Intravital 현미경 검사 법 및 대비 향상 된 초음파 마우스 Hindlimb의 인슐린 유도 된 혈관과 근육 관류 결합

The overall goal of this combined contrast-enhanced ultrasonography and intravital microscopy technique is to simultaneously visualize muscle-resistant arteries and to study skeletal muscle micro-circulation perfusion in vivo. This method helps answer key questions in the fields of Type 2 diabetes and obesity on regulation of insulin sensitivity and organ blood flow. On the day of the procedure, after disinfecting the operating area and equipment with an alcohol-based solution, equip a 10 centimeter piece of PE20 tubing with a 27-gauge needle, and attach the tube to a four-way connector.

Insert the needle into the tail vein of a 20 to 25 gram fasted in anesthetized CD57 BL6 male mouse. And use tissue adhesive gel to fix the needle to the skin tissue. Place the mouse in the supine position and use thermostable tape to fix the feet, exposing the upper-thigh area.

Using a slight exo-rotation of the hip joint, face the hind limb paws up with the knees at a 40 to 60 degree angle to standardize the stretch of the muscle at the adductor compartment of the thigh. Use a depilatory cream to bilaterally remove the hair at the groin and upper thigh areas, collecting all of the loose hair with a moist cotton swab. Then place the mouse under a stereo microscope at a 10 to 16X magnification, and make a two centimeter incision parallel to the inguinal ligament, just lateral to the abdominal curvature.

It is important to practice the placement of the skin incision, which should be cut small enough that the perifemoral does not leak away, but large enough that the vasculature can be viewed. Using a bulldog clamp, apply traction to the distal side of the incision, then dissect the adipose tissue away from the abdominal wall, gently separating the fat pad from the wall in a distal direction instead of dissecting directly through the pad to avoid bleeding. Identify the femoral artery, following the vessel down to the epigastric and gracilis arteries and the transparent deep fascia covering the muscles and the vessels.

Using sharp forceps, pull the fascia upwards, and cut the tissue with micro-scissors. Then cover the exposed muscle with a 200 microliter drop of paraffin oil to prevent the tissue from drying out and use the bulldog clamp to arrange this skin folds of the incision to make a small cavity for holding the oil bathing the vessels. Now adjust the mouse so that the gracilis artery is vertical on the computer screen at a 16X magnification and place a light source at a minimum of 20 centimeters from the hind limb to reduce the heat conduction from the light.

Applied pre-warmed ultrasound transduction gel to the upper contralateral hind limb and place the ultrasound probe perpendicular to the long axis of the femur bone. Carefully adjust the angle and direction of the ultrasound probe to obtain a cross-sectional view of the adductor muscle group, taking care to keep the position of the probe stable to maintain the same imaging plane for the baseline and hyperinsulinemic measurements. The placement of the ultrasound probe should be standardized so that the same portion of the upper hind limb will be studied in all the of the mice with the probe in the same position for the basal and hyperinsulinemic measurements.

After properly placing the ultrasound probe and having successfully prepared the gracilis artery for measurements, let the mice stabilize for 30 minutes. After a 30-minute stabilization period, proceed by making the baseline measurements. Measure and save the baseline diameter of the gracilis artery.

As the microbubbles can only be viewed in the contrast mode of the ultrasound machine, set the appropriate parameters on the imaging system for microbubble assessment. Level the location of the focal zones to the center of the region of interest, and save a five-second clip for calculation of the background signal. The address the microbubbles to a 2.5 times 10 to the 9 bubbles per milliliter concentration, and manually shake the vial containing the microbubbles to obtain a uniform suspension.

Begin infusing the microbubbles via the tail-vein cannula at a five microliter per minute rate, keeping the infusion tube on a vibrating vortex to maintain a uniform suspension of microbubbles. After five minutes of continuous infusion, use the microbubble destruct function to generate time intensity curves of the bubbles at five and 10 minutes at the start of microbubble infusion. Assess the blood-glucose from the tail vein every five minutes.

After obtaining the baseline data, start the hyperinsulinemic-euglycemic clamp using the tail cannula to administer 200 milliunits per kilogram of insulin bolus. Maintain a continuous 7.5 milliunit per kilogram per minute insulin infusion for 60 minutes. Serial blood-glucose level monitoring should be performed in blood drops obtained from the tail tip as shown earlier.

Use a variable infusion of 20%DE glucose to maintain euglycemia. Using the imaging software, measure the diameter of the gracilis artery at the desired time points after the start of the hyperinsulinemic euglycemic clamp. After 25 and/or 55 minutes, begin the second hyperinsulinemic contrast-enhance ultrasonography measurement, to monitor the microvascular blood volume at 30 and/or 60 minutes respectively as just demonstrated using the anesthesia port to infuse the microbubbles.

As demonstrated by these data, the local application of paraffin oil on the adductor muscle compartment stabilizes the vessel without changing the average baseline diameter of the arteries, and helps reduce the variation between the animals tested. Insulin-infusion consistently increases the gracilis artery diameter compared to the diameter changes induced by saline infusion, with an insulin-induced vasodilation evident after 10 minutes, and an approximately 95%maximum dilatory capacity after 30 minutes. Using contrast-enhanced ultrasonography, insulin also consistently increases the muscle microvascular blood volume compared to saline-infusion.

The signal intensity in the femoral vessels corresponds linearly with the concentration of microbubbles in the circulation, with adjustments in the femoral vessel signal, theoretically correcting for differences in the concentration of microbubbles used. Once mastered, this technique can be completed in two hours, if it is performed properly. While attempting this procedure, it is important to remember to use freshly prepared reagents and microbubbles, and to let the mouse stabilize for 30 minutes with continuous anesthesia before making the experimental measurements.

Following this procedure, new drugs can be tested to answer additional questions, such as, what is the effect of a new vasoactive component on the vasculature. After its development, this technique has paved the way for scientists in the field of vascular biology to evaluate the effects of new treatments and change of interest in norco mice. After watching this video, you should have a good understanding of how to simultaneously study the diameter of muscle-resistant arteries and to assess the perfusion of the mouse hind limb in vivo, using intravital microscopy and contrast-enhanced ultrasonography.