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

Luteinizing hormone (LH) pulsatility is a hallmark of reproductive function. We describe a protocol for remote activation of specific neuronal populations linked to serial automated blood collection. This technique allows timed hormonal modulation, multiplexing, and minimizing manipulation effects on LH levels in conscious freely moving, and undisturbed animals.

Our goal is to understand the role of a specific neuronal populations in the regulation of the pulsatile secretion of luteinizing hormone. For this, we require precise control of neuron allocation, identity and activity in combination with the sequential collection of blood samples without disturbing or stressing the animals. Current method for collecting sequential blood samples in conscious mice are limited to tail cuts, require a long training period and are susceptible to human handling and environmental stresses.

Automated infusion and blood sampling can be conducted without presence of investigators, which limits stress and allows for the capture of hormonal changes in conscious, free moving, and undisturbed mice. This technique is a asset to neuroscience research as it provides a robust support system. The Michigan Mouse Metabolic Phenotyping Center offers automated infusion and the broad sampling experiments as a core service.

To begin, inject a preemptive analgesic into an anesthetized mouse. Using clippers, shave the head of the mouse. Install the animal on a stereotaxic table and correctly position ear bars.

Attach the mouse's mouth to the mouse piece and carefully position the tongue outside to prevent suffocation. After checking the depth of anesthesia, position elevation support beneath the animal to ensure the body and head remain leveled horizontally. And use a warm pad with paper to maintain the animal's temperature.

Apply ointment to both eyes to prevent dryness. Then using iodine and alcohol, disinfect the mouse's head. Use a scalpel to make an incision along the head's midline with the cut stretching from behind the eyes to post the lambda suture.

Expose the skull and swab it with a sterile cotton tip saturated in 0.9 sodium chloride solution. Under a stereo microscope, identify the rostral rhinal vein or RRV and use a sterile pencil to mark it. Using a sterile needle as a reference, confirm the orientation of the brain.

Ensure no change in the DV and ML coordinates. Next position a sterile glass pipette containing the virus solution to the RRV reference for zero anteroposterior reference. Advance the pipette along the sagittal suture until reaching the chosen anteroposterior coordinate.

Mark this spot with a sterile pencil and lift the needle. Carefully drill a small circle around the marked position to avoid breaking the superior sagittal sinus and remove the piece of the skull with small forceps. Then using the superior sagittal sinus median as the media lateral reference, move the pipette to the position and lower it until it touches the dura mater as the dorsoventral reference.

Create a slight break in the dura mater and lower the pipette to the preferred dorsoventral position. Inject the specified quantity of adeno-associated virus or AAV vectors into the brain and remove the pipette. Free the mouse from the ear bars.

Then close the skin using surgical clips. Transition the animal into a separate warmed cage to recover. Keep an eye on the mouse's recovery.

Once it regains consciousness, return it to its original enclosure when fully recovered. To begin, inject the mouse with AAV associated vectors into the mouse's brain. After three to four weeks of virus injection, shave the ventral and back areas of the neck of an anesthetized mouse.

Clean the shaved areas using three iodine scrubs in rotation with 70%ethanol solution. Make a ventral incision in the skin between the shoulder blades. Afterward, place surgical gauze over the incision and position the animal in a supine state with its head facing toward the surgeon.

Then make a small vertical incision on the right side of the neck above the clavicle to expose the right carotid artery and jugular vein. After separating the subcutaneous tissue, expose the right external jugular vein and place the suture under the vein. Tie off the distal end of the jugular vein to halt the blood flow.

Using micro forceps and scissors, make a small incision in the collapsed vein. Now insert the venous catheter with the bevel facing downward and move it proximally towards the superior vena cava until it reaches the right atrium. Using a 7-0 silk suture, secure the catheter to the vessel.

Dissect the connective tissues to expose the right common carotid artery. Approximately pre placed two 7-0 silk sutures suture loops that remain unfastened at the level where the internal and external carotid arteries divide. To stop the blood flow temporarily pull a pre-placed suture loop.

Then using micro scissors or a 27 gauge needle, create a small opening on the vessel wall. Insert the arterial catheter proximally while releasing the suture loop and advance the catheter to reach the aorta arch without touching the aortic valve. Use the two pre-placed to fasten the catheter to the vessel.

Tunnel all catheters subcutaneously, exteriorize them at the back of the neck via the precut incision and close the ventral incision. Join the catheters to the venous or arterial ports of a silicone coated tubing connector made from 25 gauge needle tubing. After closing the ventral incision, secure the connector subcutaneously when the back skin is sealed with sutures.

Using stainless steel surgical wires, fill the catheters with heparinized saline and tightly plug the ends of the catheters. To attach the animal on an automated blood collection system 24 hours post-surgery, connect the tether hook to the metal ring on the back of the neck and connect the arterial line. Once the arterial catheter's connected to the injection line, attach the venous catheters to the sampling lines.

Set the injection time and dose at 0.5 milligrams per kilogram at 500 microliters per minute. Set the blood sampling time and frequency including the sample dilution with saline. The system will automatically replenish an equal amount of saline to replace the sampled blood.

Perform automated pre injection. Next, perform an automated intravenous injection of the drug clozapine N-oxide at an injection rate of 500 microliters per minute. Then perform post-injection blood sampling.

Luteinizing hormone patterns in adult kiss1-EYFP females that received a unilateral stereotaxic injection of AAV-hM3Dq mCherry in the arcuate nucleus are presented. Diestrous luteinizing hormone levels are generally low, but variations are usually observed because of its pulsatile release. A sharp rise in luteinizing hormone levels occurs in response to clozapine N-oxide injection.

Most mCherry neurons co-localized with Kiss1-EYFP demonstrated that the viral and neuronal activation is specific to the targeted population. A luteinizing hormone pulsatile pattern of release in diestrous wild type mice followed by the response to an IP injection of kisspeptin-10 is shown. Clear luteinizing hormone pulses typical for a female in diestrous were observed showing low basal luteinizing hormone levels.

An immediate and robust increase in luteinizing hormone was detected in response to kisspeptin administration.