Fixed Volume or Fixed Pressure: A Murine Model of Hemorrhagic Shock

To characterize the immunologic responses to hypovolemic shock, a surgical procedure is performed in which a mouse's vessels are dissected and catheterized. While monitoring the animal's physiology through digital recording and physical observation, hemorrhagic shock is induced to mimic a traumatic experience following induced shock. The mouse is resuscitated, the catheter is removed and the animal is allowed to recover.

The mouse can then be studied to assess changes in organ function, cellular communication patterns, and genetic expression through an array of techniques such as flow cytometry, hematology, immuno blots, immunohistochemistry, or gene chip microarray array. The main advantage of this technique over existing methods is that this model of fixed volume or fixed pressure hemorrhagic shock, allows the researcher the opportunity to control the degree of shock while continuously monitoring and accurately recording physiologic fluctuations. This specific method of hemorrhagic shock enhances the researcher's ability to study and understand immunologic function In preparation for the surgery.

Ensure that the following sterile equipment and supplies are on hand. Six zero sutures, cotton and tip applicators gauze male, male, three-way stopcocks transducers, 18 inch pieces of PE, 50 tubing, and five inch pieces of PE 10 tubing. Surgical tools you will need are three-way stop cocks, A one cc syringe, a 10 cc syringe, a 23 gauge needle, a 30.5 gauge needle, micro scissors, fine forceps, a surgical packet, forceps, hemostats, and surgical scissors.

Additional supplies are Lactated ringer solution and alcohol Prep. A metal board, a sterile field, sterile gloves, Betadine, sterile saline, heparinized saline, and anesthesia. On the day of the surgery, set up the right leg mirroring catheter used to measure blood pressure wearing sterile gloves.

Grab the middle of a five inch piece of sterile PE 10 tubing with both hands and stretch it about one inch to help with catheter insertion. Using the sterile scissors, cut the tubing in half, beveling the stretched end. Then insert a 30 gauge needle into the blunt end of the unstretched PE 10 tubing.

Using an alcohol wipe, sterilize the top of a 10 cc sterile vial containing the heparinized saline solution. Fill a one cc syringe with 0.6 to 0.7 cubic centimeters of the heparin solution. Attach a 30 gauge needle and catheter to the end of a three-way that is directly across from the male end.

Fill the stopcock 30 gauge needle and PE tubing with the heparin solution, making sure there are no air bubbles. Remove air bubbles by flicking, then remove the 30 gauge needle from the three-way and withdraw fluid in the three-way back to the one CC syringe to remove any bubbles that are trapped in the three-way and reattach the needle with attached tubing. Approximately 0.1 cubic centimeters of the mixture should remain in the syringe.

To set up the left leg mirroring catheter, follow the procedure just described, leaving out the three-way stop cock. First wipe the lactated ringers bag with alcohol. Then hook up a sterile transducer to a blood pressure and heart rate machine according to the micro me specifications.

Next, attach a three-way stop cock to both ends of the transducer and lay it flat on the bench top. Fill a 10 cc syringe with lactated ringer solution and attach it to the three-way. Insert a 23 gauge needle into both ends of an 18 inch piece of sterile PE 50 tubing.

Attach one end of the PE tubing to the three-way with the 10 cc syringe attached. Then fill the three-way and PE 50 setup with lactated ringer solution. Make sure to remove all air bubbles as before.

Next, reattach the three-way to the transducer and fill it. And a second three-way with lactated ringers. Finally, attach the metal male, male lure lock stopcock to the 23 gauge needle of the PE 50 tubing for attachment to the right leg mirroring catheter.

The instruments are now ready for use in the surgical and experimental procedures to dissect and catheterize the mouse's vessels in preparation for induction of shock. Begin by placing a sodium Penta barbital anesthetized mouse onto the metal surgical plate. In the supine position, immobilize the animal by a loose loop tape technique, which entails loosely wrapping thin strips of tape around each of the limbs inferior to the pore and sticking them down to the board using osteo a five clippers with the size 40 blade, shave the animals abdominal and inguinal areas.

Then using gauze, wipe the area with Betadine and alcohol to sterilize it. Place a nose cone with one cubic centimeter revise of fluorine over the mouse's nose. Once the animal's respirations begin to slow, test the depth of anesthesia using the limb extension withdrawal reflex.

Place a sterile field dressing over the animal throughout the procedure to help maintain sterility. Then make a small four to five millimeter incision in the skin parallel to the left internal oblique muscle of the abdomen and the left transverse abdominis muscle. Taking care not to damage the surrounding muscles or touch nerves.

Use dumont to grab the tissue at the abdominal connection and separate the adipose tissue from the oblique and transverse abdominal muscles. Using the other pair of Als blunt dissect along the abdominal muscles teasing away fascia and adipose tissue. Just beneath this adipose tissue, like the femoral vein and artery, along with the femoral nerve without touching it, dissect away the femoral nerve by grabbing the adipose tissue that lies next to it.

Pull this tissue laterally from the vein and artery. The nerve which is embedded in this tissue will follow as the nerve is pulled laterally. Place the other dumont point down against the artery and open and close them.

To blunt dissect the facia. The vessels are very superficial, so be sure not to dig into the underlying muscles. After the nerve is separated, keep the dumont closed and slip them dorsal to the vessels.

As the tip of the dumont appears on the other side of the vein, open them to blunt dissect the fascia. Keep the dumont dorsal to the vessel and grab the first suture. Then pull the suture back through the opening made between the vessels and the underlying muscles.

Put a total of 3 6 0 sutures around the vein and artery suture. One is the most proximal to the abdominal muscle tie knot, but leave it loose and clamp it with a hemostat. The concave edge of the hemostat should rest on the animal's abdominal cavity.

Suture two is most distal in location. Immediately tie off this suture to ligate the vessels and clamp it with a hemostat concave side down the distal and proximal sutures are used to pull the vessels tort to prevent blood loss and lift them a bit to aid in catheter insertion. Suture three should be placed between the distal and proximal sutures.

For catheter support, tie a loose knot that will be used to secure the catheter inside the vessel after insertion. After the sutures are secure, identify the artery by the thick vessel wall and white color. Using the micro scissors, make a small incision on the top of the artery close to the distal suture so that there is an ample amount of artery for the initial catheter insertion.

Place one end of the dumont into the arterial vessel lumen and close them over the vessel wall to open the hole while holding the arterial wall. Push the catheter into the lumen while pulling the vessel over the catheter. Lightly tie down the middle support suture to hold the catheter in place.

Then release the proximal hemostat to loosen the proximal suture and reopen the suture around the vessels. At this point, the arterial pressure should push blood back into the catheter. Pulsating blood should be visible in the catheter.

Hold the vessels around the catheter with one al and use the other to push the catheter into the vessel. Four to five millimeters. The tip of the catheter should rest just beneath the internal oblique and transverse abdominis muscles.

Hook the animal up to a BPA 400 analyzer and flush the arterial lines. Put one or two drops of sterile saline into the surgical opening to keep the surrounding tissue moist during the hemorrhagic shock and resuscitation procedures to induce hemorrhagic shock. Begin by placing the anesthetized mouse under a lamp and on a circulating heat pad to help maintain the temperature of 36 to 37 degrees Celsius.

Next, use a syringe to withdraw approximately half of the mouse's blood volume over a period of 15 minutes to achieve a mean arterial pressure of 28 to 32 millimeters of mercury. After 1.5 to three hours, resuscitate the animal using a syringe pump to dispense lactated ringer solution at three times the shed volume at a constant rate over 15 minutes. Once the animal has been resuscitated, remove the catheter and ligate the vessels.

Using three sutures, pull the catheter just past the proximal suture, then tie this suture completely off to prevent blood loss. Collateral flow prevents the hind limbs from becoming ischemic. Finally, sew up both hind limb openings with a sterile four zero PDSI.

I suture. Remove the loose loop tape and place the animal into a clean cage, which is kept on a circulating heat pad for several hours. Post recovery.

This diagram represents an overview of the hemorrhagic shock procedure. After bilateral cannulation and initial connection to the blood pressure monitor, a five minute baseline of blood pressure is attained. Once a reliable blood pressure is recorded, the induction of shock occurs where blood is withdrawn from the femoral catheter to obtain a blood pressure of 28 plus or minus two millimeters of mercury, the withdrawal of blood should occur over a 15 minute timeframe.

After the induction of shock, a two and a half hour shock period occurs where constant monitoring of the blood pressure in shoes. After the shock period, a controlled resuscitation occurs using a syringe pump that infuses three times the shed blood volume back into the animal. Finally, the animal is reconnected to the blood pressure monitors to attain another baseline evaluation.

This occurs for approximately five to 10 minutes. This figure denotes the pro and anti-inflammatory responses associated with hemorrhagic shock. A cytokine storm occurs where pro and anti-inflammatory cytokines, along with macrophages T cells and a plethora of other physiologic responses interact in a feedback loop in an attempt to return the body to homeostatic conditions repairing oxygen depleted organs.

This graphic representation denotes liver function in wildtype mice subject to two hemorrhagic shock time points compared to sham mice, alanine aminotransferase or a LT level is shown in graph A and aspartate amino transferase or a ST level is shown. In graph B.Liver enzymes were measured using a blood chemistry analyzer from blood plasma. These fluorescence microscopy images show lung tissue in wild type mice that have been subjected to hemorrhagic shock and control.Mice.

Foid in stain in a actin is shown in green and dappy. Staining of the nucleus is shown in blue. iNOS expression is shown in red as can be seen here.

Hemorrhagic shock mice have a distinct infiltration of iNOS, which is shown in red into the bronchial epithelium and vascular endothelium of lung tissue. This staining is absent and control mice which underwent a sham procedure. These pictor micrographs show sections of liver tissue from mice subjected to hemorrhagic shock or control mice that were fixed in para formaldehyde and stained with hematin or eoin.

The section of tissue from the mice subjected to hemorrhagic shock showed necrotic tissue as a result of depleted circulation and oxygenation that was not seen in mice that underwent the sham procedure. When attempting this technique, it's very important to remember to be very patient and deliberate with your surgical technique and hemorrhage. And keep in mind this model can be combined with other traumatic models such as bilateral femur fracture, pseudo fracture, and soft tissue trauma in order to answer questions related to the immunology of multiple traumas.