A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice

Podsumowanie

This article describes modifications of a procedure to implant a peritoneal dialysis catheter in a murine model to avoid major technical issues observed with the conventional techniques.

Streszczenie

Murine models are employed to probe various aspects of peritoneal dialysis (PD), such as peritoneal inflammation and fibrosis. These events drive peritoneal membrane failure in humans, which remains an area of intense investigation due to its profound clinical implications in managing patients with end-stage kidney disease (ESKD). Despite the clinical importance of PD and its related complications, current experimental murine models suffer from key technical challenges that compromise the models' performance. These include PD catheter migration and kinking and usually warrant earlier catheter removal. These limitations also drive the need for a greater number of animals to complete a study. Addressing these drawbacks, this study introduces technical improvements and surgical nuances to prevent commonly observed PD catheter complications in a murine model. Moreover, this modified model is validated by inducing peritoneal inflammation and fibrosis using lipopolysaccharide injections. In essence, this paper describes an improved method to create an experimental model of PD.

Wprowadzenie

End-stage renal disease burden
Chronic kidney disease (CKD) is a worldwide health problem¹. Current estimates suggest that more than 850 million people worldwide have kidney disease. The prevalence of kidney disease almost doubles the number of people with diabetes (422 million) and is more than 20 times the prevalence of cancer (42 million) or HIV/AIDS (36.7 million) patients worldwide². Approximately one in seven Americans have CKD, and two in 1,000 Americans have ESKD requiring a kidney transplant or dialysis support³. Considering the escalating burden of ESKD worldwide, optimizing dialysis technology is crucial³.

Peritoneal dialysis
PD is a significantly underutilized modality for the treatment of ESKD in the United States. According to the United States Renal Data System (USRDS), the percentage of prevalent PD patients was only 11% in 2020^4,5. PD confers several advantages over in-center hemodialysis (HD), including a better quality of life, fewer clinic visits, and a decrease in Medicare expenditures^6,7. Additionally, PD is a home-based therapy and is associated with a much lower risk of severe infections such as bacteremia and endocarditis that are often related to hemodialysis catheters. Furthermore, PD can be initiated rapidly with an urgent start protocol, decreasing the need for dialysis initiation with indwelling vascular catheters⁸. PD is considered the preferred method of dialysis in the pediatric ESKD population⁹.

Peritoneal impairment induced by peritoneal dialysis
PD entails introducing PD fluid (dialysate) into the peritoneum, which results in inflammation and remodeling of the peritoneal membrane over time. Peritoneal inflammation triggers fibrosis, culminating in the potential loss of ultrafiltration capabilities of the membrane over time. Preservation of the peritoneal membrane is a significant challenge in PD, and further research is critically important to ensure that best clinical practices are available to practitioners. There are well-established murine models to help further the understanding of pathophysiological mechanisms of peritoneal infection and inflammation, solute, water transport kinetics, and membrane failure; still, technical issues with the catheter often limit these models¹⁰.

Analyzing the peritoneal membrane changes
In ESKD patients, dialysate is traditionally introduced in the peritoneal cavity through a Tenkhoff catheter with a deep and superficial cuff. The patients can potentially experience catheter-related complications, including catheter migration, infusion pain, and poor drainage of the dialysate^11,12,13. Two major types of peritoneal catheters have been introduced for humans, coiled or straight, to minimize these complications¹². Several modifications, including an extra cuff to the conventional two-cuffed catheters, have been added to the original catheters to prolong PD catheter survival¹¹. The insertion technique varies according to several factors by preventing catheter migration to be added after survival, including the availability of the resources and the level of expertise¹⁴.

In contrast, the murine models of peritoneal dialysis have fundamental differences in techniques and purpose compared to human peritoneal catheters. For example, peritoneal catheters in murine models are used primarily to study membrane alterations and are less intended for bidirectional drainage functions. The current technique suffers from potential port dislodgement and catheter migration due to the handling of the animals. In the conventional murine models, the access ports were not fixed to the skin. This aspect created an unstable access port, which in awake animals might get dislodged, resulting in catheter migration. Given the importance of murine models in peritoneal membrane research, it is imperative to create effective surgical techniques to generate reliable models. Therefore, we set out to optimize the conventional model of PD catheter placement. It is important to note that the catheter itself causes histopathologic alterations in the peritoneal membrane, and, thus, any conclusions regarding the effect of PD solutions in animal studies must be interpreted in the context of the PD catheter as a foreign body^15,16,17.

Peritoneal membrane histopathology
PD failure is mainly related to fibrosis and excess angiogenesis resulting in the loss of an osmolar concentration gradient. In addition, the peritoneal membrane filtration capacity might be affected by peritonitis. In addition, infectious peritonitis is a well-established cause for change in the dialysis modality from peritoneal dialysis to hemodialysis.¹⁸.

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Protokół

For this study, eight female C57BL/6J mice, 8-12 weeks in age, and an average weight of 20 g were used. The mice were housed under standard conditions and were fed with chow and water ad libitum. This study was performed with the approval of the Institutional Animal Care and Use Committee (IACUC), Boston University Medical Center (AN-1549). The procedures described here were performed under sterile conditions.

1. Anesthetize the mouse in an Isoflurane chamber, and inject the analgesic subcutaneously

1. Hold the animal from the base of the tail. Keep the animal on the dorsum surface of the non-dominant hand.
2. Transfer the animal to the continuous anesthetic induction chamber filled with 3%-4% isoflurane. Confirm adequate general anesthesia by the absence of toe-pinch reflex in the right and left hind limbs. Keep the maintenance of the general anesthesia with Isoflurane 1%-3%.
3. Apply ophthalmic ointment on both eyes.
4. Administer a subcutaneous injection of Buprenorphine.
   1. Dissolve the stock of Buprenorphine at a concentration of 0.3 mg/mL in sodium chloride (NaCl) 0.9% to achieve the final concentration of 0.03 mg/mL.
   2. Inject a dosage of 0.05-0.1 mg/kg of 0.03 mg/mL Buprenorphine, together with 500 µL of sterile NaCl 0.9%, 20 min before surgery in a 20 g mouse (2 µg or 66 µL of 0.03 mg/mL Buprenorphine per mouse).

2. Skin preparation

1. Place the fully anesthetized mouse in a left lateral position, exposing its right flank to the heating blanket. Shave the right side of the abdomen, just close to the midline to the paraspinal area, and down to the animal's tail.
2. Disinfect the shaved area three times using a cotton swab with the alternate application of the antiseptic solution or scrub and either 70% alcohol or sterile saline in a circular motion, starting at the surgical incision site and moving outward. Discard the cotton swab after each use. Be careful not to excessively wet non-surgical areas of the animal with alcohol or antiseptic as this can worsen hypothermia.
   NOTE: It is important to properly dilute antiseptic solutions and not leave surgical scrubs on the skin during surgery, as they can be irritating and need to be rinsed off. Frequently check the temperature of the heating blanket during the procedure to ensure that the temperature does not fall.

3. Measure the catheter length and mark the insertion point within the abdomen and the tube tract over the prepared skin

1. Assign the access port pocket 1 cm above the animal's tail. Hold the installation segment with the non-dominant index and thumb finger over the assigned area near the tail.
2. Place the catheter above the skin and estimate the place for the catheter's tube insertion within the abdominal cavity. Mark the assigned place for tube insertion, respecting the minimal bending of the tube near the anterior midline.
   NOTE: All the procedures must be performed with sterile gloves, and the catheter should be kept sterile during the measurement. Surgical tools must be autoclaved at 121 °C before use. Refer to Supplemental Figure S1 for the instruments required for the procedure.

4. Customize the peritoneal catheter reservoir section

1. Punch a side hole over the frame of the reservoir section with the mouse ear tagger (Figure 1 and Figure 2). It should be noted that the ear punch is a surgical tool, and it needs to be sterile.

5. Place the instillation port

1. Make a horizontal 1 cm wide skin incision 1 cm above the tail. Bluntly dissect the subcutaneous plane from the underlying muscular layer to make a pouch for the catheter placement to ensure the instillation port resides in the ideal port pocket freely.
2. Keep the iris scissors' tip toward the midline to make an oblique tunnel for the tube placement (Figure 3A).
3. Pass the 3.0 suture from the customized side hole. Fix the access port to the muscular bed by tightening the passed suture, keeping the tubing course cephalad.

6. Make the catheter tip insertion site incision

1. Make a 1 cm incision over the formerly marked area near the midline. Confirm the well-developed tract by passing scissors through the tract.
2. Pick the catheter tip gently with forceps to place the catheter in a retrograde course.
   NOTE: Avoid pinching the side holes of the tube.
3. Pass the catheter tube through the prepared tract (Figure 3B). Make a 1 cm incision over the muscular layer close to the right midline.

7. Confirm the functioning of the catheter

1. Before closing all the incisions, make sure the placed catheter is functional. Check the function with a 1 mL syringe attached to the specific Huber needle for the port.
2. Inject 200 µL of normal saline into the instillation port. Look for a smooth flow with a zero tolerance for resistance.
3. Flush the port and catheter with 10% heparin to maintain patency.

8. Close the skin incisions

1. Close the skin incisions around the port reservoir (Figure 3C) with 3-0 absorbable sutures.

9. Fix the catheter tip inside the abdominal cavity

1. Place a loose purse-string suture with 4-0 round absorbable suture around the incised abdominal wall muscle. Pass the proximal felt of the catheter inside the incision.
2. Tighten the prepared purse-string suture around the tube while keeping the second felt outside the purse string, over the muscular layer (Figure 3D), and close the skin with 3-0 absorbable sutures (Figure 2).

10. Monitor the animals postoperatively and daily, administer postoperative analgesia and fluids, and maintain daily postoperative records for a minimum of 7 days and until complete recovery

1. Keep the catheter functional with a daily injection of 200 µL of normal saline through the catheter.

11. Fluid injections

1. Confirm the uneventful postprocedural process by carefully inspecting the skin incision.
2. Prepare LPS 2 mg/kg body weight for intraperitoneal injections (i.p.) by diluting 40 µg of the LPS with sterile phosphate-buffered saline (PBS) to the working concentration of 0.2 µg/µL (in essence, 10 µL for 2 µg/g body weight and 200 µL of LPS for 20 g mice).
3. Start the injections in the second week following the catheter implantation.
   1. Hold the animal gently with the non-dominant hand and restrain the instillation port while moving the index and thumb fingers in the cephalad direction.
   2. Disinfect the skin overlying the reservoir with 70% isopropyl alcohol. Use the syringe attached to the Huber needle to inject the LPS.
      1. After entering the port with the Huber needle, inject 100 µL of normal saline into the port to confirm the patent course.
      2. Inject the prepared 200 µL of LPS, followed by the 100 µL of normal saline for tube irrigation, and make sure there is no resistance.

12. Anesthetize the mice before harvesting the peritoneum and collect the peritoneal fluid

1. Following 7 days of LPS injections and 2 weeks of catheter implantation, plan for the peritoneal biopsy.
2. Plan for general anesthesia.
   1. Anesthetize the mouse in an isoflurane chamber and inject the analgesic subcutaneously.
   2. Hold the animal from the base of the tail, and keep the animal on the dorsum surface of the hand.
   3. Transfer the animal to the continuous anesthetic induction chamber filled with 3%-4% isoflurane. Confirm adequate general anesthesia by the absence of toe-pinch reflex in the right and left hind limbs. Keep the maintenance of the general anesthesia with isoflurane 1%-3%.

13. Peritoneal biopsy

1. Place the animal on the heated blanket in the supine position. Make a midline skin incision from the sub-xiphoid to the bladder.
2. Perfuse the subfascial plane with cold PBS (Figure 3E).
3. Make sure the plane is completely dissected without disturbing the integrity of the peritoneum. Start dissecting the peritoneum from the lateral peritoneal reflection at the left lower quadrant, starting from the hilum to the left flank, and bladder in the lower aspect to keep samples consistent between animals (Figure 3F).
4. Following the peritoneal harvest, euthanize the animal by cervical dislocation.

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Wyniki

All the implanted catheters were functional till the end of the study, and catheter dislodgement or kinking did not complicate any of the implanted catheters. The current, modified technique was further validated with a peritonitis-induced model using LPS. The control mice received 200 µL of daily normal saline injections, while the experimental mice were injected with 200 µL of LPS, as discussed in protocol step 11, for a total of 7 days following catheter implantation.

The peritone...

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Dyskusje

Three murine models of PD are described. This includes a blind puncture of the peritoneal surface, an open-permanent system, and a closed system¹⁰. The blind puncture of the peritoneal surface involves direct peritoneal access similar to intraperitoneal injections but does not allow drainage of dialysate. Being a blinded procedure, this method can injure the abdominal visceral organs. The open-permanent system model keeps the dialysis catheter and instillation port outside the body. However, this ...

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Materiały

Name	Company	Catalog Number	Comments
10% heparin	Canada Inc., Boucherville, QC, Canada)	Pharmaceutical product
Buprenorphine 0.3 mg/mL	PAR Pharmaceutical	NDC 42023-179-05
C57BL/6J mice	The Jackson Lab	IMSR_JAX:000664
CD31	Abcam	Ab9498
Clamp	Fine Science Tools	13002-10
Forceps	Fine Science Tools	11002-12
Dumont #5SF Forceps	Fine Science Tools	11252-00
Dumont Vessel Cannulation Forceps	Fine Science Tools	11282-11
Fine Scissors - Large Loops	Fine Science Tools	14040-10
Fisherbrand Animal Ear-Punch	Fisher Scientific	13-812-201
Hill Hemostat	Fine Science Tools	13111-12
Huber point needle	Access  technologies	PG25-500	Needle for injections
Isoflurane, USP	Covetrus	NDC 11695-6777-2
Lipopolysaccharide from E.coli	SIGMA	L4391
Microscope	Nikon Eclipse Inverted Microscope	TE2000
Minute Mouse Port 4French with retention beads and cross holes	Access  technologies	MMP-4S-061108A
Posi-Grip Huber point needles 25 G x 1/2´´	Access  technologies	PG25-500
Scissors	Fine Science Tools	14079-10
Vicryl Suture	AD-Surgical	#L-G330R24