Using 2-Photon Microscopy to Quantify the Effects of Chronic Unilateral Ureteral Obstruction on Glomerular Processes

All experiments followed the Guide for the Care and Use of Laboratory Animals and were approved by the Animal Care and Use Committee at the Indiana University School of Medicine.

1. Preparing the Munich Wistar Frömter or SD rat for UUO surgery

1. Anesthetize the rat using isofluorane (5% induction, 1.5-2.5% maintenance), then shave, wash, and disinfect the surgical area several times in a circular motion with both an iodine-based or chlorhexidine-based scrub and alcohol. Apply long-acting/slow-release analgesic for pain management as per the institutional IACUC guidelines.
2. Make an incision along the midline using a scalpel; locate the left kidney and free it from the surrounding peritoneal organs.
3. Carefully locate the renal pedicle, which comprises the renal artery, renal vein, and ureter. Separate the ureter from the other structures, taking precautions not to damage the delicate structure.
4. Using fine forceps, carefully loop a 3-0 suture around the ureter and tie it off, being careful not to tear it. Repeat this procedure a few millimeters on either side of the first knot to tie a second knot and assure complete obstruction.
5. Once the procedure is completed, carefully close the successive muscle layers. Prior to closing the final layer, add 2 mL of warm, sterile 0.9% saline into the abdomen before closing it completely. Close the outer skin with surgical staples.
6. Apply long-acting/slow-release analgesic for pain management and observe recovery closely as per the institutional IACUC guidelines. Monitor periodically thereafter and prepare for imaging at the end of the fifth week.

2. Synthesis of Texas Red rat serum albumin (TR-RSA)

1. Weigh out 100 mg of Rat Serum Albumin and dissolve it in 6.67 mL of 0.1 M sodium bicarbonate buffer at pH 8.4 in a 50 mL conical tube.
2. To a vial of 5 mg of Texas Red-X-succinimidyl ester, add 100 µL of Dimethyl Formamide (DMF, high quality) and vortex until all the dye is dissolved.
3. Place the rat serum albumin solution on a vortexer at a low/medium setting, so the solution volume is spinning well below the top of the open tube.
4. Add the dissolved dye while the tube is being vortexed.
5. Take the 50 mL conical tube, wrap it in foil, place the tube on any rocker or roller, and agitate slowly for 1 h at room temperature (RT).
6. In a 5 L bucket of 0.9% NaCl with a stir bar under gentle stirring, wet the membranes of a suitable 50 kDa molecular weight cut-off dialyzer (a membrane with clips, enclosed membrane tubes, or dialysis cassettes are all suitable).
7. Load the TR-RSA solution in the membrane system and attach it to the flotation attachments typically included with the system. Place the 5 L container with the 0.9% saline solution/TR-RSA overnight at 4 °C (in a cold room) with gentle agitation on a stir plate. Change the dialysis solution at least three times over the next 36 h.
8. Swelling of the membrane will increase the volume of the now clear TR-RSA solution. Divide the original 100 mg by the volume to obtain an approximate concentration: the dye:protein ratio will be 1:1. Aliquot into suitable volumes and lyophilize for long-term storage.

3. Preparation for 2-photon intravital imaging on an inverted microscope

1. Remove the cover of a 50 mm coverslip bottom dish (with 40 mm coverslip diameter) and place 8 pieces of autoclave tape on the inner bottom alongside the edge. Make a pyramid-shaped, empty window, using 4 pieces per side to allow the exteriorized kidney to fit snuggly in this space while maintaining contact peripherally with the autoclave tape, thereby helping to minimize motion. Adjust the spacing according to the size of the rat to assure the best contact with the kidney.
2. Place 1 thermal pad on each side of the 50 mm coverslip bottom dish. Ensure the warming pad covers the stage.
3. Use a 40x water immersion objective at 0.75x zoom and 1.5x zoom to generate 30x and 60x images, respectively, allowing for lower- and higher-magnification images. If necessary, add water to the objective using a 1 mL syringe with a long piece of PE-200 tubing that can reach the top of the objective at a downward angle to prevent wicking of the water droplet down the tubing.
4. Use 2% laser transmissivity, with blue, green, and red detectors set to predetermined levels to assure consistency in the images between studies. Set the excitation wavelength to 800 nm on the referenced laser(see the Table of Materials), which will efficiently excite all the fluorophores used in this study.
   1. Use external (non-descanned) detectors to collect blue emissions using a photomultiplier tube (PMT) (420-490 nm, gain 950).
   2. Use a Hyd detector to capture green emissions (500-550 nm, gain 100).
   3. Use a Hyd detector to capture red emissions (590-660 nm, gain 200).
   4. Adjust the offset in the PMT (blue emissions) so that only a few pixels in the blank areas of tissue have a value of zero.
      NOTE: The HyD detectors for the green and red emissions have automatic offset adjustment; only the gain can be set.
   5. Set the bit-depth to 12-bit to give images a 4,096-intensity scale between black and white.
      NOTE: It is necessary to set the lower limits of the detectors (offset in PMT) to not exclude these values to ensure collection of the low-intensity emissions within Bowman's space. If the sensitivity setting is too low, the visual warning markers will indicate this; these values are given an intensity value of zero.
5. Dilute approximately 6 mg of Texas-Red-X-Rat Serum Albumin to a total volume of 1 mL, load the solution in a 1 mL syringe, and place on the indwelling i.v catheter in step 4.1 after infusion of Hoechst 33342 in step 9.1.

4. Surgical preparation for intravital 2-photon imaging

1. Place the pre-anesthetized rat with an indwelling venous access line (femoral or jugular) on its side with the shaved left flank facing up flat and straight on the table. Ensure that the front paws touch each other, as should the rear paws.
2. Gently palpate the left flank just below the ribs to feel for the kidney to determine the natural position in the abdomen. If needed, draw a line using a permanent marker along the shaved area, bisecting the kidney center in a nose-to-tail orientation.
3. Using a pair of toothed forceps, grasp the skin and lift it upward to facilitate pinching of the permanent marker line with a pair of hemostats to crush the underlying vasculature and prevent bleeding when making the incision with a pair of surgical scissors. Repeat this for the thin outer muscle layer to minimize bleeding.
4. To make the final incision of the thin inner abdominal muscle layer, repalpate the kidney to estimate the size and position. Carefully lift the inner muscle layer with a pair of forceps and crush a line that bisects the skin above the kidney with the hemostats that is approximately 1/3^rd the estimated size of the kidney.
5. Maintaining the grip on the muscle layer with the forceps, make the final incision.
   NOTE: It is better to make a smaller incision and expand as needed than to make it too big, which will require partial closing with a suture.
6. Gently grip the kidney by the surrounding fat. Using both hands with forceps in each hand, work to grip the fat at the lower pole of the kidney by using a hand-over-hand technique to grip and hold the kidney fat, working downward.
7. Having a firm grip on the fat at the lower pole of the kidney with one hand, gently pull the fat and, if needed, very gently squeeze the kidney through the incision. If the kidney does not pass through easily, widen the incision.

5. Positioning the rat for imaging

1. Carefully place the exposed kidney against the edge of the dish, with a slight rotation so the abdominal side of the kidney is contacting the coverslip and the dorsal side is facing away from the edge.
2. To further minimize motion, take two sterile 2 x 2 gauze pads, moisten them with saline, and pack them against the dorsal side of the kidney, reinforcing the contact of the abdominal side of the kidney to the edge.
3. Look through the microscope eyepiece under epifluorescence illumination using a dual-pass Rhodamine/FITC cube. If motion is detected, make minor adjustments to the position and carefully adjust the gauze, making sure it does not push under the kidney. To further reduce motion, roll the rat over slightly, so the thorax is further away from the dish.

6. Image acquisition for quantitative analysis

1. Scan the surface of the kidney using epifluorescence illumination (step 5.3) and mark the glomeruli position(s) using the software associated with the motorized stage controller (a feature of modern systems).
2. For each color channel under 2-photon illumination, take a shallow 3D volume of the upper portion of each marked glomerulus, which will serve as the background images. Use a pseudocolor palette in the display option of the imaging software to better visualize the faint intensities of the background fluorescence of the glomerular capillary loops.
3. Using a superficial blood vessel as the focal point, slowly infuse the fluorescent albumin, allowing time to observe the rise and fall of fluorescence due to systemic distribution. Infuse enough TR-RSA to achieve an intensity in the peritubular vasculature and capillary loops that is just below saturation.
   NOTE: There is typically a 5 s delay between the infusion of material and its appearance in the bloodstream when renal perfusion is normal.
4. Wait approximately 10 min before acquiring 3D volumes (1 µm intervals) for all marked and imaged glomeruli from step 6.2.
   NOTE: Simonsen's Munich Wistar rats have fewer surface glomeruli. However, as the Frömter strain of MW rats has a greater number of surface glomeruli, up to 10 glomeruli can often be imaged.
5. Euthanize the rat via an overdose of isoflurane at the end of the study. Perform a dual pneumothoracotamy to ensure euthanasia.

7. Calculating glomerular permeability

1. Using the image viewing software associated with the microscope system, export the images to 12-bit raw images for processing and analysis.
2. Load the background 3D volumes and the raw 3D volume containing the circulating fluorescent albumin. Locate the focal plane in the 3D volume with the brightest superficial capillary loop in the glomeruli with enough space to the edge of the surrounding Bowman's capsule.
3. Using visual landmarks, locate the same focal plane found in the background volume. Select a region in the capillary loop and one within the Bowman's space noting the average intensity values of each one. Use these intensity values as background values.
4. Outline a region (at least 20 x 20 pixels in area) within the Bowman's space in the albumin-containing image and note the intensity reading (select an area that is not adjacent to a capillary loop or the Bowman's capsule to assure the cleanest measurement of Bowman's space intensities). Move the drawn region over two other regions to take an average value for the average intensity within Bowman's Space.
5. Select the brightest plasma fluorescent intensity within the capillary loop section and circle this region. Using the threshold function, highlight the bright values (usually located at the edges of the capillary loop walls), avoid circulating RBC shadows, and record the value.
   NOTE: As factors within the blood will cause an underestimation of plasma fluorescence levels, it is important to select the brightest areas.
6. Enter the values in a spreadsheet to calculate the GSC using Eq (1):
   GSC = (1)

8. Calculating red blood cell flow in surface glomerular capillary loops and renal vasculature using a linescan function

1. Find an appropriate vessel (either a capillary loop or a peritubular vessel). As the linescan function in the referenced image acquisition software (see the Table of Materials) requires the vessel to be perpendicular, rotate the image using the rotate function.
2. Once the vessel is rotated and lying flat, select the XT function on the acquisition menu. Set up to scan 4,000 lines. Place the line across the vessel to be examined; ensure the focal plane is at the maximum diameter of the segment to be imaged.
3. Left-click on the color composite image and select Take Snapshot to generate a reference image of the area where the linescan was taken. Immediately click the Start button to capture the linescan of the vessel.
4. To determine the RBC flow rate, import the linescans into the image processing software (see the Table of Materials). Open the Show Region Statistics dialog box under the Measure dropdown menu. Select the single line drawing tool and draw a line that matches the slope of the RBC shadows. Note the width and height values.
   NOTE: The pixel values obtained for width will correspond to distance; the pixels for height correspond to time.
5. Use the following formula (Eq (2)) to calculate speed.
   RBC flow rate in µm/s = (2)
   NOTE: This corresponds to acquisition parameters at 60x magnification and a 400 Hz scan rate with the referenced microscope (see the Table of Materials).
   1. Make at least five calculations and average them to report the speed for each linescan.
      NOTE: These parameters will be dependent on the pixel dimension and acquisition speed of the microscope system.

9. Calculating WBC occlusion in glomerular capillary loops

1. Administer Hoechst 33342 nuclear stain (at ~8 µg/kg rat weight) via an indwelling venous access line to identify WBCs lodged in the capillary loops.
   NOTE: The usable depth will be limited due to photon scatter and absorption by hemoglobin, especially for shorter wavelength blue emissions.
2. Center a glomerulus in the imaging field and take a 3D data set starting at the glomerular surface and end the data at least 30 to 35 µm. Use a 1 µm step size in the Z-direction.
3. Identify WBCs by comparing the blue Hoechst channel with the Texas Red albumin channel; look for exclusion of red dye in the capillary loop and a corresponding nuclear stain to positively identify the WBCs. Define white blood cells as "adhered" if they appear static over 3 optical sections. Report the values as occurrence/10 optical sections from the top of the glomerulus, taken at 1 µm intervals.

10. Scoring the presence of Rouleaux formations in surface glomeruli

1. Follow the same directions for step 9.3 and acquire a 3D data set. Look for Rouleaux formations appearing as RBCs stacked into packets that resist dissociation as they move along the capillary loops. Use a red fluorophore for better visualization of structure at greater depths due to less photon scatter for the longer wavelength red emissions. Report the values as occurrence/25 optical sections from the top of the glomerulus, taken at 1 µm intervals.

11. Isolation of glomeruli

1. Isolate three groups of glomeruli from fresh kidneys using a standard sieving technique that results in close to 90% purity of rat glomeruli¹⁵.
   1. Place the kidney cortex in cold phosphate-buffered saline (PBS) and mince it using multiple fine scissors or razor blades.
   2. Add the minced tissue to a 100 µm sterile cell strainer and gently push it through using a 5 mL syringe plunger and 50-100 mL of cold PBS.
      NOTE: Most of the tubules are retained while the glomeruli pass through.
   3. Place the enriched glomeruli fraction onto a 70 µm filter and wash them extensively with cold PBS. Wash the filter with 100-200 mL of cold PBS to remove most of the remaining tubules.
   4. Collect the glomeruli from the filter using 1-2 mL of cold PBS, centrifuge (10,000 × g, 2 min, 4 °C) and snap-freeze them in liquid nitrogen until RNA isolation.
      NOTE: Glomerular purity determined by phase-contrast microscopy is >90%, and the yield is approximately 10 mg from 2 kidneys.

12. Glomerular RNA isolation

1. Homogenize the frozen glomeruli pellet using the RNA isolation reagent by adding 400 µL of the reagent and breaking up the pellet using a 200 µL pipet tip, followed by brief vortexing and 5 min of incubation at RT¹⁶.
2. Add 40 µL of 1-bromo-3-chloropropane (BCP), vortex for 15 s, and hold at RT for 15 min.
3. Centrifuge at 12,000 × g, 15 min, 4 °C. Remove the aqueous layer, dilute the lower layer with an equal volume of 70% ethanol, and load it directly onto a spin column (see the Table of Materials).
4. After the washes, each consisting of adding the respective solution to the column followed by centrifugation at 12,000 x g, 15 s, 26 °C [3 total, first 2 with 500 µL of RPE (proprietary mild washing buffer with ethanol to remove traces of salts, 3^rd wash with 500 µL of 80% EtOH], elute the RNA by the addition of 15 µL of H₂O and centrifuge, as for the washes. Check the concentration and purity of the RNA and transport the RNA samples to the core facility for Nanostring analysis^17,18.
   NOTE: Total RNA yield is approximately 1-2 µg. Here, the 24 samples contained 200 ng of RNA at 30 ng/µL.

13. Nanostring analysis

NOTE: Nanostring technology is based on the digital detection and direct molecular barcoding of target molecules that utilize color-coded probe pairs. The Capture probe carries a biotin moiety on the 3' end, and the Reporter probe carries the signal on its 5' end.

1. Ship the Nanostring gene probe pairs and CodeSets to Michigan State's Genomic Core Facility and use as directed by NanoString.
   NOTE: There are six positions for the color codes, and each position can be one of four colors, enabling a large diversity of tags that can each be individually resolved and identified during data collection.
2. Obtain the data collected by the Genomic Core facility staff using the proprietary Nanostring nCounter Digital analyzer, which collects fields of view using a microscope objective and CCD camera and tabulates and displays the barcode counts.
3. Import the raw data into Nanostring's nSolver software for analysis. Normalize the data normalized using their default settings and compare data between groups as outlined in their manual.
   NOTE: The objective was to monitor gene changes previously shown to be altered in cortical tissue from a UUO model¹⁹, kidney disease^{17,20,21,22,23,24,25,26}. A total of 126 genes, including the recommended positive and negative controls, were analyzed in each glomeruli group (CONT, SHAM, & UUO).