Highly Sensitive Measurement of Glomerular Permeability in Mice with Fluorescein Isothiocyanate-polysucrose 70

This method can help answer key questions in the nephrology field about the role of glomerular permeability in kidney disease. The main advantage of this technique is that small and transient increases of glomerular permeability are detected by using a fluorescent lead labeled tracer. After anesthesia, place a urinary catheter in the female mouse.

Position the mouse in dorsal recumbency. Tighten the lower abdomen and find the urethral ostium. Obtain the plastic part of the catheter of a 22 G angiocatheter and lubricate it with xylocaine gel.

Use the left index finger to place careful tension on the lower abdomen and introduce the catheter three millimeters into the urethral ostium while paralleling the distal urethral axis. Turn the top of the angiocatheter 180 degrees by keeping the tip within the urethral ostium and maintaining the axis of the urethra. Then, introduce the catheter seven millimeter further into the mouse so that it is placed within the bladder.

Place a 1.5 milliliter brown tube over the top of the angiocatheter to collect the urine. Apply one milliliter of 0.9%sodium chloride subcutaneously to enhance urine production. Shave the neck of the mouse and place it in recumbency with the head toward the surgeon.

Hyper extend the head of the mouse with a tape. Disinfect the neck with 70%isopropanol. Using a tweezer and a pair of scissors make a five millimeter skin incision below the jawline.

Cut the skin approximately one centimeter in the direction of the sternum until the middle of the sternum is reached. Use a pair of scissors. Carefully dissect the skin to the right side of the neck.

Make a rectangular incision of the skin to the right to expose the soft tissue of the neck. Be careful not to damage the jugular vein. Use a pair of scissors and a tweezer to fix the skin flaps with two clamps.

Using the tip of the fine tweezer, carefully expose the jugular vein by blunt preparation. If necessary, remove tissue with fine scissors. Avoid injury to vein branches.

Place and close a ligature with a silk thread at the distal part of the visible jugular vein which is toward the head of the mouse. Fix the silk thread with a tape to put tension on the ligature. This ensures slight tension on the jugular vein.

Prepare a ligature around the proximal part of the jugular vein. After that, fill the catheter with the equilibration and fusion solution. Fix the catheter with a tape so that the catheter is aligning the jugular vein.

Control for bubbles to avoid air embolism. Lift the jugular vein with fine tweezers at the site of insertion. Align the tubing parallel to the jugular vein and puncture the jugular vein aiming for the lumen.

Insert for approximately two to four millimeters paralleling the axis of the jugular vein. Avoid any brisk movements. Close the ligature to fix the catheter.

Control for tightness of the ligature by carefully viewing through the microscope. Put a damp swab over the site of surgery. Place the tail cuff at the bottom of the mouse tail while the mouse lays in dorsal recumbency.

Start the measurements and repeat them ten times per time point. If blood pressure measurements do not seem correct and false data are produced, adjust the position of the tail cuff. Before the equilibration phase, change the urinary collection tube and place it on ice.

Fold a swab and put it around the small vessel catheter that is introduced into the jugular vein. Place a clamp over the swab to abolish retrograde blood flow or air embolism. Connect a 27 G needle with a one milliliter syringe filled with the FITC bolus to the end of the small vessel catheter.

Open the clamp and apply the 100 microliter FITC bolus. Close the clamp again and connect the larger catheter with the smaller catheter. Reopen the clamp and start the syringe pump with a flow rate of 0.008 milliliter per minute.

Continue the infusion for 60 minutes. In this phase, the effective drugs on glomerular perm selectivity is investigated. Change the urinary tube form time point zero minute to another 1.5 milliliter brown tube.

Put a swab around the small vessel catheter and disconnect the large catheter from the small vessel catheter. Fill the syringe with the experimental phase solution. Place it into the syringe pump and let the infusion pump run so that the large catheter is filled with the experimental phase solution.

Reconnect the catheters while avoiding air embolism and reopen the clamp. Start the syringe pump at the flow rate of 0.008 milliliter per minute. Continue running the syringe pump for 60 minutes and collect the urine within the urinary tube thereafter.

Place the urine tube on ice. In this study, fluorescence scan of native mouse urine shows a peak at 395 nanometers. FITC polysucrose 70 at a concentration of 40 micrograms per milliliter in mouse urine displays a fluorescence maximum at 525 nanometers.

Mouse urine does not disturb the fluorescence measurement of FITC polysucrose 70. At the admission of 525 nanometers increasing concentrations of FITC polysucrose 70 shows an increased fluorescence intensity. This method is also capable of detecting differences in glomerular permeability.

The white columns represent FITC polysucrose 70 levels before the start of angiotensin II stimulation in the experimental phase. After dosages for 60 minutes, the glomerular permeability increased compared with control group. angiotensin II wash out decreased glomerular permeability.

The FITC polysucrose 70 levels decreased after an additional 60 minutes of angiotensin II stimulation or 60 minutes of angiotensin II washout. The increase in glomerular permeability due to angiotensin II could be blocked by an angiotensin II receptor blocker. Blood pressure monitored by the tail cuff method did not show significant differences between the control and angiotensin II treated groups.

While attempting this procedure it's important to avoid air embolism when changing infusion solutions.