Monitoring Blood Glucose in Mouse Offspring After Intracytoplasmic Sperm Injection

The protocol of ICSI-ET described below follows the guidelines and has been approved by the Animal Ethical Review of the Shanghai Institute of Planned Parenthood research. Safety Procedures: Always wear appropriate personal protective equipment (PPE) when handling chemicals or biological materials. Use of Hoods: Perform all procedures involving volatile chemicals or aerosol generation within a certified fume hood or biosafety cabinet. Use female mice (6-8 weeks old B6D2F1 strain) for the superovulation procedure.

1. ICSI in mice

1. Superovulation of mice
   1. Administer an intraperitoneal injection of pregnant mare serum gonadotropin (PMSG) at a dose of 7.5 IU per mouse at around 20:00 on the first day.
   2. Administer an intraperitoneal injection of human chorionic gonadotropin (HCG) at a dose of 7.5 IU per mouse at around 20:00 (48 hours after PMSG injection) on the third day.
2. Prepare HTF medium into 1 mL aliquots and leave them uncovered for overnight equilibration at 37 °C in a 5% CO₂ incubator on the same day of HCG injection in female mice.
3. Preparation of ICSI fertilization droplets
   1. On the same day of HCG injection in female mice, divide a 50 x 9 mm cell culture dish lid into upper and lower halves. Use the upper half to create 4-6 droplets of 5 µL each, using 10% polyvinylpyrrolidone (PVP) for sperm placement. Use the lower half to create 8-10 droplets of 5 µL each, using M2 medium for the ICSI procedure.
   2. Cover the PVP and M2 medium droplets with approximately 3 mL of mineral oil (Figure 1).
4. Preparation of culture dishes
   1. On the same day of HCG injection in female mice, dispense 12 drops of KSOM medium (MR-020P-D) of 20 µL evenly in a 35 x 10 mm cell culture dish. Cover the drops with approximately 3 mL of mineral oil.
5. Sperm harvesting:
   1. Euthanize male mice by cervical dislocation at approximately 9:00 am on the fourth day (13 h after HCG injection).
   2. Isolate the epididymal cauda and thoroughly clean it to eliminate any adipose tissue and blood remnants by gently blotting with sterile gauze and rinsing in prewarmed HTF medium.
   3. Cut open the cauda epididymis to allow the sperm to freely swim out into the HTF medium. Gently squeeze the cauda using forceps to release the sperm, collect them in preequilibrated HTF tubes, and keep them uncovered in a 37 °C, 5% CO₂ incubator for sperm capacitation for further use.
6. Oocyte retrieval
   1. Euthanize female mice stimulated for oocyte retrieval by cervical dislocation at around 9:30 am on the fourth day. Carefully make an incision along the midline of the abdominal skin and muscle layer to expose the abdominal cavity.
   2. Use fine forceps to gently isolate tissue from both sides of the oviducts, especially the swollen ampulla regions, and transfer it into M2 medium in a sterilized culture dish. Repeat this procedure until the oviducts from all the designated mice are collected.
   3. Under a dissecting microscope, use a 1 mL syringe needle to puncture the transparent section of the swollen ampulla of the oviduct gently, facilitating the release of cumulus-oocyte complexes (COCs) into the medium.
   4. Using the same syringe, carefully pick and transfer the COCs into a new dish containing 100 µL of fresh M2 medium. Repeat this step until all COCs are successfully transferred from the original oviduct segments.
   5. Wipe away blood and tissue fluids, and under a dissecting microscope, tear open the ampulla of the oviduct using a 25 G needle, resulting in the release of numerous COCs.
   6. Remove the oviduct and tissue fragments, transfer the COCs into droplets of M2 medium containing hyaluronidase (HY), and prewarm them to 37 °C.
   7. After incubation for approximately 5 min, carry out gentle pipetting to facilitate the separation of cumulus cells and oocytes. Transfer the metaphase II (MII) oocytes from the HY-containing solution to fresh M2 medium; then, perform the sequential transfer into multiple droplets of M2 medium.
   8. After 2-3 rounds of washes, transfer the oocytes to droplets of KSOM culture medium and keep them in a 37 °C, 5% CO₂ incubator for further use.
7. Around 10:00 am on the fourth day, sonicate 100 µL of the capacitated sperm (see step 1.5.2.) for 5 s to separate the heads from the tails at a frequency of 46 kHz, with a power setting of 50 watts.
   NOTE: When sonicating sperm to truncate the tail, be cautious to control the time, which should not exceed 5 s. A few seconds after sonication, sperm can be observed under an inverted microscope; stop sonicating when approximately half of the sperm show truncated tails.
8. ICSI procedure
   1. Use a commercially available micromanipulation needle with a flat tip and an inner diameter of 6 µm, set at a 25° angle, for the injection. Fill the needle with the operating fluid with the filled portion measuring a length of 0.5 cm inside the needle. Ensure that the level of the fluid reaches ~0.5 cm.
      NOTE: Maintain room temperature at 19 °C during injection to avoid high temperatures that may reduce the survival rate of fertilized oocytes. The precision in ensuring the fluid level reaches 0.5 cm is crucial for maintaining the integrity of the injection process and ensuring the viability of the oocytes.
   2. Install the injection needle on the right arm of the micromanipulator, securely fixed by tightening the holding cap. Install a commercially available flat-tipped micromanipulation holding pipette designed for mouse oocytes on the left arm of the micromanipulator, and position the injection needle and the holding pipette in the microscope's field of view.
   3. Add 1 µL of sonicated sperm to a droplet of PVP solution using a pipette.
   4. Place 20 MII oocytes in a droplet of M2 medium.
   5. Use the injection needle to aspirate 10-20 sperm heads in PVP solution. Then, move the injection needle into the droplet containing the oocytes.
   6. Directly determine the position of the metaphase spindle within the oocyte under a polarization microscope to ensure that the spindle apparatus is not on the side of injection, with the polar body typically located at either the 12 o'clock or 6 o'clock position relative to the oocyte's orientation.
   7. Keep the oocyte in contact with the bottom of the culture dish. Adjust the focal plane so that the zona pellucida of the oocyte and the injection needle are in the same plane.
   8. Upon contact of the injection needle with the zona pellucida, apply a slight negative pressure to the injection needle, drawing in a portion of the zona pellucida.
   9. Simultaneously activate piezo pulses with an intensity setting of 5 and a frequency of 1, effectively creating a perforation in the zona pellucida and allowing the injection needle to pass through.
   10. Advance the injection needle into the perivitelline space, pushing the sperm head to the tip of the needle. Insert the injection needle into the oocyte until it reaches the opposite side of the zona pellucida.
   11. Use piezo pulses set to an intensity of 1 and a frequency of 1 to create a small pore in the plasma membrane, ensuring the sperm head is injected into the ooplasm with minimal entry of PVP and optimal preservation of the oocyte's integrity.
   12. Repeat step 1.8.11 until all 20 oocytes are injected, followed by a 15 min incubation at 19 °C.
   13. Transfer the surviving oocytes to KSOM medium and place them in the incubator. Observe 2-cell, 4-cell, morula, and blastocyst formation rates.
       NOTE: Timing for intracytoplasmic single sperm injection (ICSI) into the oocyte should be controlled, aiming to complete the procedure within 14-18 h after the injection of HCG.

2. Uterine transplantation of mouse blastocysts

1. Preparation of vasectomized male mice
   NOTE: Use ICR male mice aged 6-8 weeks for the study.
   1. Induce anesthesia by intraperitoneal injection of 1.25% 2,2,2-tribromoethanol at a dosage of 0.2 mL/10 g of body weight. The absence of any pain response indicates successful anesthetization.
      NOTE: While the current protocol does not include the application of vet ointment to the eyes of anesthetized animals, it is recommended as a best practice to prevent eye dryness and ensure the health and comfort of the animals during procedures. Throughout the surgical and recovery process, all surgical instruments were sterilized before use, and the surgical area was prepared using antiseptic solutions to ensure a clean operating environment.
   2. To safeguard against potential stress or injury, house the mice that have undergone surgery individually during the initial recovery phase.
   3. Administer buprenorphine at a dosage of 0.05 mg/kg of body weight to ensure optimal pain relief.
      NOTE: To ensure the well-being of the mice, closely monitor the mice during the recovery period and do not leave them unattended until they regain sufficient consciousness to maintain sternal recumbency.
   4. Prepare the surgical site at the lower abdominal region, superior to the testes. Remove the fur and disinfect the area with alcohol. Using sterile scissors, make a 1 cm longitudinal incision along the linea alba. Gently remove any blood with sterile, dust-free paper and proceed to bluntly dissect the abdominal muscles along the linea alba and access the abdominal cavity.
   5. Using curved forceps, gently grasp and expose the left testis and its associated fat pad. Identify the vas deferens and carefully isolate it using pointed forceps, creating a 10 mm segment for manipulation.
   6. Sterilize the curved forceps by holding them in the outer flame of an alcohol lamp for several seconds. Gently grasp the vas deferens, causing it to become flattened and whitened due to cauterization. Create a second cauterization point 3-5 mm distal to the first. Using sterile surgical scissors, excise the segment of the vas deferens between the two cauterization points.
   7. Carefully inspect the cauterized ends of the vas deferens, ensuring that a short, flattened, whitened segment remains. Using curved forceps, gently return the testis and its associated fat pad to the abdominal cavity.
   8. Repeat the procedure described in step 2.1.6 on the contralateral (right) side.
   9. Close the surgical site using sutures, typically placing one stitch in the muscle layer and three stitches in the skin. Ensure proper alignment of the surgical incision and disinfect the area with alcohol spray. Transfer the animal to a heating pad for observation, monitoring for 10-15 min until the animal regains consciousness. Once recovered, return the animal to its housing cage.
      NOTE: Following vasectomy surgery, mice require a 14-day recovery period, followed by a 1-week trial mating experiment.
   10. During this week, continuously mate the vasectomized males with females and observe the pregnancy status of all females presenting with copulatory plugs. Successful mating is indicated by the presence of a yellowish-white, blocky vaginal plug at the vaginal orifice of the female mouse.
       1. Pair the mice between 4:00 PM and 6:00 PM on the first day and check the vaginal plugs between 7:00 AM and 9:00 AM on the second day. Consider females with vaginal plugs to be in a pseudopregnant state at 0.5 days and use them for subsequent embryo transfer experiments. Return females without vaginal plugs to the recipient group.
          NOTE: These females without vaginal plugs can continue to be used for mating to prepare pseudopregnant females. Ensure that all plugged females do not become pregnant; only then can the vasectomized males be used for formal experiments.
       2. While vasectomized males can mate daily, allow for 1 day of rest between mating sessions.
          NOTE: Based on our experience, vasectomized males can maintain a high copulatory plug rate for over a year. However, vasectomized males that fail to produce copulatory plugs in mating females for 4-6 consecutive attempts should be removed from the study.
2. Preparation of pseudopregnant females
   1. Generate pseudopregnant female mice by mating ICR female mice (aged 7-10 weeks, weighing over 25 g) with vasectomized male mice (3-6 months old) at a 1:1 ratio.
      NOTE: The vasectomized males were used to stimulate the uterine cervix of the female mice, resulting in the activation of the corpora lutea.
3. Preparation of the pipette for transferring blastocysts
   1. Take the embryos to be transplanted out from the incubator and transfer them into a droplet of M2 medium at a temperature of 37 °C.
   2. Use a transfer pipette to aspirate a small volume of liquid, draw a small amount of air, and aspirate 6-10 embryos. Finally, aspirate another small volume of air to seal the pipette.
      NOTE: The length of the fluid in the transfer pipette should not exceed 0.5 cm.
4. Uterine transplantation (blastocysts)
   1. Weigh the pseudopregnant female mice (2.5 days post mating) and anesthetize them by intraperitoneal injection of 1.25% 2,2,2-tribromoethanol at a dose of 0.2 mL/10 g of body weight. Gently press the ear with forceps.
   2. After administering anesthesia to the mice and ensuring asepsis through alcohol disinfection, make a midline longitudinal incision on the ventral abdomen. Taking advantage of the loose subcutaneous tissue, shift the incision to the left or right side as required for the transplant. Use small scissors to perform blunt dissection of the abdominal wall muscle layer at ~0.5 cm lateral to the lumbar spine to access the peritoneal cavity.
   3. Using atraumatic forceps, carefully exteriorize the ovaries, oviducts, and uterus.
   4. Using the tip of a syringe needle, make a small, upward-angled incision on the uterine wall below the utero-tubal junction, avoiding blood vessels. This incision provides access to the uterine cavity; gently insert the oviduct 2-3 mm through the small hole.
   5. Carefully expel the small air bubble at the end of the transfer pipette while cautiously watching the transfer pipette during the slow injection of the embryos. Once the central portion of the culture medium is confirmed to be expelled, halt the procedure and gently remove the transfer pipette.
   6. Reposition the ovaries, oviducts, and uterus back into the abdominal cavity. Suture the muscle layer and skin separately. Place the mice on a heating pad until they recover from anesthesia. To ensure the well-being of the mice, closely monitor the mice during the recovery period and do not leave them unattended until they regain sufficient consciousness to maintain sternal recumbency.
5. Observation of pregnancy
   1. Following the surgery, observe the mice for 18-21 days to monitor the number of offspring born. In case of dystocia (difficult birth), perform a cesarean section. Allow the newborn mice to be fostered by lactating dams of the same age group.

3. Random blood glucose, fasting blood glucose, and glucose tolerance test

1. Tracking random blood glucose levels during mouse growth
   1. Since the birth of the mice, monitor the random blood glucose level every 4 weeks at 9 am after a good night of feeding and fun.
      1. Puncture the mouse tail tip to obtain tail vein blood.
      2. The first drop of blood has a higher sugar measurement value due to the presence of the tissue fluid mixed with it. Therefore, wipe the first drop of blood using clean absorbent paper gently.
      3. Measure the blood glucose from the second drop (~4 µL) from the tail vein with a handheld glucometer.
2. Tracking changes in fasting blood glucose and glucose tolerance during mice development
   NOTE: Considering that the first blood sampling time point is 15 min, all mice need to be injected within 15 min. Since each mouse injection took 20-30 s, to ensure that the injections are completed within this timeframe, the maximum number of mice that can be used in each test cannot exceed 30.
   1. Perform the fasting blood glucose and glucose tolerance tests every 4 weeks after birth in overnight-fasted mice.
      1. On the first day, prepare fresh irradiated shavings bedding, clean cages, lids, and water bottles. Transfer the mice into a fresh cage without food. Add do not feed tags to the cages.
      2. Prepare sufficient 10% glucose injection (sterile and pyrogen-free) and 1 mL syringes.
      3. Ensure that the mice in each group fast for 16 h, from 5:00 pm on day 1 to 9:00 am on day 2, but allow free access to water. Measure the fasting blood glucose levels as described in steps 3.1.1.1-3.1.1.3 using a glucose meter.
   2. Weigh the mice and mark their tails to facilitate rapid identification. Prepare glucose syringes with 10 µL/g of mouse weight.
   3. At 30 s intervals, inject 10 µL/g glucose into the intraperitoneal cavity of the mouse.
      NOTE: Special attention needs to be paid to injection techniques here to avoid inaccurate liquid injection volume. The key points of operation are as follows:
      1. Immobilize the mouse and restrain the tail with one hand. Keep the head slightly lower than the buttocks so that the organs in the abdominal cavity will shift towards the head, leaving enough injection space.
      2. Position the needle between the midline and the hip bone, insert it through the skin at a 45-degree angle, then gently guide the needle deeper into the abdominal cavity while keeping it aligned parallel to the skin surface for ~0.5 cm.
         NOTE: Pay attention to any resistance during injection to determine whether the internal organs are accidentally punctured.
      3. Eject the syringe into the abdominal cavity.
      4. Record the time when the injection started and use a timer to control the injection rate. Set the timer to 90 s for three mice to avoid losing time in handling the timer.
   4. Take the blood glucose measurement from the tail vein at the desired intervals (0 prior to glucose administration, 15, 30, 60, 90, and 120 min after the glucose loading).
      NOTE: Subsequent blood glucose testing should also take 20-30 s for each mouse.
   5. Return the mice to their home cages equipped with food and water after the experiment.