qPCR with a High Resolution Melting Analysis to determine the presence of mutations based on the presence of multiple peaks in the melting curve²⁵. This confirms that the gRNAs are working (Figure 7A).
      ​NOTE: If two gRNAs are used and the putative deletion is large enough (more than 20 bp), it can be detected on an electrophoresis gel. A qPCR is needed if small mutations are expected²⁵.
2. Identifying good founders
   1. If the gRNAs are working, raise the remaining embryos from step 3.4.4, following the steps 3.4.5–3.4.6. and step 1.1 (Figure 7A). If the gRNA is not working, design and test new gRNAs.
   2. Before the injected animals (F0s) become sexually mature (120 dpf), determine their sex as described in section 1.2 and pair them one male with one female. If there are unpaired F0 females or F0 males, pair them with wild-type snails of the opposite sex.
   3. Collect clutches from each pair and label them to know which F0s they are coming from.
   4. Collect some embryos from each clutch following 2.2.1–2.2.4 and individually genotype them as described in the steps 4.1.2–4.1.5 (Figure 7A).
   5. For the samples of interest, run a PCR using the primers designed in 4.1.1, purify the product using a PCR purification kit, and sequence it through Sanger sequencing to determine the specific indels present in each founder. A good founder is an animal with germline transmission and a mutation that causes a frameshift and an early stop codon (see the Representative Results section).
      ​NOTE: Several companies offer Sanger sequencing services. Prepare the samples following their specific guidelines.
   6. Save the clutches that had embryos with mutations (offspring of a good founder) and discard the clutches that had embryos without mutations or with a mutation that does not cause a frameshift. Save and hatch clutches coming from the good founders.
3. Genotyping F1 for the identification of heterozygous snails and crossing them to obtain F2
   1. Hatch the clutches coming from a good founder and raise the F1s.
   2. When the F1 juveniles are at least 6 mm in diameter, move them to a separate container, wait for them to expose their tentacles, and cut the tip of one tentacle with micro-scissors (Figure 7B).
   3. Place the snail in a 6-well plate and the tentacle in a PCR tube in 50 µL of Lysis Buffer. Make sure that it is possible to determine which snail each tentacle is coming from.
   4. Process the samples and run a PCR or qPCR as described in steps 4.1.3–4.1.5 and identify the wild-type and the heterozygous snails.
      ​NOTE: If the putative deletion is large enough, the heterozygotes will show two bands on the gel while the wild types will show one band. For small mutations, a qPCR High Resolution Melting Analysis is needed, where wild types will have one peak, and heterozygotes will have two peaks. After sequencing the samples, a qPCR probe assay can be designed and used instead of a PCR or qPCR High Resolution Melting Analysis.
   5. House at least 10–15 identified F1 heterozygous snails together in system water and raise them until they reach sexual maturity.
      ​NOTE: The clutches laid by the identified F1 snails will have 25% wild-type snails, 50% heterozygous snails, and 25% homozygous snails for the mutated gene, unless genes that do not follow the standard Mendelian distribution are being studied (Figure 7A).
   6. Analyze F2s at the time points of interest to evaluate phenotypes in the different genotypes. The genotypes can be confirmed following the steps 4.3.2–4.3.4.
4. Outcrossing heterozygous animals with wild types for line maintenance
   1. Select females before they reach sexual maturity, i.e., younger than 3 months old, and maintain them in a tank without males.
      ​NOTE: Female apple snails can store sperm for up to 5 months²⁰. Separating females before breeding ensures controlled crosses.
   2. House wild-type females and heterozygous mutant males in the same tank before the males are 2 years old. Snails older than 2 years are considered old, and they breed significantly less.
      ​NOTE: For long-term maintenance of stable mutant lines, the animals will be outcrossed from this generation onward, since inbreeding reduces fertility.
   3. Genotype the offspring as described in step 4.3, expecting 50% wild types and 50% heterozygous for the mutated gene (Figure 7A).
   4. Select the heterozygous animals and house them in a new tank representing the new generation.

Figure 7: Crossing strategy and genotyping workflow in P. canaliculata. (A) Schematic representation of the breeding workflow used to generate, analyze, and maintain genetically modified lines of P. canaliculata. After Cas9 and gRNA microinjection into zygotes, the success of the injections and the efficiency of the gRNA are tested. If these steps are confirmed to work, F0 embryos are raised to adults and bred in pairs. A PCR/qPCR followed by product sequencing will confirm germline transmission and the size of the mutation. F1 animals from the founder are raised and used for incross breeding to obtain homozygous mutants or for outcross breeding for line maintenance. (B) Tissue sampling from a juvenile snail approximately 0.6 cm in size. A small portion of the cephalic tentacle is excised for genotyping. Please click here to view a larger version of this figure.

Results

The possible outcomes of the ex ovo culture protocol are illustrated in Figure 5. To test the quality of the ePVF, 2 dpf embryos were isolated and cultured in droplets of ePVF, overlaid with paraffin oil (Figure 4). Viable embryos maintain a spherical morphology and clarity of tissues, and they grow with significant changes in their morphology every 24 h. Successful culture is represented by embryonic growth and the formation of some air bubbles on...

Discussion

This series of protocols provides a comprehensive framework for the generation and maintenance of P. canaliculata mutant lines, providing the scientific community with a new genetically tractable invertebrate model⁶. Now, this animal can be leveraged to significantly advance the studies in several fields beyond the camera-type eye regeneration one, such as comparative spiralian embryology, environmental toxicology, and comparative molluscan physiology. The methodology integrates classic g...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1.7 mL microcentrifuge tubes	VWR	87003-294
15 mL tubes	VWR	21008-670
2 mL microcentrifuge tubes	VWR	87003-298
4-well plates	Fisher Scientific	FB012926
50 mL  tubes	VWR	21008-690
6-well plates	Fisher Scientific	89093-494
96-well plate for qPCR	BioRad	HSP9601
Agarose	Fisher Scientific	BP1356500
Calcion-P	Brightwell Aquatics	N/A
Calcium chloride dihydrate	Research Products International	C36200
Capillaries with filament - Borosilicate	Harvard Apparatus	30-0038
Capillaries without filament - Borosilicate	Harvard Apparatus	30-0016
Cas9 protein HiFi V3	IDT	1081060
CellTram 4r Air	Calibre Scientific	EPE-5196000013
CFX Maestro Software for qPCR	BioRad	12013758
Chiller for microscope	Linkam	T96 PE100-ZAL System
Depression slide	Fisher Scientific	S00376
Dextran	Fisher Scientific	D3329
DNA Ladder 1kb	Promega  Corporation	G5711
Electrophoresis chamber and gel caster	BioRad	1704469
Ethanol	Koptec	UN1170
Fetal Bovine Serum (FBS) - heat inactivated	Fisher Scientific	A5670801
Filtered Tips P1000	Rainin	30389272
Filtered Tips P20/P2	Rainin	30389274
Filtered Tips P200	Rainin	30389276
Galactose	Fisher Scientific	BP656500
GoTaq(R) Hot Start	Promega  Corporation	M5123
gRNA	IDT	N/A
Hemostat locking scissor clamp	Amazon	DS01KEL
HEPES	Affymetrix	16926
Incubator DigiTherm Heating/Cooling	Tritech Research	DT2-MP-47L
Instant Ocean Sea Salt	Fisher Scientific	S81800
Inverted Microscope with 20x DIC objective	Zeiss	AxioObserver
L-cysteine	Millipore Sigma	C7352
Lettuce, organic	N/A	N/A
Loading dye GelRed	Biotum	41011
Magnesion-P	Brightwell Aquatics	N/A
Magnesium chloride	Sigma	M0250
Microcentrifuge - refrigerated	Eppendorf	5425R
MICRO-ePORE pinpoint cell penetrator	World Precision Instruments	300684
Microforge	Narishige	MF-900
Microinjector FemtoJet 4i	Calibre Scientific	5252000021
Microloader tapered tip	Calibre Scientific	930001007
Micromanipulators TransferMan 4r	Calibre Scientific	5193000020
Needle puller	Sutter Instrument Company	P-97
NP-40	Fisher Scientific	28324
Nuclease free water	VWR	E476
Paraffin oil	Millipore Sigma	PX0047
PCR Strip Tubes	VWR	490003-706
Petri dishes 100 mm	VWR	25373-100
Petri dishes 35mm	VWR	25373-041
Petri dishes 60mm	VWR	25373-085
Phenol red	Millipore Sigma	P0290
Pipettes P2	Rainin	17014393
Pipettes P20	Rainin	17014392
Pipettes P200	Rainin	17014391
Pipttes P1000	Rainin	17014382
Plate Sealing Film for qPCR	BioRad	17010701
Potassium chloride	Fisher Scientific	P217
Power Supply PowerPac	BioRad	1645050
Primers	IDT	N/A
Proteinase K	Millipore Sigma	P4850
qPCR machine	BioRad	CFX96
qPCR probes	IDT	N/A
Rift Lake dKH	Brightwell Aquatics	N/A
Scissors for microdissection	Fine Science Tools	15006-09
Sequencing Service	Genewiz	N/A
Siphon Water Changer	Amazon	11278605
Snail tank lids - 18 qt	Cambro	SFC12
Snail tank lids - 8 qt and 6 qt	Cambro	SFC6
Snail tanks - 18 qt	Cambro	18SFSCW
Snail tanks - 6 qt	Cambro	6SFSCW
Snail tanks - 8 qt	Cambro	8SFSCW
Sodium chloride	Fisher Scientific	S271500
Sodium hydroxyde	Fisher Scientific	SS2551
Sodium hypochlorite	Clorox (Bleach)	N/A
Stereo microscope with fluorescent light	Zeiss	Discovery V8
Stereo microscope with transmitted light (mirror base)	Zeiss	Stemi 508
SYBR Green Supermix for qPCR	BioRad	1725270
TAE buffer	VWR	76371-722
Thermal cycler	Eppendorf	Mastercycler x40
Tips P1000	Rainin	30389215
Tips P20/P2	Rainin	30389228
Tips P200	Rainin	30389245
Tris pH 8	VWR	10128-396
Tween 20	Sigma Aldrich	P1379
Tweezers with curved end	A2Z SciLab	686797862535
Water Recirculation System - Custom-made	Iwaki America	N/A