Mixed Primary Cultures of Murine Small Intestine Intended for the Study of Gut Hormone Secretion and Live Cell Imaging of Enteroendocrine Cells

The overall goal of this method is to isolate and culture mixed murine intestinal cells to enable the study of gut peptide secretion and live cell imaging of enteroendocrine cells. We originally developed this method to enable the in vitro characterization of enteroendocrine cells. Enteroendocrine cells monitor intestinal content and regulate body homeostasis.

But they have been difficult to study as they are scattered throughout the intestinal epithelium. The calcis described here are particularly useful when combined with genetic fluorescent tagging of enteroendocrine cells, as this enables the use of single-cell electrophysiological and live cell imaging techniques. Demonstrating the procedure will be Cheryl Brighton, a post-op in our laboratory.

To begin, transfer an isolated mouse duodenum to a 10 centimeter petri dish filled with PBS. Using forceps, carefully seize one end of the intestinal fragment, and introduce the tip of a pasteur pipette into it's lumen. Then rinse out the intestinal contents with chilled PBS.

Repeat for both intestinal ends until the majority of the contents are removed. Transfer the rinsed intestine to a clean petri dish filled with fresh chilled PBS. Under a dissecting microscope, remove any adiposed tissues and mesentary from the intestine fragment, while preventing the muscle layer from pulling off.

Next, identify a flap of the muscle layer at the proximal end of the tissue. Then, using two sets of fine foreceps, gently pull away a small amount of the layer all the way around the intestine. Grasp the intestine and as much of the muscle flap as possible, then gently pull the layers apart and start peeling off the muscle layer from around the intestine.

Keep the forceps close together, to prevent the muscle layer and epithelium from tearing. Remove the muscle layer from the entire length of the intestinal fragment. Afterward, open the intestine by cutting it longitudinally, and transfer it to a clean petri dish containing fresh chilled PBS.

Agitate the tissue to wash it of any remaining chime or mucus. Once it is washed, cut the tissue with a surgical scalpel blade to obtain squares of approximately one to two square millimeters. Then using a pre-wetted pasteur pipette with a pre-cut tip, transfer the obtained fragments to a 50 milliliter centrifuge tube filled with approximately 20 milliliters of chilled PBS.

Gently shake the tube to additionally wash the tissue fragments, and let the tissue sediment. Once the tissue is settled, aspirate most of the PBS and wash the sample with fresh PBS one more time. To digest the intestine, use a 10 milliliter serological pipette to transfer the tissue fragments to a sterile 50 milliliter centrifuge tube, containing chilled sterile DMEM, swirl the suspension, allow the tissue to settle, and then remove the medium.

Add 7 milliliters of the digestion medium to the tissue fragments and swirl the suspension. Incubate the suspension in a water bath at 37 degrees Celsius for five minutes. After incubation gently shake the tube for approximately three seconds.

Sediment the undigested tissue, and transfer the digestion medium to a 15 milliliter centrifuge tube. Allow any accidentally collected tissue to settle, and using a 10 milliliter serological pipette, transfer it back to the 15 milliliter centrifuge tube, containing the undigested tissue. Preserve a small volume of the digestion medium for the microscopic observation and discard the remains.

Monitor the contents of the digestion medium under the light microscope to ensure the suspension does not contain many crypts. Next, add 7 milliliters of fresh digestion medium, and perform the digestion procedure one more time. To prepare crypt fragment suspension, add 7 milliliters of fresh digestion medium to the intestine fragments.

Incubate the tissue in a water back at 37 degrees Celsius for ten minutes. Shaking the sample for longer and more vigorously this time, every five minutes. Sediment the undigested tissue, and transfer the digestion medium to a 15 milliliter centrifuge tube.

Allow any accidentally collected tissue to settle and using a 10 milliliter serological pipette, transfer it back to the 50 milliliter centrifuge tube containing the undigested tissue. Add 7 milliliters of fresh digestion medium to the intestine fragments, and start the next digest. Then, centrifuge the medium collected from the previous digest at 100 x g at room temperature for 3 minutes.

After discarding the supernatent, re-suspend the cell pellet in 5 milliliters of the pre-warmed culture medium, by gentle pipetting. Then, set the sample aside and collect a small aliquot of the cell suspension. Under the light microscope, confirm that the sample contains isolated crypt fragments.

Repeat digestion of the tissue an additional 2-3 times or until the majority of the tissue has been digested. Once the digest supernatants have been collected, combine all of them, and centrifuge the sample at 100x g at toom temperature for three minutes. Discard the supernatant, and re-suspend the pellet by gently pipetting, until no clumps are visible and 5 milliliters of pre-warmed culture medium supplemented with 10 micromolar of Y-27632, that prevents anoikis.

Finally, filter the cell suspension through a 100 micron filter to remove any undigested tissue. Wash the filter with an additional 2 milliliters of the pre-warmed culture medium supplemented with Y-27632. Prior to seeding the cells, remove the plate coated with basement membrane matrix from the incubator.

Remove the excess basement membrane matrix solution from the plate, and add 250 microliters of the pre-warmed culture medium supplemented with 10 micromolar Y-27632 to each well. Seed 250 microliters of the cell suspension per well of the 24 well plate, by dropwise pipetting carried out in a slow zigzag motion. Incubate the plate overnight at 37 degrees Celsius, and 5%carbon dioxide.

Presented here are the microscope images of the monolayers of the primary small intestinal cells taken after 18 to 24 hours of culture. Primary small intestinal cells that were cultured in vitro, responded to stimuli that elevate intracellular calcium, or cyclic AMP resulting in a two-fold increase in glucagon-like peptide-1 release, when the cells were treated with bombesin and eleven-fold secretion enhancement when stimulated simultaneously with forskolin and IBMX. Finally, using cultures generated from transgenic mice expressing GCAMP 3 under the control of the pro-glucagon promoter, it was shown that primary small intestinal L cells respond to bombesin and potassium chloride stimulation with an increase in cytosolic calcium, as demonstrated by the enhanced fluorescence.

The demonstrated procedure is focused on the cultural small intestinal cells, which many laps have found challenging. We also use a similar practical for the culture of other intestinal segments, of both murine and human origin. Once mastered this technique can be done in four hours, if performed properly.

No two preparations are the same, so it's important to remember to inspect aliquots of each digest under the microscope. This allows you to assess progress and tailor the intensity of the shaking and the number of digests accordingly.