A Method for Murine Islet Isolation and Subcapsular Kidney Transplantation

Eyelet transplantation has been proposed as a potential treatment for type one diabetes. However, two major limitations prevent it from being a widespread clinical reality. First, a large number of eyelets is needed per recipient.

Since eyelets are obtained from organ donors, the number of potential recipients is severely limited. Second transplantation requires lifetime treatment with strong immunosuppressants, which puts the patient, especially the pediatric population at risk for malignancy, infection, and renal failure. Eyelet transplantation under the mouse kidney capsule is performed to investigate strategies for improving clinical transplantation outcomes.

Eyelets isolated from healthy donor mice assorted into groups and prepared for transplantation. The eyelets are then transplanted into the subcapsular space of the kidney. In diabetic recipient mice eyelet graft function is assessed by monitoring blood glucose levels, which reveal that approximately 250 to 350 eyelets will restore u glycemia in a 20 gram recipient mouse within 24 hours of transplantation.

Visual demonstration of this method is critical as to steps for cannulating. The common bile duct and transfer of the eyelets to the subcapsular space of the kidney are difficult to learn. These steps require a identification of tissue structures, steady hands, and the use of technique that minimizes tissue damage.

Place a euthanized mouse in the supine position on a paper towel at the dissecting scope and spray the abdomen with 70%ethanol. Use scissors to open the abdomen with a V incision from the pubic region to both front legs. The mouse is below the dissecting scope.

Position it with the head pointing towards the observer. Locate the duodenal entry of the common bile duct with a hemostat clamp, the duodenal opening of the common bile duct. The hemostat should be positioned such that when compressed, it runs exactly along the pancreatic intestine border.

Here two methods to cannulate the bile duct will be demonstrated. The free hand method and the forceps assist method to cannulate the bile duct. Using the free hand method, pull the hemostat clamped on the duodenum away from the head of the mouse toward the tail so that the common bile duct becomes taut.

There is a confluence in the bile duct close to the liver where bile draining from the gallbladder and enzymes from the liver come together Before entering the intestines, pull the bile duct tight to expose the inside of the V position. A bent 27 gauge needle attached to a five milliliter syringe so that the needle slides right through the V and directly cannulate the lower portion of the duct. Slide the needle several millimeters beyond the end of the bevel to prevent backflow into the liver and gallbladder.

Once the cannulation is successful, perfuse the pancreas by dispensing two milliliters of liase from the syringe. Watch for the expansion of the pancreas over the stomach as an indication of optimal cannulation. Alternatively, the bile duct can be cannulated.

Using the forceps cyst method. Pull the hemostat C clamped on the duodenum away from the head of the mouse towards the tail so that the common bile duct becomes tort. Then using a bent 27 gauge needle attached to a five milliliter syringe, puncture the fascia below the common bile duct close to the liver.

Use the needle to clear the fascia from the duct with the needle still in place. Set the hemostat down and pick up a pair of forceps. Use one arm of the open forceps to support the bile duct where the needle has cleared the fascia while pulling the duct and forceps forward.

Slide the needle into the duct using the forceps as a backstop. A successful cannulation is indicated by an even expansion of the pancreas region on top of the stomach. Once the pancreas has been perfused, remove the hemostat from the duodenum.

Then use the forceps to lift the duodenum and use a second pair of forceps to separate the pancreas from the intestines. Next, pull the pancreas free from the top of the stomach and the spleen. Finally, lift the pancreas out of the abdomen and cut it free from the remaining fascia connections to the intestine, stomach and spleen.

Place the pancreas in a 50 milliliter conical tube and place it on ice. Incubate the pancreas at 37 degrees Celsius according to the instructions in the accompanying written protocol. After incubation, add 20 milliliters of media to the tubes and dissociate the tissue by shaking the tubes vigorously 40 times in 10 seconds.

This step is critical for optimal recovery of violets. Spin cells are 800 rotations per minute. Resus spend in 15 milliliters of media and condense the samples.

Pour the resuspended slurry through a 0.419 millimeter wire mesh and funnel it into a fresh 50 milliliter conical tube. Rinse the first tube with an additional 10 milliliters of media containing 10%fetal bovine serum and pour it through the wire mesh pellet the cells by centrifugation reus, bend the pellet in five milliliters of room temperature his opaque 10 77. Then vortex gently for several seconds, add an additional five milliliters of his opaque around the inside of the tube to wash the eyelets off the wall of the tube slowly pipette to overlay 10 milliliters of serum free RPMI down the side of the tube at a rate of one milliliter per 10 seconds.

A sharp interface should be evident. Then spin the samples in a swinging bucket centrifuge at 20 degrees Celsius and 900 times gravity for 20 minutes with very slow acceleration and no braking following the centrifugation, the islets will be separated from the exocrine cells using a disposable 10 milliliter serological pipette. Transfer the eyelets from the histo Paque media interface to a fresh 50 milliliter conical tube.

Several tubes can be pooled at this point to remove the eyelets residual, his opaque fill tubes with serum containing media and pellet the islets by centrifuging the tubes for two minutes at 800 rotations per minute. Then reus suspend the islets in fresh media. Repeat this step at least two times.Reus.

Suspend the pallet in five milliliters of media. Place a 0.1 millimeter nylon cell strainer upside down on a 15 milliliter conical tube. Pipette the resuspended cell slurry through it to collect the eyelets.

Then rinse the 50 milliliter tube with an additional six milliliters of media and pass it through the strainer. Place a three milliliter drop of medium in a 10 centimeter Petri dish. Invert the cell strainer and dip it into the media drop to collect the eyelets pipette an additional five to seven milliliters of culture medium through the strainer into the Petri dish.

To collect any remaining eyelets from the strainer under a four times objective, pick eyelets individually with a P 200 pipette and transfer them to a 1.5 milliliter micro fuge tube. These images show eyelets isolated from 12 week old C 57 black six mice. The purity and quality of the eyelets can be estimated microscopically upon completion of the isolation procedure.

For example, the two images shown here show a representative field where debris and exocrine cell contamination are present. When picking eyelets for an experiment, avoid picking any contaminating exocrine cells or eyelets that appear unhealthy. Eyelets with necrotic centers are unhealthy as shown in this figure taken 24 hours after isolation, necrotic centers will not be visible immediately after isolation.

Eyelets that are sloughing cells are unavoidable and can be used in eyelet transplantation in this 24 hour time lapse video of eyelets. Immediately following isolation, necrotic centers develop within many of the eyelets. These necrotic centers appear to be expelled from the eyelet near the end of the recording while picking eyelets.

Group them so that each tube has an even distribution of eyelet quality and size. For example, each group of 15 eyelets might contain four large-sized eyelets, six medium eyelets, and five small eyelets.Place. The tube on ice eyelets can be used immediately for transplantation or manipulated.

Before use. Place a sterile gel loading tip into the opening of a 1.5 milliliter micro fused tube so that there is a gentle bend, which makes a U-shape in the narrow end of the tip. Both openings of the tip should be facing directly up outta the micro refuse tube.

Gently remove a tube of eyelets from ice. Ensure that all of the eyelets have settled at the bottom of the tube. Using a P 200 pipette gently collect the eyelets in a minimal volume from the bottom of the tube and transfer them to the prepared curved tip.

After the eyelets have sedimented, remove as much of the media from the tip as possible by aspirating the media through the large opening of the eyelet bearing gel loading tip With a fresh gel loading tip attached to a P 200 pipette. Attach the non beveled end of tubing to the narrow opening of the islet bearing gel loading tip. Then attach the tip to a P 200 pipette and gently push the islets into the tubing.

Next, connect the non beveled edge of the PE 50 tubing to the needle of a 25 microliter Hamilton syringe. With the plunger withdrawn to press the plunger of the syringe until the eyelets are approximately 0.5 centimeters away from the beveled opening. Set the syringe aside so that the eyelets may settle into a single mass near the beveled opening of the tube while the kidney is being prepared to receive the eyelets.

Prepare the surgical site by clipping hair and scrubbing skin with Betadine alternating with 70%isopropyl alcohol three times. Begin the eyelet transplantation procedure by performing a toe pinch to confirm anesthesia in the recipient mouse. Place the shaft of a glass rod directly underneath the mouse and below the kidney perpendicular to the spinal cord.

Then locate the kidney through the skin and place a sterile surgical drape. Once the kidney is located, use McFerson banis scissors to make a two centimeter incision through the dermis directly above it perpendicular to the spinal cord. This incision will expose the peritoneal wall with the scissors.

Make a one centimeter incision through the peritoneal wall in the same direction as the cut through the dermal layer using the shaft of the glass rod as a backstop, force the kidney through the peritoneal opening by pressing down on the adjacent surface. Wet the surface of the exposed kidney with ice cold, sterile PBS using a soaked cotton tipped applicator. Repeat the step every few minutes as needed to prevent the kidney capsule from drying out.

Using a 27 gauge needle, make a 0.2 centimeter incision through the kidney capsule across the anterior surface of the kidney. Moving from the left lateral side towards the right lateral side, insert a flame pulled glass capillary tube. Probe through the incision made earlier and move it in a posterior direction along the dorsal lateral surface of the kidney.

To create a pouch between the kidney capsule and kidney parenchyma, move the eyelets in the prepared 25 milliliter Hamilton syringe to the very edge of the beveled opening of the flexible tube. Then insert the beveled edge facing up into the subcapsular pouch. The long edge is in contact with the kidney parenchyma.

Move the tube to the most posterior end of the capsule. Then slowly transfer the eyelets from the PE 50 tube into the subcapsular pouch by depressing the plunger of the syringe. As the eyelets fill the pouch slowly back the flexible tube out to make more room.

After removing the tube from the pouch, slide the glass probe along the outer surface of the kidney anterior to posterior to gently pack the eyelets into the proximal end of the pouch. Using bent arm forceps, lift the peritoneal lining adjacent to the opening made for the kidney. Then use a PBS soaked cotton tipped applicator to gently push the kidney back into the peritoneal cavity.

Close the mouse by applying sutures to the peritoneal wall and wound clips to the dermal layer. Return the mouse to its cage and monitor. For recovery, aceto meine water should be provided for the mouse bowel.

C mouse eyelets isolated. Using the methods described in this video and a curative dose was transplanted into allogeneic C 57 black six mice. Non-fasting blood glucose levels were monitored beginning on postoperative day one.

Rejection is to be seen around day 17 in 20 gram recipient mice that received 300 eyelets from an allogeneic donor Here. Recovery was monitored in mice that received either 150 eyelets or 300 eyelets from syngeneic donors. No rejection is seen in this transplants as the eyelets are from syngeneic donors.

The eyelet dose administered measures the primary function of the eyelets immediately following transplantation. While marginal recovery was seen with 150 eyelets, full recovery was seen when 300 eyelets were transplanted. After watching this video, you should have a good understanding of how to perform the critical steps required to isolate mouse eyelets and transplant those eyelets under the subcapsular space of the kidney.

With practice, you'll develop the ability to quickly cannulate the common bile duct, which is the most difficult step of this protocol.