A Mouse Model to Evaluate the Long-Term Structural and Functional Outcomes after the Reversal of Prolonged Unilateral Ureteric Obstruction

Abstract

Clinical management of patients with urinary tract obstruction (UTO) requires early intervention to reverse the cause of the obstruction, but despite this, patients with prolonged UTO are at increased risk of chronic kidney disease (CKD) and recurrent acute kidney injury (AKI). However, other than an early reversal of obstruction and generic therapy to delay CKD progression, no therapies have significantly improved long-term renal outcomes after the reversal of UTO. To address this, a number of laboratories have developed models of reversible UUO (R-UUO), but these are technically challenging and have not been widely adopted. In addition, while mouse models of R-UUO are attractive as they can be used to harness the power of mouse genetics to study disease pathophysiology, these have been particularly challenging as methods used often lead to irreversible UUO if the obstruction lasts >3 days. In addition, because of the nature of these models, few studies have evaluated the long-term functional outcomes of reversing ureteric obstruction. To address this, we recently developed a mouse model of R-UUO with delayed contralateral nephrectomy that allows for the analysis of the long-term effects of reversing prolonged UTO on long-term renal structural and functional outcomes. These studies show that despite near complete histological recovery 3 months after reversal, there was a permanent reduction in renal function and a marked and persistent defect in urinary concentrating capacity, indicative of a defect in renal medullary function. The model requires three major survival surgeries but results in a robust and reproducible long-term reduction in renal function that can be reproduced in different mouse strains by adjusting the period of obstruction. In this article, we provide detailed instructions for performing these surgeries, optimizing conditions for use in different mouse strains, evaluating renal functional outcomes, and harvesting renal tissues.

Introduction

Urinary tract obstruction (UTO) may be unilateral, which is often asymptomatic, presenting late with chronic obstructive uropathy¹, or may present acutely with renal colic and/or hematuria; acute pyelonephritis or post-renal acute kidney injury (AKI)². Key to the management of UTO is early reversal of obstruction². However, despite effective reversal and the use of generic therapies to delay the progression of chronic kidney disease (CKD), affected patients remain at increased risk of progressive CKD and hypertension^3,4. In addition, long-term urinary concentrating capacity (UCC) is often impaired⁵, which may predispose susceptible patients to dehydration and recurrent AKI, accelerating the progressive decline in renal function after UTO.

In part, this failure to identify therapeutic approaches to improve long-term outcomes in patients with UTO has resulted from our reliance on the use of irreversible unilateral ureteric obstruction (UUO) models to explore the pathobiology of UTO. However, these models do not replicate the clinical situation in which patients undergo reversal soon after diagnosis and cannot be used to explore the mechanisms and therapeutic approaches that could be used to improve long-term repair after the obstruction is reversed. Reversible UUO (R-UUO) models have been developed to address this, but these are more technically challenging than the irreversible UUO models, more difficult to reproduce by different laboratories, and, as a result, have not been as widely adopted by the scientific community. Mouse models are attractive since they can be used to harness the power of mouse genetics to study disease pathophysiology, but they have been particularly challenging to optimize as the methods often lead to irreversible UUO if the obstruction lasts more than 3 days⁶. A variety of approaches have been deployed to address this, including multi-surgery, sequential placement of clamps along the ureter every 2 days^6,7; UUO followed by bladder reimplantation^8,9,10,11; and use non-damaging clamps and tubing to obstruct the ureter^12,13,14. Long-term outcome studies in rats and mice have shown improved renal function, reduced fibrosis, and/or glomerular injury after reversal of obstruction, but only partial recovery if UUO lasts longer than 2 days^{6,7,8,9,12,14,15,16,17,18,19}. In addition, while the renal medulla (RM) is particularly susceptible to damage caused by UTO^20,21 until recently, there have been no long-term rodent studies in which the effects of prolonged R-UUO on RM structure and function have been studied. This is significant since UCC and the ability to excrete salt loads without increasing blood pressure (the so-called pressure natriuresis response) are dependent on maintaining anatomic integrity of the RM^22,23,24, so that incomplete repair of the RM after reversal of UTO may explain why patients have increased susceptibility to recurrent AKI and hypertension after reversal of UTO. However, many of the techniques required to evaluate this require multiple rounds of surgery (five major surgeries for 6 days of obstruction^6,7), require surgical expertise that is difficult to teach (e.g., surgical reimplantation of the ureter into the bladder^8,9,10,11), and because obstruction is unilateral, make it challenging for investigators to evaluate changes in renal function after reversal of the obstruction.

To address this, we have developed a mouse model of R-UUO that is relatively simple to perform and teach, requires a more limited number of surgeries than other techniques (two surgeries for 5 to 7 days of obstruction), and allows for the analysis of renal functional recovery without the need for invasive and/or expensive split renal function studies²⁵. For this, mice undergo three major surgeries: 1) placement of a non-traumatic vascular clamp on the proximal left ureter; 2) removal of the vascular clamp 5 to 7 days later, depending on the mouse strain; and 3) removal of the contralateral kidney 10 days later to evaluate renal function. Approximately 20 to 40% of mice die within 2–3 days of the nephrectomy, indicating that the UUO does not reverse in all mice. Long-term follow-up of the surviving mice shows that despite the near complete histological recovery of the renal medulla (RM) 3 months after R-UUO. There is reduced renal function, measured by transdermal glomerular filtration rate (tGFR)²⁶. In addition, there is a marked reduction in UCC, determined by measuring urinary osmolality after water restriction, suggesting there is a permanent defect in RM function. Time course studies show that while there is improvement in tGFR between days 28 and 56 post-RUUO, this stabilizes between days 56 and 84. In contrast, there is no improvement in UCC between days 28 and 84 after R-UUO²⁵. Using scRNA-Seq of isolated RMs, validated by cell lineage and immunohistochemistry studies, we identified strong regenerative responses that restored RM dimensions after R-UUO but that all of the major cellular compartments in the RM showed permanent changes in cell numbers and gene signatures that are likely to impact functional recovery after R-UUO. This included persistent proinflammatory responses in the RM collecting duct and loop of Henle cell populations 84 days after R-UUO, similar to the inflammatory and senescence signatures of failed repair proximal tubular epithelial cells after AKI and UUO^27,28,29,30. Similar changes were also seen in RM collecting ducts from patients with recurrent renal stone disease³¹, suggesting there is a common injury response to RM damage in both humans and mice. In this article, we provide detailed instructions on how we perform these surgeries, how we optimize conditions for its use in different mouse strains, how we determine whether the ureter is still obstructed after reversal, how we evaluate key functional outcomes, and how we harvest renal tissues to assess the renal medullary structures.

Protocol

Here, the four-step protocol used for long-term studies is described (Figure 1). As discussed at the end of this manuscript, researchers may choose only to perform three of these (sections 1, 2, and 4) or all four if interested in studying long-term outcomes. All mouse experiments in this manuscript have been approved by the Vanderbilt Institutional Animal Care and Use Committee and adhere to the animal use guidelines.

Figure 1: Long-term decrease in renal function and urinary concentrating capacity after R-UUO.

Male BALB/c mice underwent R-UUO for 7 days, followed by a contralateral nephrectomy (Nx) 10 days after reversal of UUO. (A) Typical study design for long-term studies; (B) Survival curves; (C-E) Functional studies 84 days after R-UUO comparing R-UUO/Nx with Nx alone. BUN (C), Transdermal GFR (D), and urinary osmolality from spot urine samples obtained after 18 h of water deprivation (E). Individual data points shown with means ± SEM. T-Test, p values shown if significant.

1. Placement of the ureteric clamp (10 – 15 min/mouse)

NOTE: Rely on the knowledge of mouse kidney anatomy to locate the ureter since it is not possible to see it directly without the surrounding connective tissue. The ureter runs along the lower posterior aspect of the hilar tissue as the kidney is exposed (Figure 2). The choice of vascular clamp is important. These can be expensive but can be cleaned, autoclaved, and reused multiple times. These clamps can be reused at least 100 times without issues.

1. Autoclave all surgical instruments before surgery at 121 °C for 30 min. If performing surgeries on multiple mice, remove blood and debris from instruments between use and sterilize for 10–15 s using the hot bead sterilizer. Allow the instruments to cool before using them again.

2. Weigh the mouse.

3. Anesthetize the mouse using 5–10 mg/kg Xylazine with 90–120 mg/kg Ketamine by intraperitoneal injection (see Table of Materials). This usually takes 3–5 min to take effect.

4. Administer pre-operative analgesia if using a different anesthetic agent, such as isoflurane, as, unlike Xylazine, this does not have analgesic effects.

5. Shave the surgical site from the hips to the rib cage and around the side, with enough border to keep the hair from contaminating the incision site. Perform in a preparatory area, not where the surgery is performed.

6. Cover the heated surgical field with a sterile surgical drape. It will help prevent stray hairs from entering the surgical field and provide an area to lay sterile instruments.

7. Apply ophthalmic lubricating ointment to the eyes.

8. Apply aseptic prep using betadine-soaked gauze: scrub from the center of the site toward the periphery 3 to 4 times, each time followed by wiping once with an alcohol swab. Remove all betadine from the surgical field prior to incision.

9. Make a 1.5 cm longitudinal dorsal incision on the midline through the skin and subcutaneous layers using scissors and forceps.

10. Make a small incision through the left flank muscle and fascia above the kidney and exteriorize the left kidney.

11. Carefully dissect the lower pole fat and some of the connective tissue around where the ureter should be. It is not possible to see the ureter directly without the surrounding connective tissue. Then, separate the ureter with its surrounding connective tissue from the renal pedicle, which contains the renal vein and artery, so the renal pedicle is not included with the ureteric clamp (Figure 2).

12. Use the clamp appliers to open it and place it on the ureter directly below the pelvis. Use the markings on the vascular clamp to ensure they are placed with the same pressure on the ureter for each mouse (Figure 3). Do not apply, then remove the clamp to reposition, as this can damage the ureter.

13. Once the clamp has been placed, gently push the kidney with the clamp back into the retroperitoneal space using a saline-soaked sterile cotton swab.

14. Close the muscle layer using an absorbable suture.

15. Close the skin layer using skin clips.

16. Administer Buprenorphine 3.25 mg/kg analgesia, as outlined in the drugs section, and administer 0.5 mL of sterile normal saline subcutaneously to hydrate the mice.

17. Leave the mouse on the heated hard pad surgical field or transfer it to their home cage with a warmed isothermal pad under half of the cage until the mouse wakes up.

18. Return the mouse to the animal room.

19. Closely monitor the mouse and give additional doses of Buprenorphine for pain or discomfort for 48 to 72 h. Clinical signs of pain and distress that indicate the mice need additional analgesia include depression or other behavioral changes, abnormal appearance, or postures such as piloerection, hunched posture, lack of grooming and eating, or immobility.

Figure 2: Anatomy of the mouse left renal hilum.

Illustrating the position of the ureter relative to the hilum and lower pole fat pad from the front. The red arrow indicates the ideal position of the ureteric clamp, which needs to be distinguished from the renal pedicle illustrated with the suture encircling the renal artery and vein. This must not be included in the jaws of the clamp.

Figure 3: Vascular clamp.

Markers indicate positions of maximal (15 g) and minimal (5 g) pressure.

2. Removal of the ureteric clamp (10 – 15 min/mouse)

1. Follow clamp placement procedure steps 1.1–1.10 using the original skin and muscle incisions to begin clamp removal. Remove the wound clips before cleaning the surgical area. Shaving again is usually not necessary since hair has not regrown.

2. Due to developed tissue adhesions, use forceps to carefully locate the clamp in the retroperitoneal space without first exteriorizing the kidney. Use the clamp appliers to gently open the clamp while using the forceps to pull the tissue downwards from around the clamp head to remove the clamp.

3. Now, exteriorize the kidney so that the renal pelvis can be visualized to determine obstruction. The renal pelvis should be swollen, indicating hydronephrosis.

4. Once the kidney is inspected, gently push the kidney back into the retroperitoneal space using a saline-soaked sterile cotton swab.

5. Follow steps 1.13–1.18, as outlined above.

3. Contralateral nephrectomy (10 – 15 min/mouse)

1. Shave the area if there is there is hair growth.

2. Use 1–5% isoflurane/air mix for anesthesia to allow an easier recovery due to having prior anesthetic events. Remove wound clips before cleaning the surgical area.

3. Administer Buprenorphine analgesia pre-operatively (see Table of Materials).

4. Follow procedure steps 1.1–1.9 using the original skin incision to begin.

5. Make a small incision through the right flank muscle and fascia above the kidney and exteriorize the right kidney.

6. Hold the kidney gently with smooth, curved forceps and carefully dissect the upper and lower poles of the kidney free from surrounding tissue.

   NOTE: Tissue from around the upper pole contains the adrenal gland, carrying its own blood supply, which can be pulled off the kidney but obviously should not be removed from the mouse (see Figure 2).

7. After liberating the kidney from surrounding tissue, tie the 4–0 silk suture completely around the renal vessels and ureter using a double surgical knot. If left for ~30 s, the kidney goes dark. Hold the kidney with smooth, curved forceps, and remove the kidney by cutting distally to the knot with curved scissors.

8. Gently push the remaining renal pedicle back into the retroperitoneal space using a saline-soaked sterile cotton swab.

9. Follow steps 1.13–1.18, as outlined above.

4. Tissue collection

1. Anesthetize with 5% isoflurane/air mix to achieve a stable plane of surgical anesthesia prior to the terminal procedure.

2. Open the abdominal cavity to expose the left kidney

3. Inspect the kidney to see any swelling of the renal pelvis (see step 2.3).

4. If it looks like the renal pelvis is still swollen, insert a small gauge butterfly needle connected to a syringe containing 0.5 mL of a 1 in 5000 dilution of methylene blue in phosphate-buffered saline (PBS). If the ureter is patent, the blue solution will be visible as it travels from the renal pelvis into the bladder. If not, the renal pelvis will swell with blue solution, and it will not appear in the bladder.

5. To perfusion fix the kidneys, open the thoracic cavity and expose the heart.

   1. While the heart is still beating, insert a 23–25 G butterfly needle into the left ventricle, start an infusion of normal saline, and cut the right atrium or ventricle to replace blood volume with saline.

   2. After about a minute, ensure the kidneys are white. Continue the saline infusion for another minute or swap it with an infusion of 10% formalin solution for another 1–2 min to fix the tissues.

   3. At this point, the mouse will be dead; harvest the tissues. If the tissues are collected for RNA or protein studies, do not perfusion fix them.

6. Upon removal of the kidneys from the animal (whether they have been perfusion fixed or not), use a razor blade to cut up to three 2–3 mm thick sagittal slices through the center of the kidney (Figure 4). This gives representative sections that include the renal medulla and cortex and provides enough tissue for histology, protein, or RNA assays.

7. Go on to fix/freeze samples according to the study requirements.

Figure 4: Sagittal sections of the mouse kidney.

(A) External appearance of kidney slices and their allocation; (B) Typical sections showing cortex, outer medulla (OM), and inner medulla (IM).

Representative Results

For long-term studies, we generally evaluate renal function by measuring sequential blood urea nitrogen (BUN) and transdermal Glomerular Filtration Rates (tGFR) in conscious mice (refer to a previously published article describing this²⁶), in addition to measuring urinary concentrating capacity (UCC) by evaluating urinary osmolarity from spot urine samples collected after 18 h water deprivation. While we always perform BUN studies, these are most useful to assess the severity of injury at early time points after the nephrectomy (see discussion). However, for long-term studies, BUN (and serum creatinine) measurements are not sufficiently sensitive to detect subtle changes in renal function that occur. These are usually performed at different time points after the nephrectomy (Figure 1A). Analysis of UCC provides a simple measure of renal medullary function and usually shows urinary osmolarity values that are markedly lower than nephrectomy control mice at the same time points. Figure 1B–E shows typical survival curves, BUN, tGFR, and urinary osmolarity measurements 84 days after R-UUO.

For more detailed information, including studies optimizing ureteric clamp times in different mouse strains and histological changes in the renal cortex and medulla at different time points, refer to our previously published work²⁵.

Discussion

Compared to some of the approaches that have been used to study reversal of prolonged UUO in mice^{6,7,8,9,10,11,12,13,14}, the protocol described here is relatively straightforward and does not require a high level of prior surgical expertise. It involves a relatively small number of survival surgeries (maximum 3) and can be used to evaluate long-term functional outcomes. There are a number of critical steps that need to be addressed for this technique to be successful and reproducible. These include: 1) Consistent placement of the clamp at similar points along the ureter since there is evidence that increasing the distance of obstruction from the renal pelvis slows the progression of renal injury during the period of obstruction³². Using a dorsal approach to exteriorize the kidney, the simplest way to do this is to place the clamp close to the renal pelvis. If surgery is performed by laparotomy, clamps can be placed close to the bladder^8,11. However, investigators need to bear in mind that mice may then need longer periods of obstruction to induce a similar degree of injury. This may end up causing irreversible obstruction unless the investigator reimplants the ureter proximal to the obstruction into the bladder, as described^8,11. However, reimplantation of the ureter is technically challenging, and in our experience, laparotomies are more traumatic and require longer recovery times than the use of a dorsal approach to expose the kidney; 2) Pressure exerted by the clamp must be consistent between mice to ensure there is no leakage of urine across the obstruction. This requires that the ureter is clamped at the same point between the clamp arms (Figure 3) and that a similar amount of connective tissue is left around the ureter in each mouse. Too much leaves more connective tissue, so the exerted pressure on the ureter is lower. Too little and the dissection runs the risk of damaging the ureter; 3) Removal of the vascular clamp can be challenging as it is easy to tear the tissue, which is usually thickened by the time the clamp is removed due to inflammation and fibrosis. This makes the tissue less easily malleable. This is particularly noticeable in FVB/N mice in which there was so much reactive fibrosis that it was virtually impossible to remove the clamp without tearing the ureter; 4) Timing of clamp removal is important and will depend on experimental needs. For example, if a study needs to induce survivable, severe injury that models the effects of prolonged urinary obstruction in humans, the ureter will need to be obstructed for long enough to induce injury but not so long that the ureteric obstruction is irreversible. Optimal survivable clamp times are also strain dependent²⁵, so this will need to be optimized for each mouse strain before starting definitive experiments. In general, clamp times range from between 5 and 7 days, although preliminary unpublished data suggest female mice may need even longer clamp times to have similar degrees of post-R-UUO injury. For this, we recommend that investigators perform initial pilot studies with ~5 mice/group and clamp times ranging from 5 to 9 days in increments of 2 days. They should evaluate macroscopic evidence of hydronephrosis at the time the clamp is removed (see Figure 3), observe the mice with measurement of BUN 24 h and 72 h after nephrectomy, and euthanize the mice at 72 h to confirm that the hydronephrosis has resolved (see section 4). If >60% of the mice survive, there is a transient increase in BUN at 24 h and 72 h after the nephrectomy (increases from baseline levels of 15–20 mg/dL up to 40–80 mg/dL); this is probably a good clamp time to stick with for experiments. Ultimately, the investigator will need to confirm that these mice develop long-term renal dysfunction by evaluating BUN and/or serum creatinine (which are usually insufficiently sensitive to detect long-term changes in renal function), tGFR, and UCC. Loss of the UCC compared with nephrectomy control mice provides a simple but sensitive measure of renal medullary dysfunction after R-UUO using this model.

An advantage of performing the contralateral nephrectomy in this protocol is that renal function tests can be performed at later time points without the need for more invasive and usually less sensitive, split renal function tests. In addition, the contralateral nephrectomy allows investigators to identify mice in which the ureter is patent since most of the mice with persistent obstruction die within 1–3 days of the nephrectomy, and those with partial obstruction have much higher BUN and lower GFRs than the others. This is a factor to consider if R-UUO studies are performed without a contralateral nephrectomy. For example, a recent study found that mice have reduced sodium clearance and develop salt-sensitive hypertension 4 weeks after R-UUO³³, suggesting that these mice have a pressure natriuresis defect that results from abnormal RM function in the R-UUO kidney²⁴. However, a caveat to this is that it is difficult to determine the extent to which the UUO was fully reversed in this model, as mice did not undergo a contralateral nephrectomy, so it is possible the pressure natriuresis defect is the result of persistent obstruction after the reversal.

However, a limitation of this protocol is that there is a 20%-40% mortality when R-UUO is combined with a contralateral nephrectomy, increasing animal losses and the need to include larger numbers of animals for a given study. Another limitation is that the contralateral nephrectomy increases blood flow and GFR in the remaining kidney^34,35, and compensatory cellular hypertrophy that occurs in response to the associated reduction in nephron mass, complicates the interpretation of the data³⁶. For this reason, if a contralateral nephrectomy is performed, investigators should include nephrectomy controls that are followed up for the same time period to control for compensatory responses that occur after nephrectomy. An alternative approach is to perform the contralateral nephrectomy 1 to 3 days before euthanizing the mice. This has been used to look at the long-term impact of unilateral ischemia reperfusion-induced AKI (IRI-AKI) on renal function³⁷. This allows the assessment of renal function but without sufficient time for the remaining kidney to undergo hypertrophy. The caveat to this is that there are marked changes in GFR and blood flow in the remaining kidney in the first 1 to 5 days after the procedure^34,35. This has profound effects on renal function, so minor differences in the timing of post-nephrectomy studies can have marked effects on measured renal function.

This protocol provides a tool to evaluate long-term tissue repair and functional outcomes after R-UUO. Because the model allows investigators to perform detailed functional assessment of renal recovery, future applications of this technique may include it use to explore the functional impact of therapeutic interventions on tissue repair in different cellular compartments after R-UUO. In mice, this may also include the use of genetic tools to identify new therapeutic targets that could be used to improve clinically important functional parameters after reversal of obstruction and to evaluate the effect of cell-specific gene targeting on long-term functional repair after R-UUO.