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Method Article

Effective and Safe Gene Delivery to the Mouse Kidney via Slow Retrograde Renal Pelvis Injection of Adeno-Associated Virus Vectors

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DOI:

10.3791/67716

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August 1st, 2025

 ,  ,  ,  , 
1Department of Molecular and Medical Genetics, Oregon Health & Science University, 2Department of Molecular Microbiology and Immunology, Oregon Health & Science University, 3Division of Neuroscience, Oregon National Primate Research Center

In This Article

Summary

This report details the localized injection method of AAV vectors via the renal pelvis for effective and safe gene transfer into the mouse kidney.

Abstract

The development of effective in vivo tools and methods for gene delivery to the kidney is crucial for advancing basic kidney research and gene therapy for kidney diseases. In addition, growing awareness of monogenic kidney diseases, driven by advanced genetic testing, underscores the potential of gene therapy to treat and even cure difficult-to-treat genetic kidney diseases. In this regard, adeno-associated virus (AAV) vectors have garnered increasing attention as a robust platform for in vivo gene delivery; however, the most effective and safest method for AAV vector-mediated gene delivery to each therapeutically relevant cell type in the kidney has not yet been fully established. Here, the method of slow retrograde renal pelvis (RP) injection of AAV vectors is detailed, and its high potential to transduce mouse kidneys is demonstrated when used with appropriately selected AAV capsids. Moreover, this method is shown to be effective not only with the standard cesium chloride ultracentrifugation-purified AAV (CsCl AAV) vectors but also with centrifugally ultrafiltered AAV (CU AAV) vector minipreps, which can be prepared rapidly and simply using techniques that do not require specialized AAV vector expertise. As an example, this study demonstrates slow RP injection of both CsCl and CU AAV-KP3 vectors results in robust transduction of proximal tubules in the mouse kidney with no apparent tissue damage, unlike previously reported hydrodynamic approaches that inevitably lead to tissue damage. Thus, this study highlights that slow retrograde RP injection of select AAV capsid-derived vectors is an effective and safe method for gene delivery to the kidney. This method will be applicable to a broad spectrum of kidney research, including gene therapy studies. The use of CU AAV vector minipreps will significantly increase throughput and improve the time and cost efficiency needed to generate preliminary data and obtain crucial insights.

Introduction

Effective and safe methods for gene delivery to the kidney in small and large experimental animals provide crucial opportunities to advance basic kidney research and explore innovative therapeutic approaches. Establishing such methods is particularly relevant to gene therapy for various kidney diseases. Recent advancements in clinical genetic testing have revealed a higher prevalence of monogenic renal disorders than previously estimated, affecting approximately 30% of patients with chronic kidney diseases (CKDs)1. CKD is characterized by progressive renal damage, ultimately leading to kidney failure with no truly effective disease-modifying therapies. Hemodialysis and kidney transplantation are currently available treatments; however, they have limitations that significantly impair the quality of life for patients with CKD2. Thus, gene therapy, a clinically proven powerful approach to treating genetic diseases, has garnered significant attention in recent years as a novel avenue for the effective treatment of genetic kidney diseases leading to CKD3,4,5,6. This approach can directly address the root causes of the diseases and potentially offer a cure.

Despite the steady advancements in the field, gene delivery to the kidney remains a significant challenge in both research and therapeutic settings. Efficiently targeting kidney cells with therapeutic genes is difficult due to the organ's complex structure and unique physiological barriers, including the glomerular filtration barrier, which is composed of glomerular endothelial cells, the glomerular basement membrane, and podocyte foot processes. Various strategies have been explored for kidney-targeted gene delivery, including direct renal cortex (RC) injection7,8, renal artery (RA) injection9,10,11, renal vein (RV) injection12,13,14, renal pelvis (RP) injection15,16,17,18,19,20,21,22,23,24,25 and intravenous (IV) injection19,26,27,28,29,30 of viral or non-viral gene delivery vectors, as well as ultrasound and microbubble-mediated nucleic acid delivery31,32. These approaches demonstrated variable transduction efficiencies or functional outcomes, often yielding inconsistent results that are difficult to reconcile (reviewed in references3,33,34,35). Gene delivery to the kidney is still underdeveloped, making direct comparisons between vector systems and methodologies challenging. However, localized delivery methods, which facilitate the administration of highly concentrated agents directly to the kidney, have generally achieved better results than systemic IV delivery. In addition, adeno-associated virus (AAV) vectors, with their robust safety profiles and proven track record of clinical translation36,37, have been the most studied and are regarded as the leading choice for renal gene transfer.

Common AAV serotypes and engineered capsids have demonstrated effective transduction of mesangial and mesenchymal cells in the kidney following IV injection in mice28,30,38. Notably, recent studies have shown that IV administration of AAV9 can effectively transduce podocytes in mouse models of nephrotic syndrome and Alport syndrome, although this effect is not observed in healthy kidney38,39, and that IV administration of an engineered AAV2 capsid can exclusively target glomerular endothelial cells in both healthy and inflamed kidneys in mice40. Historically, renal tubules have been challenging to transduce via the IV route. However, it has recently been discovered that RP injection of select AAV capsids, such as AAV-KP141, can achieve transduction of proximal tubules at levels more than an order of magnitude higher than those achieved with the benchmark AAV9 capsid38. Although further improvements are needed, these advancements open new avenues for basic kidney research and gene therapy, offering opportunities that were previously unattainable.

In the present article, slow retrograde RP injection of AAV vectors is detailed as an effective and safe method for gene delivery to the mouse kidney38. As a representative example of this technique, AAV-KP3 was chosen as an exemplary capsid based on the discovery that it is one of the six capsids (i.e., AAV-KP1, AAV-KP2, AAV-KP341, AAV-DJ42, AAV2G943, and AAV2.7m844) that showed enhanced renal transduction compared to AAV9 following RP injection in mice38. Notably, slow retrograde RP-injected kidney tissues showed no apparent tissue damage in contrast to those treated with hydrodynamic approaches previously reported23,24. Moreover, AAV miniprep vectors45 crudely purified by quick and simple centrifugal ultrafiltration (i.e., CU AAV vectors) achieved renal transduction levels approximately 70% of those attainable by AAV vector preparations purified by the standard cesium chloride (CsCl) gradient ultracentrifugation (i.e., CsCl AAV vectors) when administered at the same vector genome titer. Thus, the method and observations presented here provide a valuable and practical resource for researchers seeking to enhance gene delivery to the kidney in mice and potentially larger animals.

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Protocol

All animal experiments described here were approved by the Institutional Animal Care and Use Committee at Oregon Health & Science University (OHSU) and were performed in accordance with the guidelines for animal care at OHSU. In this study, eight-week-old C57BL/6J male mice were used. The reagents and the equipment used in this study are listed in the Table of Materials.

1. AAV vector production by an adenovirus-free plasmid transfection

  1. Culture human embryonic kidney (HEK) 293 cells in Dulbecco's Modified Eagle Medium (DMEM)-high glucose (4.5 g/L) supplemented with 10% fetal bovine serum (FBS), 1% Penicillin and Streptomycin Mix, and 1 mM L-glutamine, in a 37 °C incubator with 5% carbon dioxide (CO2).
  2. On Day -2 (48 h prior to transfection), plate 4 x 106 HEK 293 cells per T225 flask in 40 mL of the culture medium as described in step 1.1. This reaches ~95% confluency on Day 0.
    NOTE: The culture scale can be adjusted based on the application. For an AAV miniprep (step 2), prepare an appropriate number of flasks based on the target titer and vector concentration, considering that the purification process typically results in an approximate 50% loss of material. To achieve >6 x 1012 vg/mL of AAV9 and AAV-KP3 vectors for RP injection in mice at a dose of 3 x 1011 vg/mouse in a 50 µL vector solution, at least 2 flasks are recommended. For larger-scale AAV vector production, such as that for CsCl density gradient-purified AAV vectors (CsCl AAVs), 25 or more flasks are typically used.
  3. Day 0: Plasmid DNA transfection
    1. Preparation for transfection
      1. Make sure that cells have reached ~95% confluency.
      2. Warm up DMEM supplemented with 1% Penicillin and Streptomycin Mix and 1 mM L-glutamine but without FBS (i.e., serum-free medium) in a 37 °C water bath.
      3. Allow the linear polyethylenimine (PEI) solution to reach room temperature.
    2. Preparation of plasmid DNA mixture
      1. Mix the plasmids (15 µg each of pAAV-CAG-tdTomato, pRepCap, and pHelper) in 1 mL of phosphate-buffered saline (PBS) without CaCl2 or MgCl2 in sterile 1.5 mL microcentrifuge tubes. The total amount of DNA is 45 µg/flask.
        NOTE: Plasmid DNA solution volumes can be disregarded if they are minimal. However, volume adjustment is advised if the total solution volume is ≥225 µL. The sequences of the plasmids used in this study are available in Supplementary File 1.
    3. PEI transfection
      1. Add 90 µL of PEI solution (1 mg/mL) to the 1 mL of PBS and mix with the PBS-plasmid DNA mix (prepared as above) in a 50 mL polypropylene tube. The final volume is approximately 2 mL (5% volume of the culture medium).
      2. Vortex the tubes for 5 s and briefly spin the tubes with a centrifuge to collect the liquid to the bottom of the tubes.
    4. Keep at room temperature for 15 min to form plasmid DNA-PEI complexes.
    5. Once the 15-min incubation is complete, add the 40 mL of prewarmed serum-free medium prepared in step 1.3.1.2 to the plasmid DNA-PEI complexes. The volume is approximately 42 mL.
    6. Replace the culture medium of the T225 flask with plasmid DNA-PEI complexes containing serum-free medium described in step 1.3.4.
    7. Maintain the cells in this transfection medium until the harvest on Day 5 (no medium change is required).
    8. On Day 2, observe cells transfected with the plasmid expressing fluorescent proteins (e.g., pAAV-CAG-tdTomato) under an inverted fluorescence microscope to assess plasmid transfection efficiency.
      NOTE: The conditions described above typically result in transfection efficiencies exceeding 70%. Some morphological changes (e.g., cell elongation) may be observed, likely due to the serum-free environment.
    9. On Days 3 to 5, continue to culture the transfected cells in a 37 °C incubator with 5% CO2.
    10. On Day 5, collect both cells and virus-containing medium into a 50 mL polypropylene tube.
    11. Store the samples at -80 °C until use.

2. Centrifugal ultrafiltration (CU) purification of AAV vectors (AAV miniprep)

  1. Thaw the frozen sample from step 1.3.11 in a 37 °C water bath.
  2. Keep 1 mL of crude lysate aside for DNA dot blot assay (used as Fraction 1).
  3. Add 80 µL of 1M magnesium chloride (MgCl2, final concentration of 2 mM), 160 µL of 0.1M sodium hydroxide (NaOH, final concentration of 0.4 mM), and 16 µL of Serratia marcescens endonuclease (final concentration of 100 units/mL) into a 50 mL tube. Incubate at 37 °C for 1 h. Vortex the sample every 15 min during the incubation.
  4. Collect the Serratia marcescens endonuclease-treated sample into a 250 mL centrifuge bottle and centrifuge at 10,000 x g at 4 °C for 30 min.
  5. Transfer the supernatant to a 50 mL polypropylene tube using a 50 mL serological pipette.
  6. Filter the supernatant to remove debris using a 0.45 µm PVDF syringe filter. Keep 1 mL of 0.45 µm-filtrated sample aside for DNA dot blot assay (used as Fraction 2).
  7. Add 10 mL of molecular grade water to a 100 kDa molecular weight cut-off (MWCO) centrifugal ultrafiltration tube and centrifuge at 2,000 x g at 4 °C for 10 min. Remove the flow-through.
  8. Add 15 mL of 5% sorbitol in PBS to the centrifugal ultrafiltration tube and centrifuge at 2,000 x g at 4 °C for 10 min. Remove the flow-through.
  9. Add up to 15 mL of 0.45 µm-filtrated sample as described in step 2.6 and centrifuge at 2,000 x g at 4 °C for 15 min for concentration until the target volume of approximately 500 µL is achieved. Keep 1 mL of flow-through aside for DNA dot blot assay (used as Fraction 3).
    NOTE: Before each centrifugation, resuspend the retentate with a pipette to disrupt the concentration gradient. Do not touch the membrane with the pipette tip. If the target volume is not reached after 15-min centrifugation, extend the centrifugation as needed by adjusting the duration based on the remaining volume in the tube.
  10. Repeat step 2.9 until the entire sample is filtered, reaching the target volume of approximately 500 µL. Keep 1 mL of the flow-through from the first and second iterations of step 2.10 aside for DNA dot blot assay (used as Fraction 4 and Fraction 5).
    NOTE: A single centrifugal ultrafiltration tube can be used to process samples obtained from two or more flasks.
  11. Add 15 mL of 5% sorbitol in PBS and centrifuge at 2,000 x g at 4°C for 15 min until the target volume of approximately 500 µL is achieved. Keep 1 mL of flow-through aside for DNA dot blot assay (used as Fraction 6).
  12. Repeat step 2.11 (used as Fraction 7).
    NOTE: If higher AAV titers are required for downstream experiments, the final target volume can be further reduced to 150 µL.
  13. Transfer the ultrafiltered AAV vector to a 2 mL screw cap tube with an O-ring. Keep 10 µL of ultrafiltered AAV vector aside for DNA dot blot assay (used as Fraction 8).
  14. Determine AAV titers by DNA dot blot assay as previously described46.
    NOTE: CU-purified vectors (CU AAV) contain higher levels of empty capsid contamination and host cell-derived impurities compared to CsCl-purified vectors (CsCl AAV). These factors may impact transduction efficiency and immune responses, requiring careful consideration in data interpretation.

3. CsCl density-gradient purification of AAV vectors

  1. Thaw the frozen sample from step 1.3.11 in a 37 °C water bath.
  2. Centrifuge at 10,000 x g at 4 °C for 30 min to remove the cell debris.
  3. Collect the supernatant and precipitate AAV vectors in the supernatant by adding Polyethylene Glycol 8000 (PEG, final concentration of 8%) and sodium chloride (final concentration of 500 mM).
  4. Incubate on ice for 3 h.
  5. Centrifuge at 10,000 x g at 4 °C for 30 min to obtain the pellets containing AAV.
  6. Remove the supernatant and resuspend the pellets with 10ml of Sarkosyl containing resuspension buffer (50 mM HEPES, 150 mM NaCl, 1% Sarkosyl, pH 8.0).
  7. Transfer the pellet suspension to a 15 mL polypropylene tube and keep it at 4 °C overnight.
  8. The following day, treat the pellet suspension with Serratia marcescens endonuclease (final concentration of 200 units/mL) and MgCl2 (final concentration of 2 mM).
  9. Incubate at 37 °C for 1h.
  10. Centrifuge at 10,000 x g at 4 °C for 30 min.
  11. Collect the supernatant and add cesium chloride (CsCl, final concentration of refractive index (RI) = 1.3710 +/- 0.0005) and EDTA (final concentration of 20 mM).
  12. Load the sample into a 29.9 mL ultracentrifuge tube and add topping solution (50 mM HEPES, 150 mM NaCl, 20 mM EDTA, CsCl (RI = 1.3710 +/- 0.0005), pH 8.0) until it is full.
  13. Centrifuge at 41,000 x g at 12 °C for 22 h.
  14. Punch holes in the bottom and side of the ultracentrifuge tube to collect a total of 15 fractions of 2 mL each.
  15. Use 10 µL of each fraction to check RI and perform the DNA dot blot assay46.
  16. Based on the RI and DNA dot blot assay results, determine the best 5 fractions that contain AAV vectors.
  17. Combine the best 5 fractions (total about 10 mL) and adjust RI to 1.3710 +/- 0.0005.
  18. Load the sample into an 11.2 mL ultracentrifuge tube and add the topping solution until it is full.
  19. Centrifuge at 65,000 x g at 12 °C for 12 h.
  20. Punch holes in the bottom and side of the 11.2 mL ultracentrifuge tube to collect a total of 18 fractions of 0.5 mL each.
  21. Use 10 µL of each fraction to check RI and perform the DNA dot blot assay46.
  22. Based on the RI and DNA dot blot assay results, determine the best 6 fractions that contain AAV vectors.
  23. Combine the best 6 fractions (total of about 3 mL) and load them into a dialysis cassette.
  24. Float the dialysis cassette in 1.3 L of dialysis buffer (PBS buffer supplemented with 0.001% non-ionic surfactant) with a stir bar for 2 h to remove the CsCl.
  25. Repeat the step 3.24 two times (total 6 h of dialysis).
  26. Transfer the cassette to 500 mL of 5% sorbitol in PBS buffer supplemented with 0.001% surfactant for 2 h for buffer exchange.
  27. Collect the sample from the dialysis cassette with a syringe and filter the collected sample using a 0.22 µm PVDF syringe filter.
  28. Determined AAV titers by a quantitative dot blot assay46.

4. Quality assessment of AAV vector preparations by SDS-PAGE followed by silver staining

  1. Mix vector preparations, 1 x 1010 vg for CsCl AAV vectors, and 1 x 109 and 1 x 1010 vg for CU AAV vectors, with 6x Laemmli buffer in a total 18 µL (vector preparation:6x Laemmli = 15:3) and heat them at 100 °C for 5 min.
  2. Load the heat-treated AAV vector preparations onto a 7.5% SDS-PAGE gel and electrophorese the samples at 20 mA using Tris-glycine-SDS gel running buffer.
  3. Remove the gel from the gel cassette, rinse it with ultrapure water, and stain it using a silver staining kit according to the manufacturer's instructions.
  4. Incubate the gel loaded with CsCl AAV vectors in the developer solution for 7 min, whereas incubate the gel loaded with CU AAV vectors for 2 min to avoid signal saturation.
    NOTE: CU AAV vector preparations contain higher levels of protein impurities compared to CsCl AAV vectors. Therefore, gels loaded with CsCl AAV and CU AAV vectors were separated after gel running and silver-stained separately to prevent signal saturation due to the high sensitivity of the staining method.

5. Identification of AAV VP proteins in CU AAV vector preparations by western blot

  1. Mix 1 x 109 vg of CU AAV vector preparations with 6x Laemmli buffer in a total of 18 µL (vector preparation:6x Laemmli = 15:3) and heat them at 100 °C for 5 min.
  2. Load the heat-treated AAV vector preparations onto a 7.5% SDS-PAGE gel and electrophorese the samples at 20 mA using Tris-glycine-SDS gel running buffer.
  3. Electrotransfer proteins in the gel onto a polyvinylidene difluoride (PVDF) membrane, activated in methanol, using a semi-dry transfer system.
  4. Block the membrane with 5% skimmed milk in 1x Tris-buffered saline with Tween 20 (TBST) at room temperature for 1 h.
  5. Incubate the blocked membrane with primary antibody solution (rabbit polyclonal anti-VP antibody, diluted to 1:1000 with 5% skimmed milk in 1x TBST) at 4 °C overnight.
  6. Wash the membrane with 1x TBST for 10 min, 3 times.
  7. Incubate the membrane with secondary antibody solution (goat anti-rabbit IgG antibody conjugated with horseradish peroxidase, diluted to 1:5000 with 5% skimmed milk in 1x TBST) at room temperature for 1 h.
  8. Wash the membrane with 1x TBST for 10 min, 3 times.
  9. Visualize signals on the membrane using a western blot imaging system.

6. Renal pelvis (RP) injection

  1. Preoperative setup and anesthesia
    1. Measure the body weight of each mouse in a plastic beaker on a weight scale that is accurate to 0.1 g.
    2. Administer meloxicam (5 mg/kg) subcutaneously based on the body weight to provide presurgical analgesia.
    3. Dilute the AAV vector solution to a concentration of 6.0 x 1012 vg/mL with 5% sorbitol in PBS and keep it on ice.
      NOTE: Prepare 40 μL more AAV solution than the total injection volume (50 μL x number of mice to be injected) to account for dead space in a gas-tight glass syringe and a silicone tube.
    4. Bend and break off only the needle tip from a 30 G needle using a needle holder.
    5. Under a stereomicroscope, connect the 30 G needle tip, blunt end of the 4 cm PE-10 tube, 15 cm silicone tube, a 30 G needle and a gas tight glass syringe in this order from the tip (Figure 1D). Sterilize both tubes prior to use and perform this assembly sequence on top of a sterile drape to maintain sterility. Sterilize the autoclave-compatible silicone tube by standard steam sterilization. Before sterilizing PE-10 tube, cut one end diagonally to create a beveled tip that facilitates insertion into the silicone tube. Sterilize the PE-10 tube by immersing it in 2% glutaraldehyde for more than 10 hours and then rinse it thoroughly with sterile PBS before use.
    6. Set the connected gas-tight glass syringe onto an infusion syringe pump.
    7. Aspirate 90 µL of AAV vector solution prepared in step 6.1.3 into the gas-tight glass syringe using an infusion syringe pump.
    8. Set the infusion mode of the syringe pump as follows: the target volume of 50 µL and a flow rate of 50 µL/min.
    9. Place a mouse in a plastic chamber filled with 4% isoflurane delivered in 100% oxygen at a flow rate of 1.5 to 3.0 L/min until it is fully anesthetized.
    10. Place the snout of the mouse into a nasal cone with 1.5% to 3.0% isoflurane flow. Monitor the depth of anesthesia by toe-pinch reflex throughout the surgical procedure.
    11. Cover the eyes with ophthalmic ointment using a cotton-tipped applicator to prevent corneal desiccation.
    12. Remove the fur from the left back side of the mouse with a depilatory cream using a cotton-tipped applicator.
      NOTE: Depilatory cream must be completely removed via rinsing to prevent skin irritation.
    13. Place the mouse in a lateral position on a closed-loop heat pad connected to a heat therapy pump to maintain body temperature at 37 °C (no need to secure it with adhesive tape).
    14. Disinfect the shaved area with three alternating applications of povidone-iodine and 70% ethanol using a cotton-tipped applicator in a circular motion beginning at the incision site.
    15. Cover the mouse with a sterile surgical drape.
  2. RP injection to the left kidney
    1. Make an incision of approximately 1 cm to the skin layer with a sterile scalpel at the costovertebral angle level in the left kidney's location.
    2. After making a skin incision, cut the muscle layer at the same length.
      NOTE: If the incision length is too short, the kidney cannot be exposed, and instead, the spleen, which is smaller in size, may be accidentally exposed. On the other hand, if the incision length is too long, the kidney will not be kept in place by the incision itself and will continually slip back into the abdomen through the incision.
    3. Gently press on the abdomen with the fingers, not directly touching the kidney, so that the kidney is exposed through the incision site (Figure 1A).
    4. Minimally remove the surrounding fat so that the renal pelvis is visible as a small white area.
    5. First, clamp the ureter with a straight-type micro vessel clip, and then the renal artery and renal vein together with a curved-type micro vessel clip.
      NOTE: The kidney color turns dark red, confirming the successful blockade of blood flow into and out of the kidney (Figure 1B).
    6. Insert the 30 G needle connected to the syringe pump approximately 3 mm into the pelvic cavity with curved forceps and start an injection of 50 µL of AAV vector solution using an infusion syringe pump at a flow rate of 50 µL/min over 1 min (Figure 1C).
    7. Keep the needle and clamp for 5 min to increase the injected kidney's exposure to the AAV vector.
      NOTE: Appropriate dwelling time may vary and depend on the AAV capsids. A dwelling time of 5 min has been shown to be sufficient for transduction with AAV-KP1 or AAV-KP3 vector.
    8. Once the 5-min incubation is complete, remove the clamps.
      NOTE: The kidney color immediately changes from dark red to blood red, confirming the restoration of the blood flow in the kidney.
    9. Withdraw the injection needle carefully and slowly, simultaneously placing a cotton-tipped applicator over the injection site with pressure. Make sure that there is no bleeding from the injection site. Applying a hemostatic patch is not required.
    10. Gently place the kidney back into the peritoneal cavity.
    11. Suture the muscle layer with 6-0 absorbable suture and the skin with 5-0 monofilament suture.
    12. Administer 1 mL of saline subcutaneously into the back of mice for fluid support.
    13. Replace the used 30 G needle tip with a newly prepared one, discard the used 30 G needle according to the institution's guidelines for hazardous chemical waste management, and load 50 µL of the AAV vector solution using an infusion syringe pump for administration to the next mouse.
    14. Repeat steps 6.2.12-6.2.13 for all mice in the group, except for the last one, for which step 6.2.13 is not required.
    15. After completing all planned injections, discard the contaminated needles and tubes according to the institution's guidelines for hazardous chemical waste management.
  3. Postoperative monitoring and care
    1. Administer meloxicam (5 mg/kg) subcutaneously into the back of mice for postsurgical analgesia 24 h and 48 h after RP injection.
    2. Administer 1 mL of saline subcutaneously into the back of mice for fluid support 24 h after RP injection.
    3. Monitor the body weight and general appearance of injected mice daily for 1 week postoperatively. If a 10% body weight loss occurs, administer 1mL of saline subcutaneously into the back of mice three times a week for a week. If no improvement is observed, affected mice are euthanized.

7. Tissue harvesting and cryosectioning

  1. Preparation of fixation and cryoprotection solution.
    1. 0.2 M phosphate buffer (total 1 L)
      1. Prepare 800 mL of ultrapure water in a 1 L glass beaker.
      2. Add 6.4 g of sodium phosphate monobasic (NaH2PO4) and 21.8 g of sodium phosphate dibasic (Na2HPO4) to the solution.
      3. Adjust the pH to 7.4 with hydrochloric acid.
      4. Fill with ultrapure water to 1 L.
      5. Store at 4 °C until use.
    2. 4% paraformaldehyde (PFA) solution (total 40 mL)
      1. Mix 10 mL of 16% PFA, 20 mL of 0.2M phosphate buffer prepared in step 7.1.1, and 10 mL of ultrapure water.
      2. Aliquot 10 mL into a 15 mL polypropylene tube.
        NOTE: Make 4% PFA solution just before use for perfusion fixation.
    3. 30% sucrose solution (total 40 mL).
      1. Mix 30 mL of PBS with 12 g of sucrose.
      2. After dissolving completely, fill with PBS to 40 mL.
  2. Perfusion fixation and tissue harvesting.
    CAUTION: Perfusion fixation should be performed in a fume hood as 4% PFA solution is a potent harmful chemical if inhaled.
    1. Anesthetize the mouse with isoflurane, place it in a supine position, and confirm the complete absence of responsiveness by gently pinching the extremities.
      NOTE: The mouse is deeply anesthetized with isoflurane in an induction chamber in accordance with institutional guidelines. For terminal perfusion, anesthesia is continuously maintained by transferring the animal to a nose cone delivering inhalant anesthetic while positioned supine. Adequate depth of anesthesia is confirmed prior to thoracotomy by the complete absence of response to gentle pinching of the extremities. Inhalant anesthesia is maintained without interruption until perfusion is completed and death is ensured.
    2. Once the mouse is anesthetized, make an incision using surgical scissors from the caudal abdomen to the xiphoid. Then, make two lateral incisions along either side of the sternum and retract the chest wall anteriorly to expose the heart and major vessels.
    3. Open the right atrium via sharp dissection to decompress the circulatory system. Insert a 25 G butterfly needle into the left ventricle and infuse 50 mL of PBS at a rate of approximately 10 mL/min, directing the flow toward the ascending aorta.
      NOTE: Exsanguinating is important to obtain high-quality immunofluorescence (IF) images because blood exhibits autofluorescence. To ensure complete removal of blood, 50 mL of PBS was used in this study, although 20-30 mL is commonly used.
    4. Infuse 20 mL of 4% PFA solution prepared in step 7.1.2 at the same rate, ensuring no air is introduced during the switch of solutions.
    5. Harvest tissues and keep them at 4 °C overnight in a 15 mL polypropylene tube filled with 4% PFA solution for post-fixation. When fluorescent protein is expressed in the collected tissue, cover the tube with aluminum foil to shield it from light.
  3. Cryoembedding
    1. Transfer the tissue to a 15 mL polypropylene tube filled with 30% sucrose solution prepared in step 7.1.3 for cryoprotection.
    2. Keep at 4 °C until tissues completely sink to the bottom of the tubes.
    3. Transfer the tissue to a plastic cryomold with standard pattern forceps.
    4. Place the OCT compound over the tissue while ensuring that the tissue is correctly oriented and flush with the bottom of the cryomold.
    5. Fill the top with OCT compound. Remove any bubbles with a pipette tip.
    6. Place the plastic cryomold in a cryogenic bath containing 2-methylbutane, chilled with crushed dry ice in an ice bucket, until the OCT compound is completely frozen (the OCT compound will turn white).
    7. Store the tissue blocks at -80 °C until use.
  4. Cryosectioning
    1. Transfer the frozen tissue blocks to a cryostat microtome at -20 °C prior to sectioning and allow the temperature of the frozen tissue blocks to equilibrate.
    2. Section each frozen tissue block into a desired thickness (typically 5 µm) using the cryostat microtome. Place the tissue sections onto a microscope glass slide for immunofluorescence staining.
    3. Sections can be stored at -80 °C for later use.

8. Immunofluorescence staining

  1. Dry frozen slides at room temperature for 30 min.
  2. Isolate the section with a liquid blocker pen and dry at room temperature for 5 min.
  3. Rehydrate sections with 150 µL of PBS for 10 min.
  4. Place the slides on a flat surface and add 150 µL of 1% bovine serum albumin (BSA) diluted with PBS per section onto the slides. Incubate at room temperature for 45 min for blocking.
  5. Add 150 µL of primary antibody solution (anti-Wilms Tumor 1 (WT1) antibody, diluted to 1:100 with 1% BSA; anti-Na-K-2Cl cotransporter 2 (NKCC2) antibody, diluted to 1:100 with 1% BSA; or anti-aquaporin 2 (AQP2) antibody, diluted to 1:100 with 1% BSA) and incubate at 4 °C overnight under a humid chamber.
  6. Wash the sections with PBS at room temperature for 5 min, 3 times.
  7. Add 150 µL of secondary antibody (anti-rabbit IgG Alexa Fluor 647 antibody, diluted to 1:500 with 1% BSA) and Lotus tetragonolobus lectin (LTL)-fluorescein (diluted to 1:200 with 1% BSA), and incubate at room temperature for 60 min.
  8. Wash the sections with PBS at room temperature for 5 min.
  9. Place the slides on a flat surface and add 150 µL of Hoechst 33342 (diluted to 1:10,000 with PBS) for nuclear staining. Incubate at room temperature for 10 min, shielded from light.
  10. Wash the sections with PBS at room temperature for 5 min.
  11. Mount the slides with 20 µL of the mounting medium and a cover glass. Keep the slides in the dark at 4 °C until imaging is performed.

9. DNA extraction from tissue

  1. DNA extraction from tissues using a commercially available kit (see Table of Materials).
    1. Transfer the tissue into a ceramic bead tube and add 400 µL of DNA extraction buffer (100 mM NaCl, 20 mM EDTA, 1% SDS, and 50 mM Tris-HCl, pH 8.0).
    2. Homogenize the tissue at a speed setting of 5 m/s for 20 s in 3 cycles using a homogenizer.
    3. Centrifuge at 2,000 x g at room temperature for 2 min.
    4. Collect 200 µL of the supernatant using a P200 pipette and add 10 µL of Proteinase K solution (final concentration of 1 mg/mL).
    5. Mix by vortexing for 15 s and incubate at 55 °C for 1 h.
    6. Add 200 µL of lysis buffer from the DNA extraction kit.
    7. Mix by vortexing for 15 s and incubate at 55 °C for 15 min.
    8. Add 250 µL of 100% ethanol.
    9. Mix by vortexing for 15 s and incubate at room temperature for 5 min.
    10. Apply the lysate onto the DNA extraction spin column.
    11. Centrifuge at 6,000 x g at room temperature for 1 min.
    12. Place the spin column in a new 2 mL collection tube. Add 500 µL of the first wash buffer from the DNA extraction kit.
    13. Centrifuge at 6,000 x g at room temperature for 1 min.
    14. Place the spin column in a new collection tube. Add 500 µL of the second wash buffer from the DNA extraction kit.
    15. Centrifuge at 6,000 x g at room temperature for 1 min.
    16. Place the spin column in a new 2 mL collection tube. Add 500 µL of 100% ethanol.
    17. Centrifuge at 6,000 x g at room temperature for 1 min.
    18. Place the spin column in a new 2 mL collection tube and centrifuge at 20,000 x g at room temperature for 3 min.
      NOTE: This step ensures complete drying of the spin column membrane, as ethanol carryover into the eluate can interfere with downstream applications.
    19. Place the spin column in a new 1.5 mL microcentrifuge tube. Add 35 µL of elution buffer from the DNA extraction kit to the spin column.
      NOTE: The volume of recovered eluate is typically about 5 µL less than the volume of elution buffer initially applied to the column.
    20. Incubate at room temperature for 5 min.
    21. Centrifuge at 20,000 x g at room temperature for 1 min.
    22. Measure the DNA concentration.
  2. Vector genome copy number quantification by quantitative PCR (qPCR)
    1. Dilute the extracted DNA prepared in step 9.1 to a concentration of 25 ng/µL with molecular-grade water.
    2. Design a qPCR primer pair for the vector genome quantification (tdTomato was used in this study) and a qPCR primer pair for the mouse reference gene that will be used to quantify the diploid genomes in the extracted DNA samples (the mouse agouti gene was used in this study).
      NOTE: The primer sequences used in this study are as follows;
      tdTomato forward: 5'-TATGAACGGAGCGAGGGGAG-3'
      tdTomato reverse: 5'-TTCCACTAGACCCTGAGCCG-3'
      mouse agouti forward: 5'-GGCGTGGTCAGTGGTTGTG-3'
      ​mouse agouti reverse: 5'-TTTAGCTTCCACTAGGTTTCCTAGAAA-3'
    3. For a 25 µL qPCR reaction, mix 4 µL of diluted DNA sample in step 9.2.1 (a total of 100 ng DNA in a 25 µL reaction mixture), 12.5 µL of 2x qPCR reaction buffer, 0.5 µL of both 20 µM forward and reverse primes prepared in step 9.2.2 (final concentration of 400 nM of each primer), and 7.5 µL of molecular grade water.
    4. Measure the DNA concentration of plasmid standards for qPCR four times and determine the average value of the DNA concentration.
      NOTE: Supercoiled circular pAAV-CAG-tdTomato plasmid and a supercoiled circular plasmid containing the PCR target sequence derived from the mouse agouti gene were used for vector genome quantification and normalization, respectively, in this study.
    5. Prepare a series of 1:10 serial dilutions of plasmid DNA standards covering a dynamic range of 106.
      NOTE: Each sample and standard should be analyzed at least in duplicate.
    6. Perform qPCR using the following thermal cycling conditions: an initial heat activation step at 95 °C for 10 min, followed by 35 cycles of denaturation at 95 °C for 15 s, and annealing/extension at 60 °C for 30 s.
    7. After completing the qPCR reaction and obtaining the double-stranded (ds) copy number values of vector genomes and the mouse reference gene in the samples, determine ds vector genome copy numbers per diploid genomic equivalent (ds-vg/dge) by normalizing the ds vector genome copy numbers to the reference gene copy number in each sample based on standard curves generated from known concentrations of both the vector genome and the reference gene.
      NOTE: Ds-vg/dge can be determined based on the following approximation that 100 ng of mouse genomic DNA represents 16,900 diploid (2N) mouse cells (i.e., 5.92 pg of DNA per diploid mouse cell).

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Results

A recent study has identified a small subset of AAV capsids that exhibits remarkable enhancement of renal transduction compared to AAV9 following RP injection38. The identified subset includes AAV-KP3, a chimeric AAV capsid generated by DNA shuffling41. Therefore, AAV-KP3 was chosen for RP injection in this study, along with AAV9 as the benchmark. AAV9-CAG-tdTomato vector (AAV9) and AAV-KP3-CAG-tdTomato vector (KP3) were produced in HEK293 cells by an adenovirus-free plasmi...

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Discussion

This report has detailed slow retrograde RP injection of AAV vectors as a local administration method for gene delivery to the mouse kidney. One critical step for safely performing RP injection is the adjustment of the injection rate of AAV vectors to avoid parenchymal damage to the injected kidneys. In a previous report on hydrodynamic renal pelvis injection, 100 µL of plasmid DNA solution was injected into the mouse kidney within only 3 s24. This fast injection is essential for increasing t...

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Disclosures

H.N. receives a royalty of AAV-related technologies licensed by Takara Bio Inc. and Capsigen Inc., serves as a consultant for biotech companies, is a co-founder of Capsigen Inc., and holds shares of Capsigen Inc. and Sphere Gene Therapeutics.

Acknowledgements

We thank Guangping Gao and James M. Wilson for providing us with the helper plasmid of AAV9 and Katja Pekrun and Mark A. Kay for the helper plasmid of AAV-KP3.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
0.45 μm PVDF syringe filterMilliporeSigmaSLHVR33RS
1.5 mL microcentrifuge tube Fisher Scientific05-408-129
15 mL polypropylene tubeCorning352097
16% paraformaldehyde (PFA)Electron Microscopy Sciences15710
250 mL centrifuge bottleFisher Scientific055641S24
2F Silicone tubingSai Infusion TechnologiesSIL-2-50 
25G butterfly needleMedex Supply26708
2-methylbutaneMilliporeSigmaMX0760
2x qPCR reaction bufferFisher Scientific43-676-59Power SYBR Green PCR Master Mix
30G needleBecton-Dickinson & Co305106
50 mL polypropylene tubeCorning430291
5-0 monofilament sutureEthiconY463G
5 M NaClLonza51202
6-0 absorbable sutureEthiconJ489G
7.5% Mini-PROTEAN TGX Precast
Protein Gels, 10-well, 30 µL		Bio-Rad4561023
Amersham ImageQuant 800 Western
Blot Imaging System (Cytiva)		Cytiva29459405
Amicon 100 kDa MWCO, 15 mL tubeMillipore SigmaUFC910024
Anti-aquaporin 2 (AQP2) antibodyAbcamab199975A marker for collecting duct cells
Anti-Na-K-2Cl cotransporter 2
(NKCC2) antibody		StressMarq BiosciencesSPC-401A marker for thick ascending limb
cells
Anti-rabbit IgG Alexa Fluor 647
antibody		Jackson ImmunoResearch111-605-144
Anti-Wilms Tumor 1 (WT1) antibodyAbcamab89901A marker for podocytes
Beckman Coulter OptiSeal Tube,
Polypropylene, 29.9 mL		Fisher ScientificNC9691210
Beckman Coulter OPTISEAL TUBESFisher ScientificNC9611575
BenzonaseSigma-Aldrich1016970001
Bovine serum albumin (BSA)Sigma-Aldrich5470
Centrifugal ultrafiltration tubeMilliporeSigmaUFC9100Amicon 100 kDa MWCO, 15 mL
Ceramic bead tubeFisher Scientific15-340-154
Cesium chlorideSigma-AldrichC3032-1KG
Clamp applying forcepsFine Science Tools00071-14
Closed-loop heat padStryker Medical8002-062-012
Cotton-tipped applicatorPuritan806-WC
Cover glassFisher Scientific12-541B
Cryostat microtomeTanner ScientificTN50
Curved forcepsFine Science Tools11052-10
Curved-type micro vessel clipKleinert-Kutz65145Clamp for the renal artery and renal
vein
Dialysis cassetteFisher ScientificPIA52976
DNA extraction kitQiagen57704QIAamp MinElute Virus Spin Kit
D-SorbitolSigma-AldrichS6021-1KG
Dulbecco's Modified Eagle Medium-
high glucose		Corning15-013-CVDMEM-high glucose (4.5 g/L)
EDTAFisher Scientific15-575-020
EthanolDecon Laboratories2716GEA
Fetal bovine serum (FBS)VWR89510-186
Gas-tight glass syringeHamilton1710 TLL
Glutaraldehyde solutionSigma-AldrichG6257
Heat therapy pumpKent ScientificHTP-1500
HEPESSigma AldrichH4034-100G
Hoechst 33342InvitrogenH3570
HomogenizerFisher Scientific15-340-163
Human embryonic kidney (HEK) 293
cells		Agilent240073
Ice bucketFisher Scientific07-210-104
Infusion syringe pumpHarvard Apparatus70-4507
IsofluranePiramal Critical CareNDC 66794-017-10
L-glutamineFisher Scientific25030-081
Liquid blocker penTed Pella22309
Lotus tetragonolobus lectin (LTL)-
Fluorescein		Vector LaboratoriesFL-1321-2A marker for proximal tubules
Magnesium chloride (MgCl2)MilliporeSigmaM1028
MeloxicamVetOneNDC 86136-012-10Analgesic
Microscope glass slideFisher Scientific12-550-15
Mounting mediumSouthernBiotech0100-01
Needle holderFine Science Tools12501-13
OCT compoundSakura Finetek4583
Ophthalmic ointmentBausch & LombNDC 24208-313-34
PE-10 polyethylene tubeBD427401Inner diameter: 0.28 mm
Penicillin & Streptomycin MixFisher Scientific15140-122
Phosphate-buffered saline (PBS)Fisher Scientific10010-049
Plastic cryomoldSakura Finetek4566
Poloxamer 188 Non-ionic Surfactant
(10%)		Fisher Scientific24-040-032
Polyethylene Glycol 8000Fisher ScientificBP233-1
Polyethylenimine (PEI)Polysciences23966
Povidone iodineRicca Chemical Company395516
Power Blotter–Semi-dry Transfer
System		Fisher ScientificPB0010
Proteinase K solutionFisher Scientific25530049
Rabbit anti-Anti-Adeno-Associated
Virus (AAV), VP1, VP2, VP3		American Research Products03-61084
SarkosylFisher ScientificBP234-500
Screw cap 2 mL tubeCorning AxygenSCT-200-SS-O-S
Silver Stain KitSigma-AldrichPROTSIL1-1KT
Sodium chloride (NaCl)Fisher ScientificS271
Sodium phosphate dibasic
(Na2HPO4)		MilliporeSigmaS0876
Sodium phosphate monobasic
(NaH2PO4)		MilliporeSigmaS0751
SorbitolMilliporeSigmaS6021
Standard pattern forcepsFine Science Tools11000-12
Sterile surgical drapeSAI Infusion TechnologiesPSS5-1519
Straight-type micro vessel clipFine Science Tools00396-01Clamp for the ureter
SucroseFisher ScientificS5
Surgical scissorsFine Science Tools14058-09
T225 flaskCorning431082
Technocut Scalpel #11Myco Medical Supplies Inc.6008T-11
Tissue forcepsRoboz Surgical InstrumentRS-5155

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AAV VectorsRetrograde Renal InjectionKidney Gene TherapyProximal Tubule TransductionCsCl AAVCU AAVKidney Disease ModelsIn Vivo Transduction
Effective and Safe Gene Delivery to the Mouse Kidney via Slow Retrograde Renal Pelvis Injection of Adeno-Associated Virus Vectors
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