The overall goal of this procedure is to image leishmania parasite infections in a live host. This is accomplished by first generating transgenic bioluminescent parasites. The second step of the procedure is to infect mice with leishmania parasites.
The third step of the procedure is bioluminescent imaging of the infected mice with IVIS. The final step of the procedure is analyzing the bioluminescent data. Ultimately, results can be obtained that are quantitative measures of bioluminescence generated by the transgenic leishmania parasites in live mice through the use of IVIS.
This method allows the relative parasite burden to be analyzed at various anatomical locations and in the same animal during longitudinal studies. Hi, I'm Colin Fer, a student in the immunology graduate program at the University of Iowa. I work in the laboratory of Dr.Mary Wilson.
Hi, I'm Mary Wilson. We're in the Departments of Internal Medicine microbiology and Epidemiology at University of Iowa. We also work at the Iowa City VA Medical Center.
Hi, I'm Lori, love Holman. I'm also in the Wilson Lab. Today we'll show you a procedure for imaging bioluminescent leishmania parasites in a live animal host using an in vivo imaging system.
We use this procedure in our laboratory to study the infectious disease model leishmaniasis in mice. So let's get started. Parasites are grown in vitro and passage through animals to maintain virulence.
The parasite species dose and infection root are determined by the investigator. We will demonstrate two infection routes, an intradermal infection in the ear pinner, and an intravenous infection via the tail. For the intradermal infection, the mouse must first be sedated with an intraperitoneal injection of liquid anesthesia.
Restrain the animal manually by holding it firmly by its dorsal skin with its abdomen up and head pointed down. Clean the injection site with 70%Ethel or isopropyl alcohol. Position the needle with the bevel up and slightly angled.
The tip of the needle should just slightly penetrate the lower region of the abdominal wall during the injection. Once the animal is sedated, clean the infection site with alcohol. Load a syringe with 10 microliters of parasites.
Place a protective cover over the index finger. In this case a common 50 milliliter conical tube is used. Then place the mouse on its back and hold the ear between the thumb and protective cover.
Insert the needle at a shallow angle into the dermis of the ear pinner, and slowly depress the syringe plunger. The skin may appear to bubble slightly. There should not be any fluid found on the bottom side of the ear because that would indicate that the needle has penetrated through the ear pinner.
To perform an intravenous tail inoculation, first load a syringe with 200 microliters of parasites. Next, place the mouse under a heat lamp for a few minutes and then transfer it to a restraining device. Place a protective cover over the index finger.
Hold the animal's tail between the thumb and protective cover and keep it relatively taut. Clean the injection site with alcohol. Then insert the needle into one of the two lateral tail veins where the tail round over the protective finger cover at relatively the same angle.
As the tail slowly depress the syringe plunger, the vein will appear to flush clear. There should be no difficulty depressing the plunger or bubbling of the skin unless the needle bevel is outside of the vein. To prepare the Lucifer for injection reconstitute Lucifer in KO's PBS with or without magnesium and calcium to a concentration of 15 milligrams per milliliter syringe filter the Lucifer with a 0.2 micrometer filter.
Aliquots of the D Lucifer can be stored at minus 80 degrees until use prior to injection warmed the Lucifer to 37 degrees Celsius in a water bath. Clean the injection site on the animal with alcohol. Inject the mouse intraperitoneal with 15 milligrams per milliliter.
Lucifer solution in DPBS at a dose of 150 milligrams per kilogram approximately 10 to 15 minutes after the Lucifer injection. The animals can be imaged with the IVIS first anesthetize the animals by placing them in a holding chamber and exposing them to isof fluorine. Transfer the anesthetized animals to the imaging chamber and position them so that the nose cones attached to the manifold will deliver a continuous and regulated flow of isof fluorine.
Open the living image software program and initialize the IVIS For our experiments. We use the IVIS 200 system. The CCD camera will be automatically called and maintained at the appropriate temperature after initializing.
The IVIS. The camera settings and image acquisition parameters are set in the IVIS system control window. Important parameters include imaging mode, exposure, time spinning focus, and subject height, F stop and field of view.
Press acquire in the IVIS system control window to initiate data acquisition for a single image or press acquire continuous photos for multiple images. For our experiments, we use the 62nd default exposure setting. A closeup image may provide greater resolution, but not necessarily enhanced sensitivity compared to an image taken using a larger field of view.
After the imaging procedure, remove the animals from the imaging chamber and return them to their cage. Monitor the animals until they recover from the anesthesia. The luminescence data is analyzed using the living image software and corresponds to the light intensity expressed as the number of photons detected by the CCD camera.
These data are represented by a pseudo color image that is overlaid onto a black and white photograph of the animals. Specific areas of the image may be analyzed by creating regions of interest. One of the simplest ways to analyze the data is to select a region of interest and measure the average number of photons per second that is detected.
These data can then be easily exported to a spread and subsequently used to generate graphs. An example of a five week infection with a luciferase expressing parasite causing human visceral leishmaniasis leishmania. In phantom chaga ssu slash ir one SAT dash LUC is shown here.
After five weeks of infection, visceral parasites can be detected in the livers and bones of all mice, and in the spleens of mice one and two. Preliminary kinetic studies are essential to maximize the detection of photons emitted from the bioluminescent parasites. In the experiment just shown, a sequence of images from the infected mice was collected every two minutes after Lucifer injection, the magnitude of light emitted is quantified and graft.
To determine the maximum detection point IVIS imaging technology can be used to assess the efficiency of parasite inoculation. An example is shown here in which bulb sea mice were inoculated with a visceral species of leishmania. One hour after infection.
Mice were analyzed with IVIS. This first image is a black and white photograph of the mice taken immediately before the luminescence signal was measured. The red arrows indicate the injection site on each animal.
In this next image, the pseudo color image of the luminescence was overlaid on the photograph. Regions of interest indicated by the red circles were selected and the luminescence was quantified in each region of interest. To produce this graph, the data represent the average luminescence of infected region of interest minus the region of interest of mock infected.
IVIS can also be used to monitor the course of infection in mouse models of ly manasis. In this example bulb, sea mice were inoculated subcutaneously on day zero with 10 to the power of five. Leishmania Mexicana expressing luciferase and imaged on the sequential weeks after infection.
Representative images at 10 to 31 weeks after infection are shown here. The same data is also shown in quantitative form. This scatterplot represents net regions of interest at various time points post infection.
We've just shown you how to image T transgenic English mania parasites in a live animal host using an in vivo imaging system. Remember, when doing this procedure, it's important to wear proper protective clothing and use safety equipment to minimize potential health risks to you and your animals. So that's it.
Best of luck with your experiments. And a special thanks to the University of Iowa's central microscopy core facility and the Iowa City VA Animal Care facility.
