Remote Neuronal Activation Coupled with Automated Blood Sampling to Induce and Measure Circulating Luteinizing Hormone in Mice

All animal procedures are performed in agreement with the National Research Council Guide for the Care and Use of Laboratory Animals¹⁸ as well as federal, state, and local laws. Adult female mice (3-6 months old) were used for this protocol demonstration, including four C57BL/6J females and four Kiss1-Cre;ChR2-eYFP (Kiss1-eYFP) females. Mice were held under a 12:12 light:dark cycle, temperature-controlled at 22 °C, and fed ad libitum on a low-phytoestrogen diet. Procedures and protocols were approved by the University of Michigan Institutional Animal Care and Use Committee (IACUC, Animal Protocols: PRO00010420 and PRO00010138).

1. Stereotaxic delivery of AAVs to a specific cell population

1. Preparation for surgery
   1. Sterilize all tools. Prepare multiple packages of surgical tools to ensure that each package is used in no more than five animals. Prepare sterile gloves, a minimum of one pair for each animal.
   2. Pull glass micropipettes for injections using the following settings (see the Table of Materials) for long thin micropipettes that inject slowly and steadily and do not easily clog: Heat 1: 915, Heat 2: 630, Pull: 630. Optimize these settings for each puller.
   3. Perform surgeries in a designated surgical space. Disinfect surgical surfaces with 70% ethanol.
   4. Use sterile drapes to maintain the sterility of the surgical field. Wear a clean lab coat or disposable gown and a mask.
   5. Prepare inhalation anesthesia system. Open oxygen supply and regulate flow rate to 0.8 L/min.
2. Surgery
   1. Place the mouse in an anesthesia box and open the isoflurane to 2.5%. Reduce isoflurane flow to 2% after initial induction; maintain flow throughout the procedure. For alternative methods of anesthesia, refer to animal protocol and local ethical committee guidelines.
   2. Inject preemptive analgesic (carprofen 5 mg/kg s.c.) following animal use committee recommendation and local regulations.
   3. Shave the mouse head using clippers.
   4. Install the animal on a stereotaxic table, place ear bars, and make sure that the head is correctly fixed and stable. Fix the mouse's mouth to the mouthpiece, making sure to place the tongue to the side outside of the mouth to avoid suffocation.
   5. Verify that the animal is deeply anesthetized with a toe pinch before starting the surgery and monitor the mouse's breath and color throughout the procedure.
   6. Place elevation support under the mouse to maintain the body and head leveled in a horizontal position. Keep the animal warm with a warm pad covered in paper. Apply eye ointment to both eyes to avoid drying.
   7. Keep the surgical area as clean as possible. Wear sterile gloves. Disinfect the mouse's head with iodine and alcohol prior to opening the skin. With a scalpel, cut the skin on the head along the midline, approximately from behind the eyes until behind the lambda suture. Keep the skull exposed and clean it with a cotton swab embedded in sterile 0.9% NaCl.
   8. Find the rostral rhinal vein (RRV) and mark it with a sterile pencil. Use the stereoscope for the rest of the procedure.
      NOTE: We obtain better results using the RRV as an anteroposterior reference, but it is standard to use bregma as a reference.
   9. Using a sterile needle as a reference, make sure that the brain orientation is correct before proceeding to stereotaxic measurements. Ensure also that the heights of the skull surface at the RRV and lambda, as well as the lateral inclination, are the same (± 0.02 mm).
      NOTE: The lateral inclination becomes more relevant for injection into more lateral brain structures.
   10. Load a sterile glass pipette with the viral solution to be injected. Bring it to the RRV reference for reference 0 anteroposterior (AP). Advance along the sagittal suture to the AP coordinate of choice. Mark this position with a sterile pencil, lift the needle, and proceed to craniotomy.
   11. Carefully drill a small circle around the marked position to avoid breaking the superior sagittal sinus. Position the drill in a tilted rather than a perpendicular position to reduce pressure while drilling. Remove the piece of the skull with small forceps.
   12. Once the vasculature is exposed, use the middle of the superior sagittal sinus to use as the mediolateral (ML) reference (point 0); this is more precise than using the sagittal suture. Move to the ML position of choice. Lower the pipette to touch the dura-mater as the dorsoventral (DV) reference (point 0). Slightly break the dura-mater and descend the pipette to the DV position of choice.
   13. Inject the desired amount of AAV (50-200 nL) and leave the cannula in place for 3 min to allow adequate liquid dispersion. Remove the pipette carefully out from the brain.
   14. Release the mouse from the ear bars. Close the skin using surgical clips or any method of choice. Put the animal in a separate warmed cage for recovery. Monitor recovery, reactivity, and activity after the mouse is awake. When fully recovered, move the mouse back to its original cage.
3. Wait for a minimum of 3-4 weeks for viral expression before proceeding with the second part of the procedure.

2. Jugular vein and carotid artery canulation

1. Preparation for surgery
   1. Perform surgeries in a designated surgical space. Disinfect surgical surfaces with 70% ethanol before initiating the procedures. Prepare multiple packages of surgical tools to ensure that each package is used in no more than five animals.
   2. Autoclave all surgical instruments. Then, clean with sterile water or saline and disinfect the tools with a hot bead sterilizer for at least 15 s (according to the manufacturer's instructions) between surgeries.
   3. Use sterile drapes to maintain the sterility of the surgical field. Wear a clean lab coat or disposable gown and a mask.
   4. Prepare micro-catheters for cannulations in the carotid artery and jugular vein. Construct the artery catheter by joining a small segment of microrenathane tubing (stretched from 0.025 inches outer diameter [OD] x 0.012 inches inner diameter [ID]) with silastic tubing (0.025 inches OD x 0.012 inches ID). Construct the venous catheter only using silastic tubing (0.025 inches OD x 0.012 inches ID). Soak all catheters in 70% ethanol overnight prior to surgery.
   5. Bevel the cut tips of both catheters at 45° to preestimated lengths based on the animal's body weight and length.
2. Surgical procedures
   1. Anesthetize the animals under 2% isoflurane with a tabletop low-flow isoflurane system to precisely control the anesthesia stage and plane during surgery.
   2. Inject the preemptive dose of analgesic (carprofen 5 mg/kg s.c.) and apply ophthalmic ointment to prevent desiccation and corneal injuries.
   3. Shave the ventral and back areas of the neck and clean the skin with three iodine scrubs alternated with 70% ethanol. Cut a vertical skin incision (12 mm) between the shoulder blades and cover it with surgical gauze for later use. Place the animal in a supine position with the head towards the surgeon.
   4. Use a stereo dissecting scope for most parts of the procedures described below.
   5. Make a small vertical incision (~10 mm) on the right side of the neck, superior to the clavicle to expose the right carotid artery and jugular vein. Cut the skin with scissors and perform a blunt dissection to separate subcutaneous tissues using non-toothed micro forceps with fine tips, exposing the right external jugular vein and the right common carotid artery.
   6. Tie the distal end of the jugular vein to stop blood flow and cut a small hole in the collapsed vein using micro forceps and scissors. Insert the venous catheter bevel down and proximally using a pair of micro forceps to advance it through the superior vena cava to reach the level of the right atrium. The inserted length is ~10-12 mm for a 30 g lean mouse. Tie the catheter to fix with the vessel using 7-0 silk sutures.
      NOTE: With the catheter in the right place, blood can be easily withdrawn; otherwise, the catheter may be misplaced into the lateral thoracic vein and needs to be reinserted.
   7. Carefully dissect connective tissues to expose the right common carotid artery located in a triangle area surrounded by the sternohyoid muscle, sternomastoid muscle, and digastric muscle. Use a wire retractor to separate these muscles if needed. Tie the artery at the level of the bifurcation of internal and external carotid arteries using 7-0 silk sutures. Preplace two untied suture loops proximally.
   8. Temporarily stop the blood flow by pulling a preplaced suture loop and cut or punch a small hole on the vessel wall using micro scissors or a 27 G needle. Insert the arterial catheter bevel down and proximally to a preestimated length reaching the aorta arch but without touching the aortic valve. The inserted length is ~9-10 mm for a 30 g lean mouse. Tie the catheter to fix with the vessel using the two preplaced sutures.
   9. Tunnel all catheters subcutaneously and exteriorize them at the back of the neck via the precut incision and join them to the venous or arterial ports of a silicone-coated tubing connector made from 25 G needle tubing: modified MASA¹⁹ using two 25 G needle tubing and two 4.0 cm of PE-20 tubing, sleeved with 2.0 cm of 0.062 inch ID silastic tubing attached with a small metal wire-made ring. The total lumen volume of each catheter including the connector is 6-8 µL (Figure 1A).
   10. Close the ventral incision and fix the connector subcutaneously upon the closure of the back skin with sutures. Fill both catheters with heparinized saline (200 U/mL) and plug them tightly at the end with stainless steel surgical wires.
   11. Mice recover from isoflurane anesthesia within minutes and fully recover from surgery within 5 days. Place the animals in the automated blood collection housing chamber 24 h after surgery and connect them to the system by joining the tether hook of the system to the metal ring attached to the tubing connector implanted on the back of the neck. Connect the arterial and venous catheters to the injection and sampling lines, respectively, 24 h before the sampling begins (Figure 1B) . The total length of the sampling line is 55 cm in length or 40 µL by lumen volume.
       NOTE: The setups of injection and sampling lines, the programming of the collection method, and the cage balance mechanisms of the automated blood sampler system have been described in detail^20,21. The system keeps the catheter open by automatically delivering 10 µL of heparinized saline every 20 min.
   12. Set the preinjection sampling, injection, and postinjection sampling time and frequency via the computer program of the system. See step 3.1 below for the current settings.
   13. The system allows the animal to move freely without tangling the sampling and/or infusion lines by sensing the mouse movement while rotating the housing chamber in an opposite direction to the mouse movement²¹ (Figure 1).
   14. Refill the infusion or injection line with the compound and reconnect to the venous catheter at least 2 h before the infusion or injection begins.
       ​NOTE: Surgery complications might occur, and animals need to be monitored closely for recovery and on the days following the surgery. Refer to the animal protocol for correct monitoring, termination requirements, and procedures.

3. Automated blood collection and intravenous injection

1. Automated blood collection
   1. To follow this protocol, use a sampling volume of 20.0 µL and set an interval (i.e., 7.0 min) between each sampling to have a sampling frequency of every 10.0 min per sample. The maximum rate is to sample continuously at 3.0 min/sample; the minimum possible sampling volume is 5.0 µL. An equal amount of saline is automatically given back by the system to replace the sampled blood and maintain body fluid balance.
   2. Set the total sampling time as 30 (t = -30 – 0) min before and 30 (t = 0 – 30) min after the intravenous injection of CNO (0.5 mg/kg, Figure 2).
   3. Each collected blood sample (20.0 µL) is automatically diluted in 50 µL of saline (containing heparin at 10 U/mL) and stored individually in a microtube held in a refrigerated sample carousel by the system.
2. Automated intravenous injection
   1. Disconnect the venous line from the venous catheter to refill CNO (individually calculated dose by volume of ~50-60 µL at 0.5 mg/kg) at least 2 h before blood sampling begins.
   2. Manually withdraw the solution (slightly more than the calculated volume) from the line retrogradely with the injection syringe, leaving a small air bubble between the solution and the existing saline in the line.
   3. Reconnect the injection line to the venous catheter and set the injection rate at 500 µL/min and the injection starting time at 2 min after t = 0 sampling finishes. The total injection time for each animal is 5-6 s.
      NOTE: If required, the compound can also be delivered by a manual intraperitoneal injection (IP), but this will require disturbing the animal during or before the blood collection protocol. We also show this alternative in the results.

4. Animal perfusion and brain collection (OPTIONAL)

NOTE: This procedure is to be followed only if the brain is required to analyze the brain site of neuronal activation or downstream responses.

1. Disconnect the mouse from the blood collection system at the end of the collection protocol. Two hours after the intravenous injection, proceed to perfuse the animal with 10% neutral buffered formalin (NBF), using the preferred method or as described elsewhere²².
2. Dissect the brain and conserve it in 20% sucrose in 10%-NBF for 3 h to continue fixation.
3. After 3 h, transfer the fixed brain to 20% sucrose in PBS and conserve at 4 °C until ready for sectioning.
   NOTE: Avoid overfixation as this will mask the cFOS antigens. In the 20% sucrose solution, the brain should sink to the bottom of the container.
4. Make sections using a freezing microtome or cryostat and use the preferred method to look at neuronal activation (e.g., cFOS immunohistochemistry is shown).
   ​NOTE: The antibodies used for the current results are described in the Table of Materials.

5. Sample processing and analysis

1. Remove blood samples from the automatic sampler immediately after the end of the experiment and put them on ice.
2. Spin the samples at 14,000 × g for 30 s.
3. Collect the plasma. Store it at -80 °C until analysis.
4. Assay the blood samples by LH ELISA as described before^10,23.