益生菌酵母的转型及其胃肠道粘膜免疫组织恢复继小鼠口服灌胃

The overall goal of these procedures is to develop a strain of probiotic yeast that can be easily transformed to express and deliver heterologous protein to the murine gastrointestinal tract. The main advantage of these techniques is that they allow for rapid generation of oxytrophic strains of probiotic yeast. These mutant yeasts can then be genetically manipulated without relying on antibiotic selection.

Oral gavage of mice and harvest of Peyer's patches also allows for verification of mutant yeast adherence to the gastrointestinal immune tissues. These methods will help answer key questions in the field of oral vaccination and facilitate experiments testing probiotics for the delivery of gastrointestinal therapeutics. After preparing YPD medium and growing yeast overnight to saturation, dilute cells one to ten in water in a plastic cuvette.

Then, lank a spectrophotometer and determine the optical density of the overnight culture. With 20 milliliters of sterile distilled water, dilute the cells to a concentration of 10 to the seventh cells per milliliter. Then, pour the diluted cells into a sterile plastic petri dish, and with the lid removed, place the plate into a UV crosslinker.

Next, expose the cell solution to cumulative incremental doses of UV irradiation, extracting 500 microliter aliquots following each increment to establish a killing curve, and identify the optimal percent survival for screening. Use sterile water to serially dilute the cells at one to ten increments. Then, pellet the cells from each dilution by centrifugation at maximum speed for one minute.

Aspirate the supernatant, and resuspend the cells in a 100 microliter volume of sterile water appropriate for plating yeast cells. Then, pipette the full volume of resuspended cells onto plates containing YPD solid medium, and use a sterile spreader to evenly distribute cells across each plate. Plot a survival curve, and determine the dose of UV irradiation to be used for screening according to the text protocol.

After exposing yeast to the dose of UV irradiation selected for screening, pellet the full volume of irradiated cells, and use 100 microliters of sterile water to resuspend the pellet. Plate onto minimal medium containing 5-FOA according to the text protocol, in order to select your three minus oxytrophic newtons. Confirm the URA3 minus phenotype by using the tip of a sterile toothpick to collect part of a single colony on the 5-FOA plate, and restreak it onto YPD, 5-FOA, and uracil minus plates.

Wrap the plates, and incubate upside down at 30 degrees Celsius for two to four days. Confirm oxytrophic status by verifying growth of colonies on YPD and 5-FOA, but not Uracil minus plates. After confirming the URA-3 minus status of irradiated yeast, set up an overnight filter for lithium acetate transformation, by inoculating 10 milliliters of YPD with a single mutant colony, and incubate in a warm room at 30 degrees Celsius.

The following day, use 50 milliliters of fresh, warm YPD to dilute the overnight cultures to an OD600 of 0.16 to 0.2, and incubate the cells on an orbital platform shaker set to 200 RPM, until the culture reaches approximately one times 10 to the seventh cells per mill, usually around four hours. After washing the cells with water and lithium acetate according to the protocol, add 50 microliters of resuspended cells to a transformation mixture of plasmid and carrier DNA. Then, add 300 microliters of PEG, TE, and lithium acetate and vortex.

Incubate the preparations at 30 degrees Celsius on an orbital platform shaker at 200 RPM for 30 minutes. Continue with the transformation as described in the text protocol. After preparing yeast cultures according to the text protocol, use a hemocytometer to determine the cell concentration, and adjust to 10 to the ninth cells per milliliter.

Pellet the cells by centrifugation at 2500 times G for three minutes. Then, pour off the supernatant and resuspend the cells by adding the appropriate volume of sterile water, and gently pipetting up and down. Fix an appropriate gauge gavage needle onto a one milliliter sterile syringe, and load the yeast sample, eliminating any air bubbles.

Load an additional syringe with sterile water to gavage control mice. Using the non-dominant hand, pick up the mouse to be gavaged and, with the index finger and thumb, tightly grasp the skin around the neck. Tuck the tail under the small finger.

Be sure the grip is secure and prohibits the mouse from moving its head. Next, gently insert the gavage needle into the mouse's esophagus by angling the needle along the roof of the mouth and back of the throat, keeping slightly to the left of center. Wait for the mouse to swallow the bulb of the needle.

If resistance is felt or the mouse gasps, gently remove the needle, and try again to find the esophagus. After the mouse has swallowed the bulb of the gavage needle, gently depress the syringe plunger to administer the yeast directly into the mouse stomach. Gently remove the gavage needle from the mouse stomach and esophagus, and return the mouse to the cage.

Check that the mouse is breathing and moving normally to ensure the gavage was performed properly. After sacrificing the mouse according to the text protocol, position it with the abdomen fully exposed and use 70%ethanol to spray the abdominal area. With scissors, make a transverse incision through the fur and skin.

Then manually pry the incision open to further expose the peritoneum, the thin serosal lining that covers the abdominal organs. Gently lift the peritoneum, and make a transverse incision to expose the intestines. Then, use blunt forceps to carefully tease the small intestine away from the mesenteric arteries, fat, and other tissues.

Expose the small intestine from the stomach in the upper left quadrant of the mouse abdomen to the cecum, the large pocket of intestinal tissue at the start of the large intestine. Next, isolate Peyer's patches by looking for one to two millimeter roughly circular patches of opaque tissue along the small intestine. Then, using curved dissection scissors, cut away the dome of the Peyer's patch, leaving margins to ensure that none of the surrounding tissue is collected.

Transfer the dissected Peyer's patches into complete IMDM. Then, pour the solution with suspended Peyer's patches onto a 40 micrometer cell strainer. After washing the Peyer's patches with complete IMDM, use a plunger from a one milliliter syringe to gently break up the tissue, and collect the cells in a 50 milliliter tube.

Pellet the strained cells at 1800 RPM for seven minutes. Then, plate and analyze the cells according to the text protocol. This figure shows a plating from a serial dilution of irradiated S Boulardii cells.

The serial dilution ensures that the CF use could be enumerated at each UV dose. From these images of yeast plates, the consistent growth of mutant colonies on YPD and 5-FOA, but not Uracil minus plates, indicates a Uracil minus oxytrophic phenotype. This graph illustrates the transformation efficiency for wild type S Boulardii, compared to S Cerevisiae, using both lithium acetate and electroporation techniques.

Although lithium acetate transformation is very efficient for S Cerevisiae, transformation efficiency for S Boulardii is greatly improved using electroporation. In these Brightfield and Fluorescence images, S Cerevisiae transformed with a URA3 plasmid encoating GFE demonstrates a functional expression of heterologous protein. In this figure, an example of viable CF use detected from Peyer's patches after animal dissection is shown.

A typical yield of CF use recovered per mouse is less than 10. You should now have a good understanding of how to generate oxytrophic mutant strains of probiotic yeast, using UV mutogenesis. We have also described how to test the ability of these yeasts to express heterologous protein and adhere to Peyer's patches of the mouse small intestine.

Following these procedures, other methods, such as ELISA and fluorescence microscopy can be used to evaluate proper folding and function of heterologous proteins expressed.