Minimizing Post-Infusion Portal Vein Bleeding during Intrahepatic Islet Transplantation in Mice

Summary

Here we present refined surgical procedures on successfully performing intraportal islet transplantation, a clinically relevant but technically challenging surgical procedure, in mice.

Abstract

Although the liver is currently accepted as the primary transplantation site for human islets in clinical settings, islets are transplanted under the kidney capsule in most rodent preclinical islet transplantation studies. This model is commonly used because murine intrahepatic islet transplantation is technically challenging, and a high percentage of mice could die from surgical complications, especially bleeding from the injection site post-transplantation. In this study, two procedures that can minimize the incidence of post-infusion portal vein bleeding are demonstrated. The first method applies an absorbable hemostatic gelatin sponge to the injection site, and the second method involves penetrating the islet injection needle through the fat tissue first and then into the portal vein by using the fat tissue as a physical barrier to stop bleeding. Both methods could effectively prevent bleeding-induced mouse death. The whole liver section showing islet distribution and evidence of islet thrombosis post-transplantation, a typical feature for intrahepatic islet transplantation, were presented. These improved protocols refine the intrahepatic islet transplantation procedures and may help laboratories set up the procedure to study islet survival and function in pre-clinical settings.

Introduction

Intraportal islet transplantation (IIT) via the portal vein is the most commonly used method for human islet transplantation in clinical settings. The mouse IIT model offers a great opportunity to study islet transplantation and test promising interventional approaches that can enhance the efficacy of islet transplantation¹. IIT was first described in the 1970s and used by several groups^1,2,3,4,5. It regained popularity after the breakthrough in human islet transplantation in the year 2000^6,7. However, most islet transplant studies used the kidney capsule as a preferred site for experimental islet transplantation due to its easy success. On the contrary, IIT is more technically challenging and less frequently used for islet transplantation studies^8,9. Unlike IIT, however, islets transplanted under the kidney capsule do not suffer from the immediate blood-mediated inflammatory reaction characterized by thrombosis, inflammation, and hepatic tissue ischemia, and thus have better function than islets transplanted into the liver. The kidney capsule model, therefore, may not fully mimic the stresses encountered by islets in human islet transplantation^10,11,12.

One of the major complications of IIT in mice is bleeding from the injection site after transplantation, which could cause 10-30% of mortality among different mouse strains¹². In this paper, two refined approaches have been developed to stop bleeding more rapidly and securely and to reduce mouse mortality after an IIT. Visual demonstration of these refined details will help researchers identify the key steps of this technically challenging procedure. In addition, the location of the islet grafts in the recipient’s liver was determined by histological examination of the Hematoxylin and Eosin (H&E) stained liver tissue (whole section) bearing transplanted islets.

Protocol

All procedures were conducted with the approval of the Institutional Animal Care and Use Committees at the Medical University of South Carolina and the Ralph H Johnson Medical Center in Charleston.

1. Diabetes induction using streptozotocin (STZ)

1. Recipient mice preparation:
   1. Weigh all mice individually.
   2. Check blood glucose levels from a tail vein blood sample using a glucometer.
2. STZ dose determination for three different scenarios:
   1. For mice with fatty liver disease inject one dose of STZ [40 mg/kg/day, intraperitoneal (i.p.) injection] for 5 consecutive days.
   2. For NOD-SCID mice inject 125 mg/kg of STZ, single injection, i.p. after overnight fasting.
   3. For C57BL/6 mice inject 225 mg/kg of STZ, single injection, i.p.
3. Calculations for STZ (13.5 mg/mL):
   NOTE: This calculation is for five C57BL/6 mice with body weights of 30 g:
   1. Total body weights: 5 mice x 30 g/mouse = 150g
   2. STZ needed: 150 g x 225 mg/1000g STZ = 33.75 mg
4. STZ preparation:
   1. Weigh the STZ following the pre-calculated dose.
   2. Transfer the weighed STZ powder into a 10 mL beaker on ice.
   3. Add 3 mL of sodium citrate solution to the beaker to dissolve the STZ.
   4. Mix well, filter sterilize through a 0.22-μm pore, and use the STZ solution within 10 min of preparation.
5. STZ injection:
   1. Load the desired amount of STZ solution (enough for one mouse) into 1 mL syringe.
   2. Perform intraperitoneal injection at the lower right quadrant of mouse abdomen.
   3. Observe mice for 5 min after injection and check for any signs of discomfort during this period of time before putting them back into the cages.
   4. Monitor blood glucose level from a tail vein blood sample using a glucometer daily after the STZ injection.
      NOTE: In this experiment, mice are considered diabetic when non-fasting blood glucose is > 350 mg/dL for two consecutive days.

2. Islet preparation

NOTE: Human islets were cultured in CMRL-1066 media supplemented with 10% fetal bovine serum (FBS), and 1% penicillin/streptomycin (P/S) at a density of 10,000 islet equivalent number (IEQ) per 100 mm cell culture dish⁹. Mouse islets were cultured in DMEM with 10% FBS and 1% P/S with the same density¹³. Male NOD-SCID and C57BL/7 mice between 6-10 weeks of ages were obtained from commercial sources.

1. Detach cultured islets from cell culture dish by gentle scratching.
2. Hand-pick desired numbers of islets (e.g., 300-350 islets) using a 1cc syringe and put them into sterile 1.5 mL microcentrifuge tubes on ice.
3. Spin the tube for 10 seconds using the microcentrifuge.
4. Remove the supernatant, leaving some liquid to avoid losing the pellet.
5. Resuspend the pellet in 200 μL of HBSS with 0.5% bovine serum albumin (BSA).
6. Aspirate the resuspended islets into a 0.5 mL insulin syringe.
7. Place the syringe in the upright position. Let the islets sink down for 1 min.
8. Push the syringe to remove all the bubbles, leaving about 100-150 μL of liquid containing islets.
9. Place syringe head down and gently tap the side of the syringe to let the islets equally distribute throughout the liquid. Islets are now ready for injection.

3. Islet transplantation

1. Induce and maintain the mouse under general anesthesia with 2% isoflurane. Check for the lack of pedal reflexes to ensure proper anesthetization of the animal.
2. Shave and remove the fur in the abdomen area of the mouse.
3. Administer a single pre-operative dose of Buprenorphine (0.1 mg/kg i.p.).
4. Disinfect the surgical area with three alternating wipes of 2% iodine and 75% alcohol.
5. Perform a laparotomy with micro scissors to generate a 1-1.5 cm incision.
6. Open the peritoneal cavity with a retractor. Follow with either method A or method B as detailed below.

4. Method A: (stop bleeding with gel foam,  Figure 1A)^14,15,16

1. Mouse preparation
   1. Place a sterile gauze around the incision.
   2. Gently pull out the intestine using a forceps and keep it on the gauze.
   3. Identify the portal vein by its location and expose it well.
   4. Cover the intestine with a warm saline-wet gauze during the entire surgery.
2. Insert the islet preloaded insulin syringe needle through the portal vein near the duodenum (Figure 1B). To do so, hold the needle with the hole (bevel) facing down and position the opening surface's angle parallel to the portal vein wall before penetrating through the wall.
   1. Pull the plunger to draw some blood (20-50 μL) into the syringe to mix the islets first.
   2. Infuse the islets into the portal vein slowly while repeatedly pulling and pushing the plunge.
   3. Place a piece of gel foam (about 0.5 cm x 0.5 cm in size) to cover the injection site.
   4. Press the gel foam down with a cotton tip while pulling out the needle from the portal vein.
   5. Continue pressing on the gel for about 2 min to confirm there is no active bleeding.
   6. Rollover the cotton tip over and away from the gel foam to make sure the gel foam covers the portal vein well.

5. Method B: (stop bleeding with fat pad,  Figure 1C)¹⁷

1. Expose the portal vein thoroughly.
   1. Use two cotton tips to hold the exposed portal vein from both the left and the right sides.
   2. Identify the fat tissue pad between the duodenum and the portal vein.
   3. Penetrate through the fat pad before inserting the needle into the portal vein (Figure 1D).
   4. Infuse the islets, following the similar procedure described above in part 4.2.1 and 4.2.2 of Method A.
   5. Pull out the needle while pressing down on the fat with a cotton tip.
   6. Continue pressing on the fat pad for 1 min after removing the needle.
2. After confirming that there is no bleeding from the portal vein, gently return the intestine to the peritoneal cavity in its original position.
3. Leave 0.5 mL of warm saline (36-37 °C) in the abdominal cavity before closure.
   NOTE: Warm saline facilitates post-surgery intestine movement and recovery and prevents intestine necrosis.
4. Close the muscle layer with an 5-0 suture.
5. Close the skin layer with an 4-0 suture.
6. Place the mouse in a clean cage on a heating pad until fully recovered from anesthesia.
7. Continue to provide an analgesic (e.g., buprenorphine 0.1 mg/kg i.p.) every 12 h and supplemental heat for 48 h post-surgery.
   ​NOTE: The islet transplantation procedure takes approximately 15-20 min to complete.

6. H&E staining and photograph of whole liver section

1. Liver perfusion
   1. Put the mouse under anesthesia as described above in part 3.1.
   2. Carefully expose the portal vein and cut the inferior vena cava.
   3. Manually perfuse the liver using 20 mL of 10% paraformaldehyde via the portal vein for about 5 minutes, using a 20 mL syringe with 25G needle¹⁸.
      NOTE: Liver perfusion can remove blood from liver tissue and improve liver fixation without disturbing the islet grafts.
   4. Dissect the perfused whole liver from other organs.
   5. Fix the perfused liver tissue in 10% paraformaldehyde for 24 h.
   6. Embed the tissue in paraffin.
   7. Cut tissue sections of 5 µm thickness each and put them on a glass slide for staining.
   8. Perform H&E, insulin, fibrin, and polymorphonuclear neutrophil (PMN) staining using standard methods^15,16.
   9. Scan whole liver section under a microscope.

Results

We performed syngeneic and xenogeneic islet transplantations via the portal vein. Islet graft function was observed in a dose-dependent manner in both islet transplantation models. In the syngeneic islet transplantation model using C57BL/6 mice, transplantation of 250 islets led to transitory normoglycemia before mice returned to hyperglycemia. Mice receiving 500 islets reached and maintained normoglycemia beyond 30 days after transplantation (Figure 2A). Mice in both groups showed increased...

Discussion

In this study, two improved procedures that can prevent bleeding and may reduce mouse mortality during mouse IIT have been demonstrated. This study enables researchers to visualize the islet transplantation model that is unique in studying the instant blood mediated inflammatory response after transplantation. The IIT model is a distinctive model for studying islet cell survival and hepatic ischemic injuries in response to islet transplantation¹⁹. Here, we refined the procedure based on previous s...

Materials

Name	Company	Catalog Number	Comments
10% Neutral buffered formalin v/v	Fisher Scientific	23426796
1 mL Syringe with needle	AHS	AH01T
20 mL Syringe	BD	301031
25G x 5/8" hypodermic needles	BD	305122
Alcohol prep pads, sterile	Fisher Scientific	22-363-750
Animal Anesthesia system	VetEquip, Inc.	901806
Buprenorphine hydrochloride, injection	Par Sterile Products, LLC	NDC 42023-179-05
Centrifuge tubes, 15 mL	Fisher Scientific	0553859A
CMRL-1066	Corning	15110CV
DMEM	Corning	10013CV
Ethanol, absolute (200 proof), molecular biology grade	Fisher Scientific	BP2818500
Extra fine Micro Dissecting scissors 4” straight sharp	Roboz Surgical Instrument Co.	RS-5882
Fetal bovine serum (FBS)	Corning	35011CV
FreeStyle  Glucose meter	Abbott	Lite
FreeStyle Blood Glucose test strips	Abbott	Lite
Gelfoam (absorbable gelatin sponge, USP)	Pharmacia & Upjohn Company	34201
Graefe forceps 4” extra delicate tip	Roboz Surgical Instrument Co.	RS-5136
Heated pad	Amazon	B07HMKMBKM
Hegar-Baumgartner Needle Holder 5.25”	Roboz Surgical Instrument Co.	RS-7850
Insulin syringe with 27-gauge needle	BD	879588
Iodine prep pads	Fisher Scientific	19-027048
Isoflurane	Piramal Critical Care	NDC 66794-017-25
Penicillin/streptomycin (P/S)	HyClone	SV30010
Polypropylene Suture 4-0	Med-Vet International	MV-8683
Polypropylene Suture 5-0	Med-Vet International	MV-8661
Sodium chloride, 0.9% intravenous solution	VWR	2B1322Q
Streptozocin (STZ)	Sigma	S0130
Surgical drape, sterile	Med-Vet International	DR1826
Tissue Cassette	Fisher Scientific	22-272416