This protocol describes a method to inject plasmid DNA into the mouse kidney via the renal pelvis to produce transgene expression specifically in the kidney.
Hydrodynamic injection creates a local, high-pressure environment to transfect various tissues with plasmid DNA and other substances. Hydrodynamic tail vein injection, for example, is a well-established method by which the liver can be transfected. This manuscript describes an application of hydrodynamic principles by injection of the mouse kidney directly with plasmid DNA for kidney-specific gene expression. Mice are anesthetized and the kidney is exposed by a flank incision followed by a fast injection of a plasmid DNA-containing solution directly into the renal pelvis. The needle is kept in place for ten seconds and the incision site is sutured. The following day, live animal imaging, Western blot, or immunohistochemistry may be used to assay gene expression, or other assays suited to the transgene of choice are used for detection of the protein of interest. Published methods to prolong gene expression include transposon-mediated transgene integration and cyclophosphamide treatment to inhibit the immune response to the transgene.
The hydrodynamic tail vein injection technique has become a commonly used way to achieve high levels of gene expression in mouse liver1,2. The kidneys are also transfected by this technique at a much lower level, approximately 100-fold less3. The hydrodynamic renal pelvis injection described here provides a simple way to control the specificity of organ expression through physical means using the same hydrodynamic principles that have been established previously in liver4,5, muscle6, and other organs7,8. This method transfects cells in live animals in vivo by using pressure and speed to force fluid containing DNA into the cells, simultaneously inducing damage to the organ that is quickly resolved9. Using well-established surgical techniques to visualize the kidney via a flank incision10 along with a single injection by insulin syringe, we have found successful transfection of various types of kidney cells, mainly interstitial fibroblasts, tubules, and collecting duct11. Dissection of these mice has shown that other organs are not transfected at levels high enough to visualize by luciferase imaging techniques11. Since the technique is non-viral, use of plasmid DNA for transfection permits fast and easy preparation of the reagents required for injection.
We have used localized hydrodynamic injections to express the antioxidant glutathione S-transferase A4, the insulin-like growth factor-1 receptor, and the hormone erythropoietin in the kidney, all with the expected biological effects11,12,13. Detailed evaluation of route of administration, injection volume, DNA dosage, and promoter choice has been performed11. Additionally, both the piggyBac transposon system and/or cyclophosphamide treatment to suppress the immune reaction to the transgene have been shown to improve long-term gene expression outcomes11. Other investigators have used a renal vein approach in rat with success, achieving high transfection efficiency for time periods of greater than one month14. However, genetic correction of phenotypes mimicking human disease are usually performed in mice first as a proof-of-concept since most mammalian genetic models are mouse models. We compared renal vein injection to renal pelvis injection and found that injection into the renal pelvis was superior to the renal vein for gene expression (approximately ten-fold higher) and survival11. The renal pelvis is an ideal route of entry into the kidney because it is flexible enough to tolerate fluctuations in urine production and is often able to maintain its structural integrity even when dilated during hydronephrosis. Additionally, injection into the renal pelvis allowed access to the kidney without piercing the kidney capsule, allowing the injected fluid to be visibly retained by the kidney better than intraparenchymal injection. Other mouse organs do not have a route of entry other than the vasculature, but the urinary space of the kidney is an ideal injection site. Additionally, injection into the renal vein resulted in leakage of blood into the abdominal cavity. The total kidney volume of wild-type mouse kidneys has been estimated by magnetic resonance imaging to be approximately 0.2 cm3, so the volume of a single kidney is approximately equal to the amount of fluid injected by renal pelvis hydrodynamic injection (100 µL)15. Herein, we have made available all of the detailed nuances of the hydrodynamic renal pelvis injection protocol to achieve reproducible transfection of the kidney.
All methods described here have been approved by the Institutional Animal Care and Use Committees (IACUCs) of Baylor College of Medicine and Vanderbilt University Medical Center.
1. Prepare the DNA solution for injection
2. Perform the hydrodynamic renal pelvis injection surgery
Figure 1. Correct incision site and needle placement for hydrodynamic renal pelvis injections. A) The incision (red line) should be located approximately 1 cm from the spine and approximately 1 cm below the ribcage of the mouse. B) After the kidney is exposed via the flank incision, the renal pelvis should be located as a small yellowish clear/white dot midway down the kidney. The injection should not disturb the renal vein, renal artery, or ureter. The needle of the insulin syringe is inserted directly into the renal pelvis as shown to a depth of approximately 0.5 cm and quickly depressed in 2-3 s. Please click here to view a larger version of this figure.
Figure 2. The surgical steps to perform renal pelvis hydrodynamic injection of plasmid DNA. A) Forceps pinch the skin to allow the surgeon to make a ~1 cm flank incision with a scalpel, first through the skin layer, then through the muscle layer. B) Using two pairs of closed forceps to open the surgical wound, the kidney is visualized within the abdomen if possible. C) With gentle pressure on the abdomen, without touching any organs directly, the kidney is exposed through the flank incision. D) Fat is gently dissected from the kidney, disturbing it as little as possible to achieve access to the renal pelvis. E) Pressing on the right side of the left kidney to better visualize the renal pelvis, the syringe is held with the thumb on the depressor and the needle is carefully but firmly placed into the renal pelvis. F) Following the <3 s injection, clearing may be observed in the areas of the kidney that received the bulk of the injection. G) Sterile purple vicryl absorbable sutures are used to make 2-4 independent knots in the muscle layer. H) Sterile blue nylon non-absorbable sutures are used to make 2-4 independent knots in the skin layer. Please click here to view a larger version of this figure.
Cause | Onset | Number of mice affected | Symptoms | Immediate action | Long-term solution | ||
DNA contained endotoxins | 6-40 h post-injection. | Likely to affect every mouse given the contaminated DNA preparation. | Breathing problems, signs of severe pain, organ failure, death. | Euthanize the affected mice. Test each injected component for endotoxins with Limulus Amebocyte Lysate. | Use endotoxin-free maxiprep kits and only sterile and new or NaOH-treated labware. Use a commercially purchased, endotoxin-tested buffer to dilute the DNA. | ||
Anesthesia overdose | During surgery, either before or after DNA injection. | May affect only the younger, smaller, or leaner mice. | Cease respiration while on heating pad, urination. | Decrease anesthesia for remaining mice. | Double-check preparation and protocol for dosage accuracy. Exclude “runts.” Consult veterinarian if appropriate dose was used. | ||
Air bubble in the syringe or needle used for injection | Immediately following renal pelvis injection. | Unless all syringes were loaded carelessly, this only affects one mouse. | Gasping for breath. | Check remaining syringes for signs of air bubbles. | Prepare syringes carefully, tapping the syringe to remove bubbles at the bottom. Improve lighting conditions to better visualize bubbles. | ||
Opening of surgical site | 12-72 h post-injection | One or more mice. Sometimes the entire cage. | Gaping wound, usually no other distress | Repeat suturing to repair the wound carefully under isoflurane anesthesia with sterile technique. May need to irrigate with saline or remove wound borders with scissors. | If all mice have very short or missing sutures, there may be one mouse removing them, so mice may be separated. Improve suturing technique. Use independent knots. | ||
Hernia at surgical site | 48+ h post-injection | One or more mice. | Mound is visible at surgical site. | Under isoflurane anesthesia with sterile technique, cut the healed skin to reveal hernia of muscle layer. Replace organs in the peritoneum, repair muscle layer with absorbable sutures and close the site. | This indicates poor suturing of muscle layer. Improve suturing technique. Use independent knots. | ||
Kidney failure | 48+ h post-injection | One or more mice. | Weight loss of >20%, possibly becoming uremic, hunched posture | Provide saline and increase or prolong analgesia. If no improvement is observed, sacrifice the affected mice. | Alter the disease state of the animal to make it less severe. Alter the transgene to be less strong or inducible. Inject mice at an earlier timepoint in disease progression. | ||
Abcess or infection | Days to weeks following surgery | One or more mice. | Palpable abcess or signs of sepsis | Euthanize the affected mice. Request necropsy to confirm suspected infection. | This may occur when the surgical conditions and injections are not sufficiently sterile. The procedure shown is for mice with a normal immune system but further precautions must be taken in the setting of immunocompromised animals such as those treated with cyclophosphamide. |
Table 1. Table of potential health problems encountered during the renal pelvis injection protocol. Although the listed health problems are not common, there are a number of investigator-related errors that can occur during the course of the procedure. This table may be of aid in prevention and diagnosis of the health problems, as well as for implementation of potential remedies to prevent such problems from occurring in the future. With practice, investigators should expect infrequent health problems and mortality due to the procedure.
3. Assess injection efficiency and transgene effects
The surgery and injection technique are simple to perform once mastered, requiring no major equipment or expensive materials. If new to flank-incision kidney surgery, one training day on several mice scheduled for euthanasia should be allowed in which the mice are not recovered following surgery because the first attempt at this surgery may take much longer than normal. Alternatively, investigators familiar with similar techniques may find it quite simple. Carefully follow the illustration in Figure 1 and the pictorial directions in Figure 2. To properly place the incision, the surgeon can use fingers to push on the mouse's stomach while it lays on its side. The kidney is visualized by a small lump that rises near the spine with gentle abdominal pressure, and the incision should be made over this area (Figure 1 and Figure 2). It is important to make the incision through the skin and muscle layers the correct size (Figure 1). If the incision is too big, the kidney will glide back into the abdominal cavity easily, making injection difficult. If the incision is too small, it will be difficult to push the kidney out of the abdominal cavity and organ damage could occur. Another important point is placement of the needle in the renal pelvis. The surgeon should push the kidney down with closed forceps, then insert the needle into the renal pelvis at an angle parallel to the working surface such that most of the injection volume goes directly into the kidney parenchyma (Figure 2). It is important that the injection be performed as quickly as possible to induce the hydrodynamic pressure required for kidney transfection.
Mice of different strains, ages, weights, and/or sex will have differences in the position of the kidney and amount of fat surrounding the kidney, so some adjustments may be necessary to work with the mice of interest. If possible, the first surgical attempt from which mice are recovered should include an immediate readout of kidney-specific gene expression, such as luciferase imaging (Figure 3A). If young mice are used, the expected result is that all mice will have a radiance that is within one order of magnitude (Figure 3B). Occasionally, even after a second injection of luciferin, it is apparent by imaging that a mouse or mice failed to become transfected at a level comparable to the others or that some injected mice are even completely lacking in gene expression. Such mice may be excluded from the study due to failed transgene delivery. If many of the mice lack gene expression, then the surgeon should decrease the time of injection of the DNA solution. Failure to induce the necessary damage through the high pressure of the injection is the most likely culprit for consistent lack of gene expression. Mice should be carefully monitored following surgery for potential health problems (see Table 1 for potential health problems with accompanying solutions and consult with the institution's veterinary team as necessary). Following an initial training period, the surgery itself should take less than ten min. The surgical period should be made as short as possible by operating on one mouse at a time to prevent desiccation of tissues during surgery. Most of the time investment in this procedure is due to the induction of and recovery from the ketamine/xylazine anesthetic. Two experienced personnel working as a team should expect to perform up to ten surgeries per day, with fifteen as a reasonable maximum.
Researchers should expect a low number of cells to stain highly positive for the transgene within the first few days following the injection. These positive cells will be localized in patches surrounding areas where the injection fluid infiltrated the kidney (Figure 3C-D). Some parts of the kidney will be completely negative for transfected cells. Such negative areas will also be lacking in erythrocytes and have completely normal histology since the injection did not reach these areas. Staining protocols and promoters should be optimized carefully to capture gene expression. Investigators should expect that the cell types transfected will be variable, the gene expression levels within the low percentage of cells transfected are high, and the localization of the proteins within the cells may not be as expected due to high levels of expression (Figure 3D).
Figure 3. Hydrodynamic renal pelvis injection results in kidney-specific gene expression of luciferase and TdTomato. (A) FVB 7-8 week old male mice were given hydrodynamic renal pelvis injections of 20 µg of pTEeL, a plasmid expressing enhanced firefly luciferase from the EF-1α promoter. Mice 2, 3, and 5 were also injected with 20 µg of fluorescent reporter plasmid. (B) Dot plot of the radiance obtained from the mice shown in (A). (C and D) Mice were given either hydrodynamic delivery buffer alone (C) or (D) fluorescent microspheres (bright green dots) and pEF1α-TdTomato expressing the red fluorescent reporter TdTomato from the EF-1α promoter. Proximal tubules were stained by Lotus tetragonolobus agglutinin (LTA; green) and transfected cells are shown after staining with anti-RFP antibody to amplify the TdTomato signal (red). Nuclei are stained by DAPI (blue). The images shown in (C) and (D) are 13.2 mm x 17.5 mm. Error bars represent the standard error of the mean. Please click here to view a larger version of this figure.
In this protocol a robust method for achieving reproducible gene expression specifically in the kidney is described. In the hands of a moderately experienced surgeon we have found the percentage of mice transfected by this technique to be in the range of 50-100%, depending on mouse age and the sensitivity of the readout of the transgene. The level of luciferase gene expression was above background for several months in mice receiving piggyBac transposons and completely maintained for several weeks in immunocompromised mice receiving the same transposons11. While many cells stained brightly for the transgene in areas near the injection site, the overall transfection efficiency was relatively low (Figure 3D). The most important material to achieve the desired outcome is the injection syringe itself. This must be a 29G x ½" insulin syringe with an attached needle and no safety to impede the injection. The injection must be fast, within three s. The plasmid DNA preparation should be pure, with very low endotoxin and bacterial genomic DNA levels19.
It is conceivable that injections of other substances, such as RNA, viruses, or protein, may be done in a similar manner. The renal pelvis was superior as a route of entry to other injection sites tested, such as the renal vein or direct injection of the kidney parenchyma. Those injections led to bleeding and lower gene expression11. Because the renal pelvis is an elastic structure sometimes required to hold urine backflow, it may be better able to stretch and accommodate the needle. Leakage of DNA solution or urine from the hole created by the insulin syringe has not been observed, suggesting that the renal pelvis is repaired quickly. The exact mechanism by which the injected fluid transfects the kidney has not been studied in depth as it has for the liver, for example. Future electron microscopy studies may elucidate this mechanism further. The retrograde nature of the injection has precedence in that the hydrodynamic tail vein injection method reverses the flow of the hepatic vein20 and other successful methods of kidney transfection have injected via the renal vein12,13,14,21. Kidney damage is likely required for optimal transfection efficiencies so we hypothesize that transfection occurs via damage to the cell membrane caused by the fast, high-volume injection. The podocytes, tubules, and collecting ducts all have direct interface with the urinary space as their major function is to filter and concentrate the urine.
Although reproducible and kidney-specific, this method is limited by the relatively low transfection efficiency observed (Figure 3D). Hydrodynamic renal pelvis injection is not a substitute for creation of a transgenic animal driving the transgene from a kidney-specific promoter, in which every cell of a certain subtype in the animal would express the gene of interest. Rather this method is best adapted to gene transfer approaches for diseases in which a small number of corrected cells would have a dramatic effect on the phenotype, for secretion of hormones or other substances normally produced by the kidney, or for creation of a population of rare cells that will exert some positive effect on the kidney to repair or regenerate it. Alternatively, it could also be used in the opposite way to develop a new model of kidney injury by introducing a transgene with a negative influence.
The authors have nothing to disclose.
A Career Development Award from the Department of Veterans Affairs [BX002797] supported L.E.W. and the National Institutes of Health [R01-DK095867] and American Heart Association [15GRNT25700209] supported J.C. The National Institutes of Health [DK093660], Department of Veterans Affairs [BX002190], and the Vanderbilt Center for Kidney Disease supported M.H.W. This material is the result of work supported with resources and use of facilities at the VA Tennessee Valley Healthcare System.
AnaSed Xylazine | Patterson Veterinary | 07-808-1947 | Anesthetic – Not controlled substance |
BD Insulin Syringe 0.5 mL 29G 1/2 Inch | Cardinal Health | 309306 | Required syringes |
Buprenex | Pharmacist/Veterinarian | Analgesia – Controlled Substance | |
Dynarex Disposable Towel Drape | Thermo Fisher Scientific | 19-310-671 | Place over heat pad |
EndoFree Plasmid Maxi Kit | Qiagen | 12362 | Use only endotoxin-free plasmid DNA |
Endosafe Gel-Clot LAL Rapid Positive Control | Charles River | PC200 | Positive control for endotoxin test |
Endosafe Gel-Clot LAL Rapid Single Test Vial | Charles River | R13500 | Endotoxin test |
Extra Fine Micro Dissecting Scissors | Roboz Surgical Instrument | RS-5882 | Surgical tool |
Fisherbrand Instant Sealing Sterilization Pouch – 9" | Thermo Fisher Scientific | 01-812-51 | For autoclaving surgical tools |
Gaymar Heat Pump | Paragon Medical | TP-700 | Water-circulating heat pump |
Germinator 500 | Roboz Surgical Instrument | DS-401 | To reuse surgical tools during surgery |
Graefe Forceps | Roboz Surgical Instrument | RS-5136 | Surgical tool |
Graefe Tissue Forceps | Roboz Surgical Instrument | RS-5153 | Surgical tool |
Halsey Needle Holder, 5" Length | Roboz Surgical Instrument | RS-7841 | Surgical tool |
Heat pads – 15" x 21" – need at least 3 | Paragon Medical | TP22G | For use with Gaymar Heat Pump |
IsoFlo (Isoflurane, USP) | Abbott Animal Health | 5260-04-05 | For imaging and euthanasia |
Isotec Isoflurane Delivery System Vaporizor | Smiths Medical | VCT3K2 | For imaging and euthanasia |
Ketamine | Pharmacist/Veterinarian | Anesthetic – Controlled Substance | |
Kimwipes | Kimberly-Clark Professional | 34120 | Laboratory tissues |
Living Image software | Caliper Life Sciences | For live animal imaging | |
Luciferin | Perkin Elmer | 122796 | For live animal imaging |
Nanodrop 2000 | Thermo Scientific | ND-2000-US-CAN | Spectrophotometer for DNA measurement |
Prevantics Swabs | Thermo Fisher Scientific | 23-100-110 | For skin surgery prep |
Prolene 5-0 sutures Taper 30" | Thermo Fisher Scientific | NC0256891 | Non-absorbable sutures for skin |
Puralube Brand Opthalmic Ointment | Patterson Veterinary | 07-888-2572 | To keep eyes moist during surgery |
Trans IT – QR Hydrodynamic Delivery Solution | Mirus Bio | MIR-5240 | Hydrodynamic delivery buffer for diluting DNA |
Vicryl 5-0 Sutures J303H | Thermo Fisher Scientific | NC9816710 | Absorbable sutures for muscle layer |
Wahl Mini Arco Clipper | Med-Vet International | 8787-1550 | Shaver for skin prep |
Xenogen IVIS 200 | Caliper Life Sciences | For live animal imaging |