The overall goal of this procedure is to transurethral induce mouse urinary tract infection. This is accomplished by first growing bacterial strains of interest and diluting cultures to the desired concentration. The second step of the procedure is to transurethral inoculate the mouse bladder with the bacterial strains of interest.
The third step of the procedure is to collect urine samples and harvest kidneys and bladders. The final step of the procedure is inoculation of auger plates with urine and homogenized kidney and bladder samples. Ultimately, results can be obtained that show successful infection of the urinary tract based on the presence of bacterial colonies in plated samples.
Hi, I'm Michael Shea and welcome to my lab in the Department of Urology at the Stanford University School of Medicine. I am Annu ti also from the Shea lab. Today we will show you a procedure for mouse transurethral catheterization.
We use this procedure in our lab to study the induction of urinary tract infection in mice. So let's get started. In a lanar flow hood, cut a 30 centimeter length of polyethylene tubing, secure 30 gauge needle on a sterile three cubic centimeter syringe.
Use sterile forceps to pick up one end of the cut tubing and slide it onto the needle until it meets the hub. Cut the tubing so that approximately two centimeters extend beyond the tip of the needle. This helps to prevent injury to mice during catheterization.
Remove the catheter from the syringe and place it into a sterile petri dish. Make one catheter for every mouse to be infected. Sterilize the catheters with ultraviolet irradiation for at least 30 minutes.
Seal a dish with perfil to keep the catheter sterile for long-term storage. Streaking appropriate auger plate with an inoculum of frozen stock of the bacterial strain of interest. Here, an inoculum of U pathogenic E coli or UPEC is streaked out for overnight growth at 37 degrees Celsius.
Pick single colonies with the proper morphology, which in the case of UPEC are solid yellowish, translucent colonies on LB auger and place in at least five milliliters of liquid broth for overnight culture at 37 degrees Celsius with agitation at 200 revolutions per minute. Add 10%of the first broth culture to fresh, medium, and grow overnight. Again, to enhance type one pill eye expression, which is associated with euro pathogenesis in vivo, measure the OD 600 of the culture to determine the number of colony forming units.
Abbreviated CFU per milliliter of culture. Dilute the culture with sterile one XPBS so that the final concentration is between 10 to the sixth and 10 to the eighth, CFU in 10 to 50 microliters to ensure that inocular are not voided out immediately. After installation, deprive mice of water for 30 minutes before anesthesia, scruff the mouse and gently press the lower abdomen to encourage voiding.
Place the anesthetized mouse on its back with its head inserted in a nose cone connected to isof fluorine containing oxygen. Massage the abdomen to evacuate any remaining urine from the bladder. Use a spray bottle to soak the lower abdomen and perineum with 70%ethanol.
Next, use a sterile one milliliter syringe to draw up the bacterial inoculum. Secure a prepared sterile catheter on the syringe. Use scissors to cut the tubing so that extends only one to two millimeters beyond the tip of the needle.
This helps to prevent injury to mice During catheterization, tap the syringe and depress the plunger. To remove air bubbles, lubricate the catheter by dipping it into a dollop of bacterial static lubricating jelly. Be sure to use a different dollop of jelly for each bacterial strain that will be tested under a dissecting stereo microscope.
Use fine toothed forceps in the non-dominant hand to gently grasp the clitoral hood and raise it to expose the deep pink urethral ATU in the dominant hand. Grasp the syringe with attached catheter. With a pencil grip, engage the tip of the catheter in the urethral ATU at a 45 degree angle relative to the mouse's abdomen.
In the midline sagittal plane. Stretch the clitoral hood along the long axis of the syringe. Effectively loading the urethra onto the catheter.
The catheter should slide easily into the bladder if necessary. Gently twirl the catheter to help navigate the redundant folds of the urethra. When the catheter is seated up to the hub, lower the syringe parallel to the mouse's abdomen.
Drop the forceps and use the non-dominant hand to stabilize the catheter and syringe. Slowly inject the inoculum over at least five seconds. If fluid is seen flowing around the catheter during injection, the bladder is full or the catheter is in the vagina.
Stop the injection and examine the catheter position. Reposition the catheter in the urethra if necessary. After the injection, slowly withdraw the catheter.
Keep the mouse under anesthesia for half an hour after inoculation. To reduce the risk of voiding after surgery, allow the mouse to recover on a clean warming blanket with continuous observation to confirm the return of normal breathing patterns and activity. Once it is fully recovered, the ability to crawl, return the mouse to a bedding containing cage.
Depending on the combination of bacterial and mouse strains, it may take between six hours and week for a full infection. To develop, place a sterile Petri dish on ice for each infected mouse over the course of several hours. Scruff the mouse over the dish to collect urine for culture.
50 microliters of urine should be sufficient. Pipee the urine into a micro centrifuge tube, cap it and keep the urine on ice. Under the dissecting scope, place a euthanized mouse on its back on a dissection tray.
Spray the abdomen with 70%ethanol. Use scissors to make a three to four centimeter incision in the skin and underlying abdominal wall. If necessary, use pins to retract the abdominal wall away from the abdominal contents.
If earlier attempts at urine collection were unsuccessful, use a 30 gauge needle and syringe to aspirate urine directly through the bladder wall. Use forceps and scissors to remove the kidneys. Transfer them to a sterile Petri dish and mince each one into at least four pieces, approximately two to three millimeters each in size.
Transfer each kidney fragment to a cryo vile containing five to six sterile 0.5 millimeters, zirconium oxide beads, and 400 microliters of sterile PBS. Keep the kidney samples on ice. Use forceps and scissors to remove the bladder.
Mince the bladder using the same method As for the kidneys. Transfer each bladder fragment to a cryo vial containing 1.6 millimeter stainless steel beads and 200 microliters of sterile PBS and keep the tubes on ice. Place the vials in a bullet blender Homogenizer homogenize the tissue at speed setting eight for at least one minute.
It is common for some tissue fragments to remain after homogenization. All samples should be homogenized for the same amount of time in a microtiter plate. Make serial tenfold dilution of the collected urine and tissue homogenate with ice.
Cold PBS spread. The dilution over. Appropriate auger plates grow overnight at 37 degrees Celsius.
The CFU per organ or milligram of tissue or milliliter of urine can be back calculated from the number of colonies on each plate. We've been using these techniques to study how mutations in bacterial stress responses affect the ability of UPEC to cause UTI in vivo, as well as resist antibiotics given to infected mice. This table shows the ability of wild type UPEC to infect the mouse bladder after the transurethral inoculation.
We've just shown you how to transversely induce urinary tract infections in mice. When doing this procedure, it's important to catheterize the mice gently. Proper technique is associated with easy catheterization but may require practice.
So that's it. Thanks for watching and good luck with your experiments.
