Pancreatic Islet Isolation and Purification from Lewis Rats Using Enzymatic Digestion and Density-Gradient Separation

Our research focuses on developing a standardized high throughput Lewis Rat eyelet isolation protocol to optimize yield, purity, viability, and functional longevity. Existing methods yield variable impure short-lived eyelets. This protocol standardizes digestion, purification and pooling to improve consistency and longevity.

To begin, place a euthanized rat under a microscope. Using skin scissors, make a V-shaped incision starting from the pubic synthesis toward each lateral rib cage to access the abdominal cavity. Using flat forceps, retract the small and large intestines to the left side of the animal to clear the abdominal cavity workspace.

Then use the flat forceps and curved hemostats to unfurl the upper duodenum. Ensure the common bile duct and hepatic portal vein are within the field of view. Now use the curved hemostats to retract the liver in the cephalon direction.

Clamp the common bile duct and hepatic vein one to two millimeters from the liver. Under the microscope view, with flat forceps and flat hemostats, gently mobilize the contents within the duodenum to clear the sphincter of audi. Then gently clamp the duodenum approximately five millimeters from each side of the sphincter of audi to isolate the surrounding region.

Ensure minimal compression to preserve tissue integrity. Carefully insert the needle of a syringe containing sterile WDM through the duodenal wall. Pass the needle through the sphincter body and advance it into the main pancreatic duct for perfusion.

While maintaining constant pressure, perfuse seven to 10 milliliters of WDM into the pancreas at a rate of about 500 microliters per second until the pancreas is fully distended. Now, remove both flat hemostats from the duodenum. Then gently dissect and free the pancreas from surrounding structures starting from the spleen, stomach, large intestine, duodenum, and mesentary.

Gently lift the perfused pancreas with flat forceps and use fine dissecting scissors to sever any remaining connective tissue attachments. Immediately transfer the excised pancreas into a Petri dish containing ice cold rinse solution and proceed to harvest the next pancreas while the excised pancreas is being cleaned. To clean the pancreas's use a pair of fine curved forceps and fine dissecting scissors to carefully remove lymph nodes, adipose tissue, mesentary, and large vasculature from the perfused pancreas if present.

Transfer the cleaned pancreas into a sterile 600 milliliter beaker containing 25 milliliters of cold WDM. Cover the beaker with sterile aluminum foil to maintain cleanliness during transfer from the surgical workspace to the cell culture workspace. Transfer the beaker into an orbital shaker water bath set at 130 revolutions per minute for 10 minutes.

At the five minute mark, manually swirl the beaker clockwise and then counterclockwise to aid tissue dissociation. At the 10 minute mark, rapidly transfer the beaker and submerge it in ice to slow enzyme activity. Then add an equal volume of quenching buffer to the solution in the beaker to stop the digestion.

Using a 50 milliliter syringe fitted with a 14 gauge spinal needle, gently aspirate and expel the tissue suspension repeatedly to dissociate large tissue fragments. Transfer the suspension into 200 milliliter conical tubes. After centrifuging the suspension, carefully decant the supernatant and resuspend the pellet in 100 milliliters of washing media.

Now filter the suspension through a sterile 500 micrometer sieve placed over a sterile 600 milliliter beaker to remove large undigested tissue. Transfer the filtrate into new 200 milliliter conical tubes before centrifuging again. For eyelet purification, use a pre wedded 10 milliliter serological pipette to resuspend each tissue pellet in 15 milliliters of gradient solution per two milliliters of pellet volume.

Mix gently until the suspension is uniform. Carefully dispense 15 milliliters of the cell suspension into the bottom of each 50 milliliter conical tube avoiding contact with the tube walls to ensure clean gradients. Then use a syringe pump to dispense 10 milliliters of each gradient at a rate of five milliliters per minute while positioning the 16 gauge needle tip against the inner wall of the tube to prevent mixing.

After centrifuging the tubes, identify the eyelet layer located at the interfaces between the gradient layers. Using a wedded past pipette, collect the eyelet layer, transfer the collected eyelets into a pre chilled 200 milliliter conical tube containing five milliliters of washing media. Bring up the volume to 200 milliliters with washing media and perform the following washing and centrifugation steps.

Then evenly plate the eyelets into non tissue culture treated 100 millimeter Petri dishes using one plate for every five pancreas'processed. Across the test isolation, yield was 3, 330 eyelet equivalents per pancreas with a median eyelet diameter in the 50 to 100 micrometer bin. Automated sizing yielded 98, 832 total objects, which strongly correlated with manual eyelid equivalents measured at day zero.

The endocrine purity was about 98.56%based on dithozone staining at day one. Stimulation indices were 9.37 at day one, 8.34 at day three and 6.7 at day seven with no significant differences between time points. This method enables the study of eyelet transplantation, beta cell function, diabetes reversal, eyelet encapsulation, and eyelet immune or microenvironment interactions.

This protocol can be applicable to preclinical transplantation, diabetes modeling, encapsulation studies, and invitro functional assays requiring high purity Lewis Rat eyelets.