Characterization of Inflammatory Responses During Intranasal Colonization with Streptococcus pneumoniae

This procedure involves colonization of the murn nasopharynx with streptococcus pneumonia and the subsequent extraction of colonizing bacteria as well as adherent or recruited cells. This is accomplished by first preparing a bacterial inoculum of the S pneumonia. Strain of interest.

The live bacteria is then administered intranasally into anesthetized mice while they are in a restrainer apparatus After the desired length of colonization, mice are then euthanized and nasal lavages are performed by cannulating their exposed tracheas and flushing out the contents through the nas. Bacterial numbers present in nasal lavages can be quantified using microbiological techniques. Alternatively, immune responses can be assay using either flow cytometry techniques to phenotype recruited cells or quantitative PCR techniques.

To measure levels of expressed mRNA of interest prior to beginning, you’ll need to prepare mouse restrainers for the intranasal colonization portion of the procedure and cannulated needles for the nasal lavages. A mouse restrainer can be prepared simply by cutting off the tapered tip of a 50 milliliter falcon tube. Once you have created the aperture, make sure to use scissors to polish down any sharp ends.

For the preparation of cannulated needles, you will need the following PE 20 polyethylene tubing with an inner diameter of 0.38 millimeters. One mil syringe is capped with 26 and three eight gauge beveled needles, a pair of sharp, fine iris, scissors, and a pair of forceps. Cut the PE 20 tubing with the fine scissors into pieces roughly 2.5 centimeters or one inch in length, ensuring that each end has a beveled tip.

Using the forceps carefully slide one of these pieces onto the tip of your needle. Avoiding puncturing the tubing side, cannulated needles can be kept in 70%ethanol until needed and flushed with PBS directly before use. For intranasal colonization of mice with S pmo, we recommend using a concentration of 10 to the nine colony forming units per mil delivered in 10 microliters.

For a final dose of 10 to the seven CFU, keep the inoculum on ice until just prior to use and make sure to agitate in between each colonization. In addition to the inoculum, you will also need a P 10 or P 20 pipette sterilized pipette tips and your mouse restrainer. Isolate an individual mouse by its tail and restrain it by placing it into your previously prepared mouse restrainer apparatus.

If you’re having trouble directing the mouse into the restrainer, it may help to position it in front and above the mouse, allowing the animal to crawl upwards. Once the mouse is inside the restrainer, secure it by the base of its body gently pushing it upwards so that its nose emerges out of the tapered end of the restrainer. Using a P 10 or P 20 pipette inoculate each mouse by depositing 10 microliters of the prepared culture, distributing it evenly between both.Nares.

Allow the inoculum to drip into the nose by pulsing the inoculation gradually taking the time necessary for the mouse to inhale the inoculum. The mouse may sneeze some of the inoculum out of its nose, which is expected. Note that mice should be anesthetized for this procedure to ensure the bacteria remains localized to the nasopharynx and does not spread to the lower respiratory tract.

For the purposes of this demonstration, we are inoculating with PBS, but whenever working with live bacteria, make sure to conduct the procedure according to institute approved guidelines. For example, most institutions will require colonization to be conducted in a biosafety level two hood. If utilizing weight indicators as part of your endpoint monitoring way, MA immediately after colonization, monitor mice every 12 to 24 hours for clinical symptoms including lethargy, ruffled fur, and weight loss.

Once mice are colonized, keep them for the desired length of time. At the end of this period, mice will be euthanized and nasal lavage is performed as cervical dislocation can potentially damage the trachea. This method of euthanasia must be avoided.

Prior to performing nasal lavages, ensure that you have the following on hand. Previously prepared cannulated needles, 70%aqueous ethanol, a pair of forceps, a pair of sharp fine iris scissors, phosphate, buffered saline, RNA lysis buffer, and one mil einor tubes. Once you’re ready to begin, lay out the animal flat on its back, and using 70%aqueous ethanol sterilize the front of its fur, particularly the neck area.

Taking care to prevent ethanol from accessing the nares. Make a single longitudinal cut along the midline of the neck of the animal, allowing yourself to create an opening to envision the trachea. Carefully peel back skin to either side revealing the neck tissue beneath.

At this point, the trachea should be visible surrounded by longitudinal muscles on either side. Carefully snip these to provide a clear view of the trachea itself, taking care not to sever the surrounding vasculature. In the event you do accidentally cut the vasculature and blood is present in the region prior to proceeding.

Allow the bleeding to stop and then cleanse the area several times by dispensing sterile PBS and using sterile gauze to gently soak up excess moisture. Once the trachea is properly exposed, make a transverse semilunar cut in the trachea about halfway up. Drop sterile PBS into a cannulated syringe.

Insert the cannula into the trachea towards the nose, keeping the beveled edge pointing downwards for ease of insertion. Once the cannula is in place, rotate at 180 degrees and gently probe upwards until you feel light resistance. Place the EEND DPH tube designated for sample collection just beneath the nose of the mouse.

Test correct placement of the cannula by dispensing a minimal amount of PBS lavage fluid. Following this, a drop of fluid should form around the NAS of the mouse if the cannula is placed correctly. If the test PBS emerges directly out of the mouth of the animal, pull the cannula back and reposition it again by gently probing forward until very slight resistance is felt.

Take care not to push the cannula too far past this resistance as you’ll move it past the nasal palate and through to the oral cavity. Once you confirm that the cannula is correctly positioned, proceed to dispense 200 microliters of PBS rapidly to help displace and collect the maximum amount of cells. Content should flow out through the nares of the mouse and into the collection tube.

Place the sample immediately on ice. These samples can be subsequently assayed for bacterial load as well as cellular recruitment. To collect samples for RNA analysis, use a separate cannulated needle containing 500 microliters of RNA lysis buffer to perform a secondary nasal lavage.

Note that the RNA lysis buffer will denude the epithelium and destroy surrounding tissue. So care must be taken to avoid contact with organs such as the lungs. If retentions of these tissues is desired to quantitate nasopharyngeal bacterial load, you’ll need the following.

Epicor tubes, microbiological plates containing triptych soy egg are supplemented with 5%sheep’s blood. A vortex mixer, PBS sterile pipette tips and P 20 and P 200 pipettes. First working in a BSL two hood, prepare a serial dilution series for each mirin nasal lavage sample.

In general, concentrations of S pneumoniae in the nasopharynx can be expected to be between zero and 10 to the four CFU. Therefore, conduct three tenfold serial dilution, add 10 microliters of the neat nasal lavage sample to the first tube to a concentration of 10 to the minus one CFU per mil, and continue downwards until you have diluted three samples excessively vortex thoroughly in between each dilution. Please note that this video is just for demonstrative purposes.

Proper lab technique, including the changing of pipette tips between dilu should always be adhered to Using a sharpie, divide it back to your logical plate into quadrants and label the quadrants with the corresponding dilution to be plated on it. Plate out three drops of 10 microliter samples of each of your dilution on the agar plate. Allow these to dry for 15 to 30 minutes uncovered then cover plates and place upside down in a bacterial incubator with optimal conditions for pneumococcal growth, typically 37 degrees Celsius and 5%CO2.

Incubate these plates for 24 to 48 hours. After this period of time, remove the plates and determine the number of colonizing bacteria by averaging the colonies formed on the plate. For each dilution, animo colonies can be identified by their distinctive beige color, their zones of alpha hemolysis, which are small halos around each colony, created by bacterial digestion of the blood supplement in the agar.

And finally, by small depressions in the center of each colony that give it the appearance of an erythrocyte. After watching this video, you should have a solid understanding of how to establish a nasopharyngeal es pmo, colonization in a mouse, how to isolate bacteria and cells from the nasopharynx of a mouse after colonization, using a nasal lavage and how to quantify the corresponding bacterial load using microbiological techniques. We hope the methods described in this video will encourage you to use an intranasal model of colonization to study host responses to pathogens in the context of this understudied region.