Our protocol couples an accessible method for preparing tesaglitazar-loaded liposomes with a thorough approach for assessing liposome uptake at a cellular level. This technique provides an unbiased method for assessing the uptake of fluorescently-labeled liposomes that are prepared using low temperatures and easy to acquire equipment. To prepare fluorescence-labeled liposomes loaded with calcium acetate and tesaglitazar, first use a half inch probe to emulsify an ether chloroform solution of lipids by sonication for 30 seconds at room temperature, 20 kilohertz, and 50%power.
Immediately transfer the homogenized water and oil emulsion solution into a rotary evaporator with a special adapter, nanometer gauge, and pressure regulator valve and connect the evaporator to a vacuum line. To remove the organic solvents, set the rotation rate to 100 revolutions per minute and the vacuum to 0.5 atmospheres. After a gel forms and disappears, increase the vacuum to 0.9 atmospheres and filter the liposome aqueous dispersion back and forth through a 200 nanometer pour polycarbonate filter in a liposome extruder equipped with two gas-tight syringes multiple times.
After verifying the size distribution by dynamic light scattering, use a two milliliter spin column to remove any unentrapped tesaglitazar from the liposomes and use a spectrophotometer to determine the entrapped drug concentration. Then add no more than 500 microliters of liposome sample to the dry column gel bed. To administer the liposomes by retro-orbital injection, gently press down on the skin above and below the exposed eye to lift the eye above the plane of the face.
Then carefully insert the tip of the needle at the front corner of the eye, making sure that the needle is below the orb without touching it, and slowly inject the liposomes into the retro-orbital space. To harvest the blood sample, at the appropriate experimental endpoint carefully make an incision in the skin at the edge of the caudal end of the mouse's rib cage and cut a one centimeter straight line up towards the head of the animal until the pectoralis muscles are exposed. At the initial incision site, make two small cuts perpendicular to the line towards the head and carefully cut away the pectoralis muscle on one side of the ribcage in the exposed area to allow better access to, and visualization of, the needle insertion site.
Insert the needle between the third and fourth ribs keeping it as close to the center line of the rib cage as possible, and gently pull up on the syringe to begin collecting blood. When as much volume as possible has been collected, transfer the blood to a micro centrifuge tube containing 10 microliters of 0.5 molar EDTA on ice. To extract the inguinal post-tissue pad from each side of the mouse, use one set of forceps to hold the peritoneal membrane to the right side and the second set of forceps to hold the edge of the skin above the peritoneal membrane on the other.
Gently pull the skin away from the membrane to separate the layers from each other. Pin down the outer edge of the skin to better access the adipose depot and locate the inguinal adipose tissue depot along the skin. And locate the inguinal lymph node in the center of the adipose depot using forceps and scissors to remove the lymph node as needed.
After the lymph node has been removed, use the forceps to carefully grasp the end of the adipose depot nearest to the pinned point and begin making small cuts at the connective membrane between the adipose tissue and the skin. Lift the adipose tissue away from the skin while making cuts for better access to the membrane, and place the depot in a polyethylene vial containing HEPES buffer on ice. To extract the epididymal adipose depots from the caudal end of the peritoneal cavity, use forceps to gently pull the first epididymal adipose depot from the dorsal end of the mouse and locate the epididymis and vas deferens attached to this depot.
Then carefully cut between the adipose depot and the epididymis and vas deferens to separate the adipose from the other tissues and place the depo in a polyethylene vial with HEPES buffer on ice. When all of the tissues have been harvested, use one to two pairs of scissors to mince the adipose tissue samples into 0.5 millimeter tissue fragments. Next, add 1.5 milliliters of two milligrams per milliliter collagenase buffer to the vials of adipose tissue pieces and place the vials into a shaking incubator at 37 degrees Celsius and 150 revolutions per minute for 30 to 45 minutes.
After 30 minutes, use a one milliliter pipette to triturate the samples several times. If the tissue pieces are still too large for easy pipetting, return the samples to the incubator for an additional 15 minutes. Once the samples are fully digested, triturate the fragments another 10 times to ensure the creation of a single cell suspension and strain each sample through individual 70 micrometer filters into one 50 milliliter tube per sample.
Rinse each file with five milliliters of FACS buffer to collect any remaining cells and filter each wash into the appropriate vial of cell sample on ice. When all of the samples have been filtered, collect the cells by centrifugation and aspirate the adipocyte supernatant from the samples. Next, carefully aspirate the supernatants leaving the stromal vascular fraction pellets intact, and resuspend the pellets in one milliliter of FACS buffer per tube.
Then transfer the suspensions into new 1.5 or 1.7 milliliter tubes on ice until flow cytometric analysis by standard protocols. Dynamic light scattering reveals an average liposome diameter of 163.2 nanometers and a zeta potential of minus 19.2 millivolts. Cryogenic electron microscopy imaging reveals circular liposomes with a relatively small standard deviation obtained from the average diameter as measured by dynamic light scattering.
Flow cytometric analysis of samples from PBS-treated mice can be used as a DiD fluorescence minus one control with which to create DiD positive gates. As illustrated in this representative analysis, a 10 milligram per milliliter liposome loading concentration of DiD is too high, surpassing the quantifiable range of the cytometer in all three tissues tested. A 0.1 milligram per milliliter concentration demonstrates some signal, but a clear population beyond the PBS-treated cells is not observed.
Therefore, the one milligram per milliliter concentration, which demonstrates a clear distinction from the PBS control while still being readable by the analyzer, is recommended for use in liposome preparation. Slightly different gating strategies are required for each tissue type, but most of the same cell types can be identified in each. Once specific populations of interest are identified, the total size of each cell population and the frequency at which they are DiD positive can be quantified.
Preparing reagents and equipment and anticipating outcomes for each step is crucial. Unexpected results early on may ruin future outcomes. Histological methods can be used to assess liposome localization within whole tissue, while imaging flow cytometry can be used to verify liposome internalization on a per-cell basis.
