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

Summary

Here, we present a protocol to test glomerular permeability in mice using a highly sensitive, nonradioactive tracer. This method allows repetitive urine analyses with small urine volumes.

Abstract

The loss of albumin in urine (albuminuria) predicts cardiovascular outcome. Under physiological conditions, small amounts of albumin are filtered by the glomerulus and reabsorbed in the tubular system up until the absorption limit is reached. Early increases in pathological albumin filtration may, thus, be missed by analyzing albuminuria. Therefore, the use of tracers to test glomerular permselectivity appears advantageous. Fluorescently labeled tracer fluorescein isothiocyanate (FITC)-polysucrose (i.e., FITC-Ficoll), can be used to study glomerular permselectivity. FITC-polysucrose molecules are freely filtered by the glomerulus but not reabsorbed in the tubular system. In mice and rats, FITC-polysucrose has been investigated in models of glomerular permeability by using technically complex procedures (i.e., radioactive measurements, high-performance liquid chromatography [HPLC], gel filtration). We have modified and facilitated a FITC-polysucrose tracer-based protocol to test early and small increases in glomerular permeability to FITC-polysucrose 70 (size of albumin) in mice. This method allows repetitive urine analyses with small urine volumes (5 µL). This protocol contains information on how the tracer FITC-polysucrose 70 is applied intravenously and urine is collected via a simple urinary catheter. Urine is analyzed via a fluorescence plate reader and normalized to a urine concentration marker (creatinine), thereby avoiding technically complex procedures.

Introduction

Functional or structural defects within the glomerular filtration barrier increase glomerular permeability to albumin, resulting in the detection of albumin in the urine (albuminuria). Albuminuria predicts cardiovascular outcome and is an important marker for glomerular injury¹. Even low levels of albuminuria, lying within the normal range, are associated with an increased cardiovascular risk¹.

Under physiological conditions, albumin is filtered through the glomerulus and is almost completely reabsorbed in the tubular system^2,3. In mice, the detection of albumin in the urine is usually performed by an albumin enzyme-linked immunosorbent assay (ELISA) from 24 h of urine collection. If urine from a 24 h urine collection or spot urine is used, small differences in albumin concentrations may be missed due to assay sensitivity problems. Most researchers, therefore, use animal models in which albuminuria is induced by robust renal injury due to toxins, drugs, and renal surgery.

Therefore, the finding of a sensitive method to detect small and transient changes in glomerular permeability is very important to the field. Rippe et al. have presented a rat model to test glomerular permeability by applying a fluorescently labeled tracer, namely FITC-polysucrose 70 (i.e., FITC-Ficoll 70), at the size of albumin⁴. The tracer application allows the testing of short-term changes in glomerular permeability (within minutes) and is very sensitive⁴. Two studies have used the tracer method in mice^5,6. Despite its benefits, this method, unfortunately, has disadvantages: it is technically very complex, radioactive, and invasive. Further analysis of the urine is only accomplished by using gel filtration or size-exclusion HPLC^5,6.

Within this paper, we present an alternative, sensitive, nonradioactive, and fast method to measure glomerular permeability in mice using fluorescently labeled FITC-polysucrose 70. By introducing a transurethral catheter, urine collection is less invasive than bladder puncture, urethrotomy, and suprapubic catheter application, and allows urine collection at least every 30 min. Urine analysis is performed from small amounts (5µL) using a fluorescent plate reader. Tracer concentrations in the urine are normalized to creatinine concentrations in the urine using an enzymatic creatinine assay.

Therefore, this novel method offers a sensitive tool to study early glomerular injury with increased glomerular permeability.

Protocol

The investigations were conducted according to the guidelines outlined in the Guide for Care and Use of Laboratory Animals (US National Institutes of Health Publication No. 85-23, revised 1996). All animal experiments were performed in accordance with the relevant institutional approvals (state government Landesamt für Natur, Umwelt und Verbraucherschutz [LANUV] reference number 84-02.04.2012.A397).

1. Preparation of instruments, solutions, and equipment

1. Reconstitute FITC-polysucrose 70 with 0.9% sterile sodium chloride (NaCl) to a final concentration of 10 mg/mL (i.e., 100 mg in 10 mL of NaCl).
2. Dialyze FITC-polysucrose 70 solution to remove free FITC molecules overnight at 4 °C (molecular weight cut-off [MWCO] at 10,000). Use 1 L of 0.9% sterile NaCl per 10 mL of FITC-polysucrose 70 under constant stirring. Protect from light. Aliquot the dialyzed FITC-polysucrose 70 and store it at -20 °C.
3. For the FITC-polysucrose 70 bolus, add 4 µL of 10 mg/mL FITC-polysucrose 70 solution to 996 µL of 0.9% NaCl (the final concentration of FITC-polysucrose 70: 40 µg/mL).
4. For the equilibration infusion solution, add 20 µL of a 10 mg/mL FITC-polysucrose 70 solution to 9.98 mL of 0.9% sterile NaCl yielding to a final concentration of 20 µg/mL.
5. For the experimental solution, add drugs or substances to the infusion solution (e.g., for angiotensin II [Ang II] [100 ng/kg/min] for a 25 g mouse, add 3 µL of Ang II of a 1 mM solution).
6. For the surgery, prepare one shaver, two surgical clamps, one pair of surgical scissors, two tweezers, two fine tweezers, one pair of fine scissors, and swabs. Prepare two 10 cm silk threads (4-0 to 6-0) for ligation procedures.
7. For the placement of a central venous catheter, prepare a 10 mL syringe with a 21 G needle. Place the tip of the needle in a 30 cm-long catheter (with an inner diameter [ID] of 0.58 mm). Connect the 0.58 mm catheter to a 10 cm catheter (with an ID of 0.28 mm). Cut the tip of the smaller catheter oblique to create a sharp tip that is introduced into the jugular vein.
8. Prepare the anesthesia (i.e., intraperitoneal anesthesia ketamine, 100 mg/kg of body weight, and xylazine, 5 mg/kg of body weight).
9. Prepare a 22 G angiocatheter by discarding the needle and marking the catheter 1 cm from the tip. Preheat the heating pad to 37 °C.
10. Prepare a blood pressure device and change the blood-pressure-measuring membrane if necessary.

2. Preparation phase

1. Urinary catheter
   NOTE: Section 2.1 follows the protocol as described by Reis et al.⁷. Figure 1 and Supplemental Figure 1 show the placement of a urinary catheter in female mice.
   1. Anesthetize the mouse with ketamine/xylazine (see step 1.8). Use the toe-pinch test to confirm proper anesthesia. To maintain anesthesia, repeat the anesthesia (e.g., with half the dosage) after 60 min. Proper anesthesia is confirmed if toe pinch test does not result in reflex withdrawl.
      NOTE: Female FVB mice are used in this protocol.
   2. Position the mouse in dorsal recumbency on a 37 °C heating pad. Tighten the lower abdomen and find the urethral ostium (e.g., under a microscope).
   3. Use the plastic part of the catheter of a 22 G angiocatheter and lubricate it with xylocaine gel. Introduce it carefully 3 mm into the urethral ostium while paralleling the distal urethral axis (Figure 1A).
   4. Turn the top of the angiocatheter 180° by keeping the tip within the urethral ostium and maintaining the axis of the urethra (Figure 1B).
   5. Introduce the catheter 7 mm further into the mouse so that it is placed within the bladder (Figure 1C). Do not force the catheter beyond resistance. Correct the position of the catheter from the beginning if resistance is felt. Take note that if the position of the urinary catheter is correct, urine might already appear within the catheter.
   6. Place a 1.5 mL brown tube over the top of the angiocatheter to collect the urine. Apply 1 mL of 0.9% NaCl subcutaneously to enhance urine production.
2. Central venous catheter
   1. Shave the neck of the mouse and place it in recumbency with the head toward the surgeon. Hyperextend the head of the mouse with a tape.
   2. Disinfect the neck with 70% isopropanol. Make a small skin incision (5 mm) below the jawline, using a tweezer and a pair of scissors. Cut the skin approximately 1 cm in the direction of the sternum until the middle of the sternum is reached.
   3. Carefully dissect the skin on the right side of the neck, using a pair of scissors. Make a rectangular incision of the skin to the right side of the mouse to expose the soft tissue of the neck. Use a pair of scissors and a tweezer. Fix the skin flaps with two clamps.
      NOTE: The jugular vein runs along the left side of the thyroid gland or is slightly covered by the right lobe of the thyroid gland. After this step, use a microscope for surgery.
   4. Carefully expose the jugular vein by blunt preparation, using the tip of the fine tweezer. Avoid injury to vein branches.
      NOTE: It might be necessary to remove tissue with fine scissors. Be careful when using scissors to remove tissue as it increases the risk of bleeding.
   5. Place and close a ligature with a silk thread (4-0 to 6-0) at the distal part of the visible jugular vein (toward the head of the mouse). Put tension on the ligature by fixing the silk thread with a tape to ensure slight tension on the jugular vein. Prepare a ligature around the proximal part of the jugular vein.
   6. Fill the catheter with the equilibration infusion solution (see step 1.4) and fix the catheter with a tape so that the catheter is aligning the jugular vein. Control for bubbles to avoid air embolism.
   7. Lift the jugular vein with fine tweezers at the site of insertion (the insertion site is 1–2 mm proximal of the ligature). Align the tubing parallel to the jugular vein. Puncture the jugular vein, aiming for the lumen, and insert for approximately 2–4 mm, paralleling the axis of the jugular vein. Avoid any brisk movements.
   8. 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.
3. Blood pressure measurement
   1. Place the tail-cuff at the bottom of the mouse tail while the mouse lays in dorsal recumbency. Start the measurements and repeat them 10x per time point. Build a mean from the measurements.
   2. Adjust the position of the tail-cuff if blood pressure measurements do not seem correct and false data are produced.

3. Equilibration phase

1. Change the urinary collection tube before the equilibration phase starts and place it on ice.
2. Introduce a 10 mL syringe filled with FITC equilibration phase solution (see step 1.4) with a 21 G needle and the larger catheter (with an ID of 0.58 mm) and place it into the syringe pump.
3. Fold a swab and put it around the small-vessel catheter (with an ID of 0.28 mm) that is introduced into the jugular vein. Lay the catheter inside the swab. Place a clamp over the swab to abolish retrograde blood flow or air embolism. Connect a 27 G needle with a 1 mL syringe filled with the FITC bolus to the end of the small-vessel catheter.
4. Open the clamp and apply the FITC bolus (100 µL). Close the clamp again and connect the larger catheter with the smaller catheter. Start the syringe pump with 0.008 mL/min (0.480 mL/h). Continue the infusion for 60 min.

4. Experimental phase

NOTE: In this phase, the effect of drugs on glomerular permselectivity can be investigated.

1. Change the urinary tube (time point 0 min) to another 1.5 mL brown tube. Put a swab around the small-vessel catheter and close a clamp as indicated in step 3.3.
2. Disconnect the large catheter from the small-vessel catheter. Change the syringes to the experimental phase solutions (see step 1.5). Let the infusion pump run so that the large catheter is filled with the equilibration solution.
3. Reconnect the catheters while avoiding air embolism and reopen the clamp. Start the syringe pump at 0.008 mL/min.
4. Continue running the syringe pump for 60 min and collect the urine within the urinary tube thereafter (urine 60 min). Place the urine tube on ice.
5. Sacrifice the mouse by cervical dislocation during anesthesia. Prior to disposal the mouse is observed for 5 minutes to ensure it is dead.

5. Urine analysis

1. Fluorescence measurement
   1. Thaw the urinary pool and dilute it 1:10 with phosphate-buffered saline (PBS).
      NOTE: The urinary pool is a pool of urine that has been collected from healthy mice in a 12–24 h urine collection in a metabolic cage.
   2. Prepare standards for the fluorescence measurement. Take 10 brown 1.5 mL tubes and label them for blank (1:10 diluted urine pool) and the following FITC-polysucrose 70 concentrations (0.625, 1.25, 2.5, 5, 10, 20, 40, 80, and 160 µg/mL).
   3. Pipet 246 µL of diluted urine pool into a 1.5 mL brown tube and add 4 µL of the FITC-polysucrose 70 stock concentration (see step 1.1) to get a final concentration of 160 µg/mL FITC-polysucrose 70. Pipet 100 µL of the diluted urine pool into all other tubes.
   4. Dilute 1:2 by pipetting 100 µL of the 160 µg/mL FITC-polysucrose 70 tube into the 80 µg/mL tube and continue with the other concentrations. Assure proper mixture of the diluted concentrations (e.g., by vortexing the tube before continuing).
   5. Dilute the mouse urine samples (0 min, 60 min) 1:10 with diluted urinary pool.
   6. Pipette 5 µL of each standard FITC-polysucrose 70 concentration and of the mouse urine samples as triplicates in a black 384-well plate. Centrifuge the plate at 1,000 x g for 15 s to avoid bubbles.
   7. Analyze the 384-well plate in a plate reader. Perform excitation at 496 nm and measure the fluorescence at 525 nm.
2. Creatinine measurement
   1. Follow the manufacturer’s instructions. In brief, prepare creatinine standards by diluting 10 µL of the 100 mM creatinine standard solution with 990 µL of creatinine assay buffer to prepare a 1 mM standard solution.
   2. Add 0, 2, 4, 6, 8, and 10 µL of the 1 mM creatinine standard solution into a 96-well plate to generate 0, 2, 4, 6, 8, and 10 nmol/well standards, respectively. Add creatinine assay buffer to each well to bring the volume to 50 µL.
   3. Prepare the reaction mixes by adding 42–44 µL of creatinine assay buffer, 2 µL of creatinase, 2 µL of creatininase, 2 µL of creatinine enzyme mix, and 2 µL of creatinine probe. For the blank, use 44 µL of creatinine assay buffer, 2 µL of creatinase, 2 µL of creatinine enzyme mix, and 1–2 µL of creatinine probe.
   4. Add 50 µL of the appropriate reaction mix to each well in the 96-well plate and incubate it for 60 min on a horizontal shaker at 37 °C. Protect the plate from light during the incubation. Measure the absorbance at 570 nm on a plate reader.
   5. Calculate the creatinine concentrations by subtracting the blank value from all readings. The result will have the unit nanomoles/microliter. Multiply the concentration of creatinine by the molecular weight of creatinine (113.12 ng/nmol) to receive the unit nanograms/microliter.

6. Data analysis

1. Calculate the FITC-polysucrose 70 urine concentration from the standard curve.
2. Reference the FITC-polysucrose 70 urine concentrations to creatinine concentrations in the representative urine sample.
3. Reference the urine 60 min samples to urine 0 min samples of controls and treated mice.

Results

As depicted in Figure 2, the method to test glomerular permeability in mice is built up in three phases. The first phase is called the preparation phase, in which a urinary catheter and a central venous catheter are placed. The second phase is called the equilibration phase, starting with an intravenous bolus injection of FITC-polysucrose 70 and followed by the continuous infusion of FITC-polysucrose 70 for 60 min. The last phase is called the experimental ph...

Discussion

The presented method enables the investigator to test glomerular permeability in mice in a very sensitive manner using a tracer. With this method, short-term increases in glomerular permeability can be diagnosed using only small amounts of urine. The most critical steps for successfully mastering this technique are 1) developing manual expertise in mouse surgery, especially in the cannulation of a central vein, 2) placing the urinary catheter without harming the mucosa, and 3) manual expertise in handling 384-well plates...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Motic SMZ168 BL	Motic	SMZ168BL	microscope for mouse surgery
KL1500LCD	Pulch and Lorenz microscopy	150500	light for mouse surgery
Microfederschere	Braun, Aesculap	FD100R	fine scissors
Durotip Feine Scheren	Braun, Aesculap	BC210R	for neck cut
Anatomische Pinzette	Braun, Aesculap	BD215R	for surgery
Präparierklemme	Aesculap	BJ008R	for surgery
Seraflex	Serag Wiessner	IC108000	silk thread
Ketamine 10%	Medistar		anesthesia
Rompun (Xylazin) 2%	Bayer		anesthesia
Fine Bore Polythene Tubing ID 0.28mm OD 0.61mm	Portex	800/100/100	Catheter
Fine Bore Polythene Tubing ID 0.58mm OD 0.96mm	Portex	800/100/200	Catheter
Harvard apparatus 11 Plus	Harvard Apparatus	70-2209	syringe pump
BD Insyte Autoguard	BD	381823	urinary catheter
Multimode Detector DTX 880	Beckman Coulter		plate reader
384 well microtiterplate	Nunc	262260	384 well platte
Creatinine Assay Kit	Sigma-Aldrich	MAK080	to measure creatinine concentration
96 well plate	Nunc	260836	for creatinine assay
FITC-labeled polysuccrose 70	TBD Consultancy	FP70	FITC-ficoll
Angiotensin II	Sigma-Aldrich	A9525	used to test glomerular permeability
BP-98A	Softron		for blood pressure measurement
OTS 40.3040	Medite	01-4005-00	heating plate for mouse surgery
Instillagel 6mL	Farco-Pharma GmbH		for urinary catheter
Exacta	Aesculap	GT415	shaver