In Vitro and In Vivo Approaches to Determine Intestinal Epithelial Cell Permeability

The overall goal of this experiment is to study the diseases caused by epithelial barrier dysfunction by measuring the intestinal epithelial cell permeability in vitro and in vivo. This method can help answer key questions in the intestinal barrier function field, such as those related to the inflammatory bowel disease. The main advantage of this technique is that intestinal epithelial cell permeability can be assessed both in vitro and in vivo.

The implications of this technique extends towards therapy or diagnosis of gastrointestinal diseases, because intestinal epithelial barrier dysfunction contributes to those diseases. Though this method can provide insight into the intestinal barrier function, it can also be applied to other systems, such as drug toxicity studies and barrier integrity of other cell types. Visual demonstration of this method is critical, as some steps are difficult to learn.

An accurate operation of this method can now be understood by textual description. To begin, grow Caco-2bbe cells in a T75 flask with medium. Feed the flasks regularly, depending on the cell density.

When the cells are 80%confluent, remove the medium and use 1-2 milliliters of sterile PBS without calcium to rinse them. Add 1.5 milliliters of Trypsin-EDTA, and gently rock the flask. Then, place it in the 37 degree Celsius incubator for 20 minutes, without rocking.

While the cells are trypsinizing, place inserts containing porous polycarbonate membranes into 24 well plates. Then add 1 milliliter of culture medium into the basal chamber which is the lower space of the membrane. Add 5 milliliters of medium to the flask, and vigorously pipette the cells against the inside of the flask 5 to 10 times, to separate the culture into loose individual cells, or two to three cell clumps.

Plate 0.166 milliliters of the cells into the apical chamber which is the upper space of the membrane. Then incubate the plates at 37 degrees Celsius for up to three weeks. Feed the cells three times weekly, by using a pressure pump to carefully aspirate the medium from the basal compartment of each well.

Then gently drip one milliliter of medium into the apical chamber of each insert. For cytokine studies, one day before transepithelial electrical resistance or TER measurements, replace the basal medium with medium, containing 10 nanograms per milliliter of interferon gamma. On the day of the experiment, replace the medium with HBSS containing 2.5 or 7.5 nanograms per milliliter of TNF.

To correct the meter, insert the correction electrode into the input port and choose Ohm mode. Then use a screwdriver to adjust the R Adjustment screw until the meter displays a reading of 1, 000 ohms. Sterilize the electrodes in 70%ethanol for 15 to 30 minutes and allow them to air dry for 15 seconds.

Then rinse the electrodes in experimental cell culture medium. Next, turn on the power and choose Ohm mode. While keeping the electrodes vertical, carefully place the long ends of electrode bridges into the basal chamber, ensuring that they touch the dish.

Then place the short ends into the apical chamber, ensuring that they stay below the surface of the medium, but above the tissue culture inserts. Measure the resistance of the sample and blank inserts at 0, 1, 2, 3 and 4 hours after cytokine treatment. Then record the resistance.

To achieve consistency across different plate formats, calculate the product of the resistance and the effective membrane area as shown here. For 24 well inserts, the effective membrane area is 0.33 square centimeters. To induce colitis, add DSS to autoclaved water to a final concentration of 3.5%weight per volume.

Then use the DSS solution to replace the drinking water in the cages of 8-week-old male C57 black six mice for a total of seven days. Give regular drinking water without DSS to control mice. After day seven, switch the DSS water to regular drinking water.

Each day, weigh the mice and assess the clinical scores as defined according to the disease severity by four parameters:rectal prolapse, stool consistency, bleeding, and activity. Sum the scores from these parameters for a final clinical score. To analyze the histopathological condition of colon tissues seven days post DSS treatment, after euthanizing mice according to the text protocol, isolate the colon and cecum, and measure the length of the colon.

Cut 0.5 centimeter segments from the distal colon, and fix the tissue in a 15 milliliter falcon tube containing 10 milliliters of 10%formalin overnight. Wash the fixed tissues with graded ethanol and xylene. Then embed the tissues in paraffin and cut six millimeter sections for the hematoxylin and eosin staining.

To measure epithelial barrier permeability in DSS-induced mice, seven days after the start of DSS administration, fast the mice for three hours. Autoclave a gavage needle to ensure sterility, then use 150 microliters of 80 milligrams per milliliter four kilodalton FITC dextran in sterile water to gavage the mice, and keep the unused FITC dextran to measure the standard curve after serum collection. Next, weight the mice for the permeability calculation.

Four hours later, after anesthetizing the mice by IP injection, confirm proper anesthetization by the lack of reflexes and whisker movement. Then place the mice on a heat block for five minutes. Using a pair of scissors, clip a one centimeter piece of the tail and collect 100 microliters of blood from the tail into a serum collection tube.

Spin the collected blood at 10, 000 times g and room temperature for ten minutes. With water, dilute the serum one to four. Then, to make a standard curve, use water to prepare the dilutions of the unused FITC dextran.

Add 100 microliters per well of the serum and the standard curve samples into 96 well plates. Using a plate reader, read the fluorescence at an excitation of 485 and an emission of 528. Calculate the permeability values based on the standard curve, and multiply by four to correct for the dilution.

Divide the concentration of FITC dextran by the weight to normalize the values. This helps to normalize the difference in FITC dextran delivery if mice are sick, and have lost weight. The Caco-2bbe cells shown here were labeled with nuclei and F-actin stains to show the difference between undifferentiated and differentiated cells.

Stress fibers are clearly seen in undifferentiated cells. Differentiated cells have a smaller volume, larger nuclei, and fewer stress fibers than undifferentiated cells. TNF is central to intestinal barrier loss through MLCK-dependent tight junction regulation.

As seen in this figure, TNF significantly decreases the TER of the Caco-2bbe monolayer in a dose-dependent manner, suggesting that TNF increases epithelial permeability. Compared to their initial body weight, mice treated with DSS lose a significant amount of weight. The severity of colitis is scored by rectal prolapse, stool consistency, bleeding, and activity.

Cross-sections of colonic tissues stained with hematoxylin and eosin show colonic mucosal damage in DSS-treated mice. In addition, the length of colon crypt, as well as the colon itself, is decreased in DSS-treated mice. Finally, there is an approximately two-fold increase in the levels of FITC dextran staining in DSS-treated mice, compared to control mice.

While attempting this procedure, it’s important to remember to minimize error, by standardizing operational procedures and utilizing statistical methods. After its development, this technique paved the way to researchers in the field of intestinal barrier function to explore gastrointestinal diseases in both model organisms and cell lines. After watching this video, you should have a good understanding of how to determine intestinal epithelial permeability by measuring the TER, gavaging FITC dextran, and observing morphological and histochemical changes.