Unraveling the Role of Discrete Areas of the Rat Brain in the Regulation of Ovulation through Reversible Inactivation by Tetrodotoxin Microinjections

Procedures involving animals were approved by the Ethics Committee of Facultad de Estudios Superiores Zaragoza, UNAM. This institution operates in strict accordance with the Mexican rules for animal handling, Official Norm: NOM-062-ZOO-1999, which agrees with international guidelines.

1. Construction of bilateral cannulas

1. Extract the stainless-steel shaft from the hub of two 23 G hypodermic needles using pressure tweezers and then remove any remaining glue using a scalpel blade.
2. Draw a line 15 mm apart from the blunt end of the shafts with a fine permanent marker. Use cutting tweezers to remove the beveled ends.
3. Hold the 15 mm segments with fine hemostats and press them perpendicularly with a cut-off disc attached to a rotatory tool until 14 mm segments of tubing are obtained. This step is done in order to eliminate the occluded portion of the shafts, creating open and blunt ends. Finally, insert a 30 G needle through the segments to eliminate any internal obstruction.
4. Determine the distance between the structures of interest in the left and right hemispheres of the brain with the aid of a brain atlas²⁷. Use moldering clay to attach the two 14 mm segments to a microscope slide and ensure that both are at the same horizontal level. Observe through an ocular micrometer (10x) and adjust the segments with fine tweezers until the desired distance is obtained.
5. Mix solder paste with 10% hydrochloric acid in a 2:1 proportion and add a drop of the mixture 2 mm below the blunt end of the segments. Solder both segments with a single point using a soldering iron and 0.5 mm-diameter solder wire. Ensure that the solder does not obstruct the lumen of the segments.
6. Create a support to attach the cannula to the stereotaxic holder by cutting a 10 mm segment of resilient 0.3 mm stainless steel wire. Use moldering clay to place 2 mm of the wire in contact with the previous solder point and the rest laying above the blunt end of the cannulas. Solder the wire to the cannulas.
7. Clean the surface of the cannulas with 70% ethanol to remove the excess of the solder paste. Flush the interior of the cannulas with sterile water to remove metallic particles. Repeat this process until no particles can be detected under the microscope at a 10x magnification.

2. Construction of obturators and caps

1. To build the obturators cut two 16 mm segments of round stainless-steel soft wire (0.35 mm of diameter). Hold a bilateral cannula with hemostats, perpendicular to a lab bench, and insert one of these segments into each cannula until they reach the bench. Bend the remnant at a 90° angle.
2. For the caps cut two 2 mm segments of silicone tubing (inner diameter=0.76 mm) and apply a drop of silicon glue at one of the ends of each segment. Do not let the glue to enter to the tubing. Let dry for at least 24 hours.

3. Construction of microinjectors

1. Repeat step 1.1, using 30 G hypodermic needles instead to create the shafts.
2. Draw a line 18.5 mm apart from the blunt end of the shafts and remove the remnant of the beveled ends with cutting tweezers.
3. Repeat step 1.1 with a single 23 G needle to build adaptors by cutting two 6 mm-long segments starting from the blunt end.
4. Eliminate the occluded portions of the 6 mm segments by pressing them perpendicularly against the cut-off disc until two 4 mm adaptors are obtained. Eliminate any internal obstruction.
5. Insert the beveled end of the 30 G segments into the adaptors. Look through a stereomicroscope to ensure that the end of both, segments and adaptors, are at the same level. Apply cyanoacrylate glue to the distal joint using a cotton swab and let dry for 15 minutes.
6. Soak two Teflon tubing connectors in 70% ethanol for 5 minutes and then attach them to the microinjectors through the adaptors. Wait until the diameter of the connectors shrinks for at least 24 hours and then sterilize the microinjector with a low temperature method to avoid damage of the connectors (ethylene oxide sterilization is recommended).

4. Animal maintenance and vaginal smears

1. Use cyclic adult female hooded rats (Rattus norvegicus, CIIZ-V strain) weighing between 230 and 260 grams. House the rats in groups of four in standard polypropylene cages in a room with a 14:10 light–dark photoperiod. Set the temperature and humidity at 22 ± 2 °C and at 40%, respectively. Provide food and water ad libitum.
2. Take vaginal smears every day between 10:00 and 12:00 h.
   1. Sterilize a modified bacteriological loop with an inner diameter of 1 mm using an alcohol lamp and cool it with sterile water. Hold the rat with a secure grip and introduce 5 mm of the bacteriological loop into the vagina, touching its internal walls. Remove the bacteriological loop. If successful, a cloudy drop will be seen at the tip. Place this drop on a microscope slide.
   2. Repeat this process for each rat sterilizing and cooling the loop between each animal.
   3. After the drops dry off, stain with hematoxylin-eosin and observe the samples under a microscope at 10x.
   4. Determine the proportion of leukocytes, epithelial nucleated cells and keratinized cells on each smear and classify it according to the criteria of estrous cycle stages reported in Figure 1, which agrees with previous literature^28,29.

5. Stereotaxic implantation of the cannulas

NOTE: Perform the implantation of the cannulas following a regular aseptic stereotaxic surgery and abiding to institutional norms.

1. Before surgery
   1. Sterilize surgical instruments, cannulas, surgical screws, obturators, gauze and wooden cotton swabs 24 h before surgery using a steam autoclave. Sterilize the tips of metallic instruments between consecutive surgeries by cleaning them with 10% hydrogen peroxide followed by water and then place them in a hot bead sterilizer.
   2. Prepare the working areas and the stereotaxic instrument before removing the animal from the homeroom. Prepare the animal for surgery in an area that is located as far as possible from the surgery table to avoid contamination during the procedure.
   3. Clean the preparation and surgery areas with 70% ethanol followed by an application of a 10% chloride solution for ten minutes. Clean the base, frame and manipulators of the stereotaxic instrument with 70% ethanol and sterilize the tips of the ear bars using a hot bead sterilizer and air-cool them before surgery.
   4. Perform the surgery wearing a dedicated lab coat, face mask, head bonnet, surgical sleeves, shoe covers and surgical gloves. Ask the assistant to prepare the animals for surgery and check their general state during the procedure.
   5. Attach the cannula to the stereotaxic holder. Ask the assistant to bring the first animal to be operated into the room. Select rats in diestrus for surgery, since observations from our lab indicate that these animals recover proper estrous cycles quicker than rats operated in other stages, probably because the response to stress changes along with the estrous cycle.
   6. Weigh the animal, and induce anesthesia with 4% isoflurane in 100% oxygen inside an induction chamber for rats connected to an isoflurane vaporizer. To avoid stressing the animals, do not prefill the chamber. Confirm a surgical plane of anesthesia by checking pupil dilation and the loss of pain as measured by the tail and ear pinch tests after you observe the loss of the righting reflex.
      1. Anesthetize the rat by an intraperitoneal injection of a mixture of ketamine (100 mg/kg) and xylazine (10 mg/kg) if an isoflurane vaporizer is not available.
   7. Remove the rat from the induction chamber and use a nose cone in the preparation table to maintain the effects of anesthesia with 2.5% isoflurane in 100% oxygen. Trim the hair of the scalp with clippers and remove loose hair with a lint roller. Inject a preoperative subcutaneous dose of 2 mg/kg of Meloxicam and 5 mg/kg of Enrofloxacin as a non-steroidal anti-inflammatory/analgesic and antibiotic, respectively.
      1. Apply hypromellose artificial tears to each eye to avoid desiccation during the surgery.
   8. Mount the animal in the stereotaxic instrument above a warming pad using non-rupture ear bars and an anesthesia mask. Adjust the nose clamp to ensure that the head of the animal is flat. Perform a surgical scrub in the shaved area alternating between povidone-iodine with soap and 70% ethanol one time and then using povidone-iodine and 70% ethanol two times. Insert a rectal thermometer to record the temperature of the animal during the procedure and then cover it with a sterile surgical field.
2. During surgery
   1. Use a scalpel to make a 2 cm incision in the skin and muscle in the middle of the shaved area. Remove the periosteum from the skull using a cotton swab coated with 2% hydrogen peroxide to reveal bregma or lambda, the landmarks that will be used to calculate the target coordinates. Dry the skull with air for a better visualization of the landmark.
   2. Use the manipulators of the stereotaxic instrument to move the cannula to a position directly above the landmark of choice. Register the anterior-posterior and the medial-lateral coordinates from the stereotaxic instrument and use them to calculate the coordinates of the target area according to the brain atlas²⁷.
   3. Set the calculated coordinates in the stereotaxic instrument and place the tip of the cannula on the surface of the skull. Register the dorsal-ventral coordinate and use it to calculate the depth to the target area.
   4. Remove the arm of the manipulator by unscrewing it from the frame. Use a dental burr attached to a rotatory tool at a speed of 15,000 rpm to make a mark in the place where the craniotomy will be performed. Then use the burr to make three holes, forming an equilateral triangle around the mark. These holes must not pierce the skull. Insert the surgical screws into the holes, ensuring that they are tightly placed.
   5. Make the craniotomy wide enough to accommodate the distance between the target areas in each hemisphere. It should be as small in diameter as possible but big enough to allow the lowering of the cannula without touching the borders of the skull, since this will modify the trajectory. Perform the craniotomy gradually, applying the lowest pressure possible. The rodent skull is just a few millimeters thick and it is pivotal to preserve the integrity of the tissues below.
   6. Once the dura mater is visible, use a sterile 21 G needle with the tip curved at a 90° angle to remove splinters of bone and make an anterior-posterior cut in the meninges. Use Wirtshafter and colleagues³⁰ technique to avoid damage to the superior sagittal sinus if the target area is located close to the midline.
      1. Place the holder with the cannula in the frame and use the dorsal-ventral manipulator to reach the ventral coordinate. Check that the needle does not touch the borders while descending and increase the diameter of the craniotomy if needed.
         NOTE: It is important that the tip of the guide cannulas targets a region ranging from 0.5 to 2.0 mm above the region of interest. This is due to the inflammatory reaction of the brain in response to the introduction of foreign bodies and to the destruction of the tissue. This is especially critical if the region of interest is small and the working distance must be calculated empirically in advance.
   7. Apply dental cement to attach the cannula to the skull and let it dry. Ensure that the dental cement does not cover the blunt end of the cannula since this will irreversibly obstruct it. Wait until the cement solidifies completely.
   8. Insert the obturators to avoid further obstructions and to ensure patency during the study. Put on the silicone cap to avoid contamination.
3. After surgery
   1. Replace lost fluids by an intraperitoneal injection of 1.0 mL of sterile saline solution at a physiological temperature. Place the animal in a recovery cage with thermal support until it recovers from anesthesia. Check the temperature and hearth/respiratory rate of the animal periodically. Isoflurane effects last for a few minutes after interruption of the gas flux while ketamine/xylazine effects lasts for 40-50 min.
   2. Provide post-operative injections of the antibiotic, analgesic and anti-inflammatory drugs (at the same doses and routes stated before) for 48-72 h and closely inspect the animals on a daily basis for the rest of the experiment. Report any sign of pain, stress or loss of weight to vet services or to a trained member of the lab. Do not perform any other manipulation to the rats during this period.
   3. Start taking vaginal smears after the post-operative treatment and continue doing it until the animal shows three consecutive estrous cycles of four days.
      NOTE: Chronic jugular catheters can be surgically implanted, allowing the researcher to take serial blood samples to analyze the secretion of hormones such as estradiol, progesterone, testosterone, luteinizing hormone, follicle-stimulating hormone and corticosterone. We recommend placing such a catheter once the animal recovers from the brain surgery, since this will improve survival rate while also avoiding clogging of the catheter that may result from an extended recovery time.

6. Tetrodotoxin handling and preparation of solutions

CAUTION: TTX is one of the most toxic substances known. It acts on skeletal muscles and nervous tissue. Intoxication by contact is unlikely but any open wound is a potential pathway for inoculation into the body. The main concerns when working with TTX are the puncture of the skin with sharp instruments that have been in contact with the toxin and the generation of aerosols that may reach the mouth, eyes and mucous membranes. Depending on the dose, TTX may be lethal if inhaled, swallowed or inoculated by the skin. It will cause strong irritation of the eyes. The LD50 has been tested in mice by oral and intravenous administration and it is 334 µg/kg and 7.3 µg/kg, respectively. There is no antitoxin available at this time.

NOTE: Inactivating substance is a 1.0% NaOCl with or without 0.25 N NaOH, or a 10% bleach solution. Complete inactivation occurs after 30 min exposure. TTX cannot be completely inactivated by any chloride derivate at a concentration below 10%, by autoclaving nor by dry heat sterilization at a temperature lower than 500 °F. All the procedures involving TTX must be performed by two knowledgeable individuals wearing inner and outer pairs of nitrile gloves, dedicated lab coat, safety glasses, disposable face mask, closed shoes and full-length pants.

1. Preparation of the stock solution
   1. Place a pad that is soaked with the inactivating substance in the fume hood to prevent contamination in case of a spill. Be sure that the fume hood is working properly before starting. Prepare a receptacle with the inactivating solution to dispose of pipette tips.
   2. Resuspend sterile TTX citrate lyophilized powder according to manufacturer instructions by an extremely careful and slow titration with sterile artificial cerebrospinal fluid, rinsing down the walls of the vial in the process. Avoid the formation of foam and aerosol. Follow an aseptic technique to avoid contamination of the TTX solution since it will be injected into the brain of live animals. Use a syringe with a needle instead of a micropipette if your TTX vials have an outer membrane to prevent aerosol formation.
   3. Aliquot the stock solution into sterile micro tubes. Make aliquots as small as possible since freezing/thawing cycles may alter the stability of the molecules. Do not fill more than half of the tubes because water increases in volume when frozen, which can lead to opening of the lid and contamination of the tube and other samples stored in the container.
   4. Decontaminate the exterior of the tubes and the surfaces where TTX was used with the inactivating substance. Store the tubes at -20 °C in a vial box inside a secondary container.
2. Diluting the stock solution to a working concentration
   1. Prepare freshly diluted solutions the day they will be used since diluted solutions are less stable. Place a pad that is soaked with the inactivating substance in the fume hood and a micro tube-rack on top of it. Prepare a receptacle with the inactivating solution to dispose of pipette tips.
   2. Pipette the stock solution at the bottom of a sterile microtube and then add the calculated amount of artificial cerebrospinal fluid to achieve a concentration of 10 ng/µL. As described for the re-suspending of the TTX powder, perform an extremely careful and slow titration, rinsing down the walls of the vial and avoiding the formation of foam and aerosol.
   3. If samples that have been on the freezer will be used for microinjection, they must be allowed to equilibrate at room temperature for at least one hour prior to the experiment.

7. Microinjection of TTX or vehicle solutions into freely moving rats

1. Configure the microinjection pump with the infusion rate and total time of injection according to the experiment. Calculate these parameters in advance considering the volume occupied by the target structure and the amount of solution to be injected. For this experiment inject 200 nL of solution at a rate of 50 nL per minute for a total infusion time of 4 minutes.
2. Fill two 10 μL Hamilton syringes with sterile distilled water, insert a piece of sterile Teflon tubing into the tubing connector of each microinjector, ensuring that the length of the tubing does not constrain the movement of the rat (tubing can be sterilized using ethylene oxide).
3. Place a pad that is soaked with the inactivating substance and a 2 cm x 2 cm square of paraffin film above it. Pipette a sufficient amount of TTX to fill the needle of the injector and 1 cm of the tubing above the film. Absorb the TTX drop by gently retracting the plunger. Mount the syringes in the microinjection pump and use its controls to manipulate the plunger until a drop of TTX can be seen at the tip of each microinjector. Discard the drops into the pad soaked with inactivating substance.
4. Select the rats that will be microinjected. Use only rats that showed at least three consecutive cycles after the surgery. Consider their stage of the cycle and the time of the day. Both the time and the stage depends on the specific hypothesis that you will test, for this experiment 14:00 hours of proestrus selected since the nervous preovulatory signals governing phasic GnRH secretion occur at this moment. Transport them to the room where the injection will take place inside their own housing cages to avoid distress.
5. Hold the rat with a firm grip, remove the cap and obturators from the cannulas, insert the microinjectors into the guide cannulas and return the animal to the cage.
6. Turn on the pump and wait until the microinjection finishes. Observe the animal during this period in order to detect a possible detachment of the microinjectors and to avoid the twisting of the Teflon tubes.
7. Leave the microinjectors in place for two additional minutes to avoid reflux of the solution. Remove them, clean the surface of the implant with iodopovidone antiseptic solution, avoiding its flow inside the guide cannulas. Insert sterile obturators, put on the cap and return the animal to the colony room. Observe the animal periodically to detect any possible side effects of the drug.
8. Turn on the pump with the same parameters as during microinjection and check for a correct flow to ensure that patency of the system was retained during the procedure.
9. Press the plunger to expel the TTX/vehicle out of the microinjector until the air bubble reaches the tip of the needle. Recollect the TTX in its microtube. Fill the syringe with distilled water and use it to clean the interior of the tubes. Discard the water into the inactivating substance and repeat this process at least three times. Clean the outside of syringes, tubes and microinjectors with a cloth soaked in the inactivating substance.

8. Euthanasia and tissue processing

1. On the day of the predicted or vaginally confirmed estrus, inject the rat with an intraperitoneal overdose of sodium pentobarbital (75 mg/kg). Check for loss of consciousness and inject 200 nL of a 0.5% methylene blue solution through the guide cannulas as described in steps 7.1 to 7.9. Check for pain signs using the toe or tail pinch test and, if none are detected, decapitate the rat using a rodent guillotine.
2. Use scissors to open the abdominal cavity and find the ovaries. Use fine iris scissors to dissect each ovary, cutting at the utero-tubal junction with the aid of a magnifying glass. Avoid damaging the oviduct because this will result in the loss of oocytes.
3. Put the ovaries under a stereomicroscope and use a razor blade to remove all the fat tissue without damaging the organ. Remove the oviduct from each ovary by cutting with the razorblade as far as possible from the ampulla region. Mount each oviduct on a different slide and cover with a drop of water. Store the ovaries in a vial containing Bouin´s solution.
4. Gently dry the oviducts with an absorbent paper towel and search for the ampulla under the stereomicroscope. With the non-dominant hand, hold the oviduct in place by pinching far from the ampulla with a 23 G needle, then use another 23 G needle in the dominant hand to make a 1 mm incision in the middle region of the ampulla. The cumulus-oocyte complexes will protrude from the incision as a drop of viscous fluid (Figure 2D). Use the needle in the dominant hand to gently and slowly pull the drop far from the oviduct. Press the remnant of the ampulla to ensure that no oocytes are left inside. Extract the oocytes from the oviducts as fast as possible as desiccation of the oviduct will irreversibly trap the oocytes inside.
5. Remove the oviduct from the slide and wait until the drop of fluid containing the oocytes dries. Stain the sample with hematoxylin-eosin. Use mounting medium to coverslip. Observe under the microscope (10x) to determine the number of oocytes shed by each ovary.
   NOTE: Sacrifice by cardiac perfusion is not performed in this protocol since the oocytes would be trapped in the oviducts and hence histological methods would be necessary to assess ovulation. If the user must perfuse to analyze other tissues, the ovaries can be removed first and the descending arteries clamped with thick hemostats below the liver to avoid leaking of fixative.
6. Remove the skin of the head and submerge the skull in a glass container with a 10% formalin solution. Allow fixation of the head for at least ten days before removing the cannula to avoid distortion of the tissue. To remove the cannula, use scissors to detach all the muscles in the region of the skull that surrounds it. Cut between the nasal and frontal bones with bone trimmers and then remove the occipital bone. Insert the tip of the trimmers into the foramen magnum and start cutting through the sagittal crests reaching the remnant of the nasal bone.
7. The isolated region of the top of the skull must contain the frontal and parietal bones. Remove the implanted cannula attached to the top of the skull by slowly pulling it up perpendicularly to the head. Cut any attached portion of the meninges with fine iris scissors to avoid deformation of the brain.
8. Make an anterior-posterior cut on the meninges and put them aside to remove the brain from the base of the skull. Insert blunt tweezers below the olfactory bulbs and gently pull up the brain until the optic nerves are visible. Cut the nerves with fine iris scissors and pull out the brain. Preserve the brain in fixative solution.
9. Obtain 50 µm thick sections of the brain using a vibratome, mount them in poly-L-lysine coated slides (0.1% in distilled water) and stain with the Nissl technique. Observe the slides under the microscope (10x) to determine the final position of the cannulas (Figure 2A-2C) and the spread of the dye with the aid of a rat brain atlas²⁷.