by the core staff will include blood collection, routine health measurements including regular body weight measurements, quantitative fecal collections when required, and oversight of all breeding activities and housing needs in collaboration with the animal care staff. A technician familiar with the experimental protocols related to each project has been assigned by project, and this technical staff member will be responsible for the specific duties for that project. This group of technicians will benefit from interacting with Dr. Wallace and with other staff members to make the husbandry aspects of the entire program project more efficient. In many cases, the diets to be fed to the mice are made especially for the individual project, and the technical staff of the core will provide the liason function between the project, the diet kitchen, and the Diet Analysis laboratory that provides quality control assays, assuring timely delivery of diet. C. Personnel and Responsibilities Jeanne Wallace, D.V.M., Dipl. A.C.L.A.M., is the leader of the Animal/Surgical Core. Dr. Wallace serves as the Director of the Animal Resources Program (ARP) at the Bowman Gray Campus and has over 15 years of experience in working with experimental animals. Dr. Rudei will participate as required and will assist Dr. Wallace in oversight of the surgeries, as he has many years of experience with these procedures. Ms. Janet Sawyer is a Research Assistant with over 15 years of experience in the evaluation of atherosclerosis. She developed the techniques to evaluate aortic atherosclerosis in mice. She is also an experienced small animal surgeon who has worked with Dr. Rudel for over 20 years. Under the guidance of Drs. Wallace and Rudel, she will perform the surgeries required for each of the three surgical protocols. Mr. Ramesh Shah is a Research Assistant who has worked with Dr. Rudel for over 25 years. He has designed the mouse liver perfusion set-up that now has been used successfully in over 20 consecutive liver perfusions, and he will carry out all of the liver perfusion experiments for projects 1, 4, and 5. Ms. Kathryn Kelley is a highly experienced technician with over 10 years of experience in animal work. She will be responsible for all procedures done in project 1, including blood sample collections. She also will be responsible for scheduling and assisting with liver perfusion and thoracic lymph duct cannulation experiments for project 1. She will supervise the breeding and maintenance of the mouse colony as required for project 1. Ms. Li Hou is an experienced technician and will oversee the breeding and maintenance of the transgenic mice needed for project 2. PHS 398/2590 (Rev. 05/01) Page "_5_ Principal Investigator/Program Director (Last. first, middle): Rudel. Lawrence L. Ms. Elena Boudyguina is an experienced technician with many years of experience working with mice, and she will oversee the breeding and maintenance of the mice needed for project 3. Ms. Manal Zabalawi is a veteran technician, who has worked with mice with Dr. Sorci-Thomas for over three years, and she will oversee the breeding and maintenance of the mice for project 4. She will be responsible for scheduling and assisting with the liver perfusion experiments for project 4. Ms. Jamie Haywood is an experienced technician who has been working with mice in project 5 for about 2 years, and she will oversee the breeding and maintenance of the mice for project 5. She will be responsible for scheduling and assisting with the liver perfusion and intestinal/bile duct cannulation experiments for project 5. D. Methods Methods used by the Core are described below. All animal care procedures are carried out in full compliance with state and federal animal welfare laws and the standards and policies of the Department of Health and Human Services. Our laboratory animal medicine facility is fully accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC). All procedures have been approved by the Institutional Animal Care and Use Committee. Diet Preparation: Each diet has a diet formulation card listing all pertinent identifying information and the amounts of each ingredient per batch. As each ingredient is weighed and added to the mixing pot, it is checked off on the card. Weighed portions of most fats are melted over low heat on an electric range and care is taken to minimize oxidation of fats. If crystalline cholesterol is required, it is added to the already melted fat just above the melting point for cholesterol (about 150[unreadable]F). Weighed portions of dry ingredients are placed in a large stainless steel mixing pot and mixed with a heavy- duty commercial mixer for about 1 minute. The melted fats are then added and mixed for an additional 1 to 2 minutes. A measured amount of water is added last and mixing continues for about 2 to 3 minutes until the ingredients are thoroughly blended. The diet attains the consistency of dough and is then divided into portions, wrapped in waxed paper or sealed in plastic bags, packed in a box labeled with the diet number, date and experiment number and is then transferred for storage into a freezer at -20[unreadable]C. The diet formulation card is dated, a log is kept of all diets prepared, and a sample of each diet prepared on a given day is sent to Dr. Martha Wilson in the Diet Analysis Laboratory for analysis of cholesterol and fatty acid content before the diet is fed as a quality control procedure. Animal Care and Housing: The number of mice that will be in each study are the minimum number required to obtain statistically significant differences among groups based on power calculations described in the write- up for each project. In addition, a number of mice will be used as breeders, as the mouse models of this program are not available from commercial sources and must be raised in our facility. Support for the expenses of these breeding efforts is requested within this core. The veterinary care of the animals will be supervised by Jeanne Wallace, D.V.M. She will be assisted in this activity by the additional veterinary faculty of the Animal Resources Program, post- DVM fellows and residents, and certified laboratory animal technicians. Costs for these serv=ces are included in the animal per diem rate. Ketamine/Xylazine administered IM is used for restraint in mice for blood sampling and the surgical procedures as described below. Blood Sampling: For small blood samples from the mice, intraorbital blood collection is routinely done on anesthetized mice. In some cases, terminal blood samples are needed, and these samples are taken via heart puncture while the mice are anesthetized with Ketamine/xylazine. Data Recording and Sample Coding: When blood samples are collected, data sheets are prepared listing the study number, investigator, location of the mice sampled, date of sample collection, analyses requested, strain designation, mouse numbers and sequential sample numbers. The sample tubes are labeled with PHS 398/2590 (Rev. 05/01) Page _ Principal Investi.qat0r/ProqramDirector (Last, first, middle): Rudel, Lawrence L. the sequential sample numbers, study number, and date of collection. Although this Core is responsible for verifying assignment of animals to groups in addition to collecting research data, there is no chance for bias. All samples are collected in a standard fashion and according to established schedules. Experimental Surgery Each procedure is done while the mice are anesthetized with ketamine/xylazine (150 & 30 mg/kg body weight). [unreadable] The lymph duct cannulation protocol is as follows: A midline laparotomy is performed after the mouse is anesthetized. The mesenteric lymph ducts are full of chylomicrons and easily visualized if the surgery is started by 8:30 AM. These lymph ducts lie on the left side of the dorsal abdomen on either side of the mesenteric artery and are exposed by gently retracting the small intestine. A small PE-10 cannula is placed through an incision in the mesenteric duct and tied in place with 6-0 silk suture. A second silastic cannula is sutured into the duodenum. The cannulae are tunneled through the right lateral abdominal walland exteriorized through a stab wound in the skin in the middle of the back at the level of the 6u' vertebrae. The abdomen is closed with silk suture and the mouse placed in a restraining cage. After the mouse is fully awake, liquid diet consisting of nonfat dry milk solids suspended in water containing vegetable oil (n-6 PUFA) as 10% of energy is administered through the duodenal cannula. About 0.1 ml of diet mixed with radiolabeled sterol (about 5 pCi) dissolved in ethanol is infused at a rate of 0.3 ml/hour. Infusion of additional diet is continued at the same rate for the remainder of the study. The infusion rate of liquid diet is estimated to represent the number of calories that would be consumed over 24 hours. In typical experiments, lymph collection will be carried out for 6 to 8 hours after which animals will be sacrificed for tissue collection. [unreadable] The liver perfusion protocol is as follows: The mouse liver perfusion set-up is based on over 20 years of experience with primate liver perfusion. The mouse liver perfusion system contains all of the components of the monkey liver system, including a reservoir made with a 10 ml syringe to attached via silastic tubing to a peristaltic pump. The tubing from the pump is connected to a Hamilton lung which is coiled silastic tubing in a small plastic container continuously gassed with oxygen, and the tubing from the lung then passes through a heat exchanger, a clot filter, a bubble trap, finally connecting to the portal vein at the liver. The total volume in the tubing of this system is about 5 ml. A total of 10 mi of perfusate is used for each liver. The perfusate is Krebs-Henseleit original Ringer bicarbonate containing 1% human serum albumin, 0.1% (w/v) glucose and 0.2% (v/v) complete amino acid mix with glutamine. Perfusate is made to a 10% hematocrit with mouse red blood cells isolated fresh from citrated blood and washed the day prior to the experiment. The liver is cannulated with inflow through the portal vein and outflow through the superior vena cava, and perfusate is circulated through the liver at a rate of 1.5 ml/min. Recirculating perfusion is done in situ in a closed chamber in which humidity and temperature (37 [unreadable]) are maintained constant throughout the experiment. For experiments in which bile is collected, the gall bladder is tied off and the bile duct is also cannulated with PE10 tubing. During bile collection, taurocholate is added at a constant rate to the perfusate, which is 62.5 nmole/min, a rate that represents the physiologic rate. Typical experiments are carried out for 3 hours. [unreadable] The surgical preparation for intestinal bile acid perfusion and bile duct cannulation is as follows: Mice will be fasted overnight, and then anesthetized. The jugular vein will be cannulated with a PE-10 catheter connected to an infusion pump. For maintenance of hydration, the mice will be infused intravenously with 0.9% saline at 200 pl/h. After laparotomy, the bile duct will be cannulated. After exposing the liver and gallbladder, the lower end of the common bile duct will be ligated and the common bile duct cannulated below the entrance of the cystic duct using a PE-10 polyethylene catheter. After catherization, the cystic duct will be ligated and cholecystectomy will be performed. A PE-10 catheter will then be inserted into the duodenum 5 mm below the pylorus and secured with 6-0 sutures and cryoacrylate adhesive. The duodenal catheter will be externalized through the abdominal wall and connected to a second infusion pump. During surgery and bile collection, mouse body temperature is maintained at 37 [unreadable] C with a heat lamp and temperature probe. Continuous anesthesia will be maintained with periodic i.m. injections of ketamine/xylazine. Hepatic bile will be collected by PHS 398/2590 (Rev. 05/01) Page _ PrinciDal Investiqator/Proqram Director (Last, first, middle): Rudel, Lawrence L. gravity every 15 min into preweighed tubes. The mice will be euthanised at the end of the experiment. Atherosclerosis Evaluations: Atherosclerosis evaluations of the mouse aorta have been done according to standard protocols developed in this program project. To study the extent and severity of aortic atherosclerosis in mice, the heart and aorta will be removed in block at the time of necropsy. Three different atherosclerosis endpoints are compared. 1. Evaluation of aortic surface atherosclerosis: At the time of necropsy the aorta is cleaned of adventitia, pinned flat on a black wax board, and submersion fixed in 10% neutral buffered formalin. The image of the aortic surface is captured using a Scion LG-3 Scientific Frame Grabber interfaced to a Hitachi color video camera. Areas of aortic surface with lesions and total aortic surface are digitized using a Wacom digitizing tablet and Scion Image software (version 1.62a). Atherosclerosis extent is expressed as percent of intimal surface covered with lesion. 2. Histologic evaluation of aortic atherosclerosis: To study the extent and severity of aortic atherosclerosis using morphometric techniques, 3 mm blocks will be cut in cross section at specified sites from the aortic arch, and the proximal abdominal aorta just distal to the celiac artery. The tissue blocks will be dehydrated through increasing concentrations of ethanol and embedded in paraffin. One 5 pm section will be cut from each block to provide a cross-sectional view of the artery and stained with Verhoeff van Gieson stain. The artery image from the stained slide will be captured using a Scion LG-3 Scientific Frame Grabber interfaced to a Nikon Labophot 2 microscope. Utilizing the Scion Image 1.62a version of NIH Image software and a Wacom digitizing tablet, measurements of intimal area (lesion area), maximum intimal thickness (lesion thickness), internal and external elastic lamina length and area (artery size) and areas of calcification and necrosis will be made using a Macintosh G4 computer. Percent lumen stenosis (amount of artery occlusion) and mean intimal thickness (average lesion thickness based on artery size) will be calculated from raw measurements. 3. Evaluations of aortic atherosclerosis expressed as cholesteryl ester accumulation: Using a dissecting microscope, the fixed aorta will be cleaned of all adventitia and blocks will be cut as described above. The remaining aorta will be used for analysis of cholesterol by GLC, before and after saponification, and the amount of esterifled cholesterol will be calculated by difference. Results will be expressed as milligrams of free or esterified cholesterol per gram of artery protein. E. Facilities Available The Animal Resources Program facilities, located at the Bowman Gray Campus, consist of 32,000 sq.ft, of space on the 7th floor of the Hanes Research Building and the adjacent floor of the Nutrition Research building and 5,500 sq. ft. of space on the Ground and E floors of the Gray Building. In addition to animal housing and procedure rooms, the facilities include the following special support laboratories. Necropsy Laboratory - Terminal liver perfusions and necropsies will be performed in this laboratory located on the Ground floor of the Gray Building. This laboratory is equipped with a stainless steel necropsy table, stainless steel benches, a Castle portable operation light, Masterflex peristaltic pump and a Technicon Autoanalyzer heat exchanger. A Wise model 650 double headed surgical microscope is also available in this room. Diet Laboratory - The Diet Laboratory is located at the CMCRC in Building 14 and consists of separate rooms for ingredient storage, diet preparation and diet storage. In the ingredient storage room, dry ingredients are stored on shelves or palates and ingredients requiring refrigeration are kept in a Kelvinator commercial refrigerator. The diet preparation area is equipped with Sartorius and Mettler balances for accurate measuring of ingredients and a Hobart Commercial Diet Mixer to ensure thorough mixing of diets. Prepared food is stored at -20[unreadable]C in one of eighteen diet storage freezers located in the diet storage room on the Ground floor of Building 14. Diets are shipped to the Hawthorne Campus on a weekly basis and stored in freezers on the 7th floor until needed. Diet Analysis Laboratory - The diet analysis laboratory is combined with the lipoprotein analytic laboratory in 450 sq. ft of space. Available for use are homogenizers, heating blocks, drying PHS 398/2590 (Rev. 05/01) Page _ _"!
NIH ExPorter
NIH ExPorter