Uncontrolled Hemorrhagic Shock Modeled via Liver Laceration in Mice with Real Time Hemodynamic Monitoring

Summary

Uncontrolled hemorrhage, an important cause of mortality among trauma patients, can be modeled using a standard liver laceration in a murine model. This model results in consistent blood loss, survival, and allows for testing hemostatic agents. This article provides the step-by-step process to perform this valuable model.

Abstract

Uncontrolled hemorrhage is an important cause of preventable deaths among trauma patients. We have developed a murine model of uncontrolled hemorrhage via a liver laceration that results in consistent blood loss, hemodynamic alterations, and survival.

Mice undergo a standardized resection of the left-middle lobe of the liver. They are allowed to bleed without mechanical intervention. Hemostatic agents can be administered as pre-treatment or rescue therapy depending on the interest of the investigator. During the time of hemorrhage, real-time hemodynamic monitoring via a left femoral arterial line is performed. Mice are then sacrificed, blood loss is quantified, blood is collected for further analysis, and organs are harvested for analysis of injury. Experimental design is described to allow for simultaneous testing of multiple animals.

Liver hemorrhage as a model of uncontrolled hemorrhage exists in the literature, primarily in rat and porcine models. Some of these models utilize hemodynamic monitoring or quantify blood loss but lack consistency. The present model incorporates quantification of blood loss, real-time hemodynamic monitoring in a murine model that offers the advantage of using transgenic lines and a high-throughput mechanism to further investigate the pathophysiologic mechanisms in uncontrolled hemorrhage.

Introduction

Trauma is the leading cause of death and disability among young people worldwide.¹ Uncontrolled hemorrhage remains a leading cause of mortality among severely injured trauma patients.² Management of the hemorrhaging trauma patient is two-fold: control of surgical bleeding, and resuscitation and replacement of lost blood.

Animal models of hemorrhagic shock have been the cornerstone in trauma research and can be used in the evaluation of the pathophysiology and treatment of traumatic/hemorrhagic shock.^3,4 Shock in animal models can be achieved broadly by two methods: controlled-hemorrhage and uncontrolled-hemorrhage.^5,6 Controlled-hemorrhage is performed by removal of a fixed volume of blood or by blood removal to achieve a certain blood pressure (fixed-pressure). While these models are useful in the evaluation in the mechanisms and immune alterations in hemorrhagic shock, they are not applicable to the testing of hemostatic agents and do not mimic the clinical scenario of hemorrhage following trauma. To this degree, we sought to develop a model of uncontrolled hemorrhage that would allow us to test hemostatic changes and pro-coagulant agents in a murine model. The liver is an attractive option for uncontrolled hemorrhage in part because of the dual blood supply to the liver and it is one of the most commonly injured intrabdominal organs in both blunt and penetrating trauma. Given the high clinical relevance, the liver has been utilized as a model of uncontrolled hemorrhage, most commonly in rat and porcine models but recently in primates as well.^{7,8,9,10,11,12} Murine models have also incorporated liver injury, such as a crush model or blunt trauma; however, these models do not result in hemorrhagic shock secondary to the liver injury.^13,14

The rat and porcine models of uncontrolled liver hemorrhage, while valuable in looking at resuscitation practices and hemodynamic monitoring, are less advantageous than a murine model for various reasons such as cost, number of animals utilized, and importantly the relative lack transgenic lines available for analysis of specific cellular and molecular signaling. The present murine model shares important similarities to existing liver hemorrhage models including standardized liver laceration, blood loss quantification, hemodynamic monitoring, and the ability to perform survival analysis. Many existing models only incorporate some of these aspects whereas our model was developed to measure many of the physiologic variables simultaneously and in multiple mice. As well, development of a murine model opens the door to investigations beyond resuscitation and into greater pathophysiology mechanisms in uncontrolled hemorrhage with the potential of a cost efficient, high-throughput model using advanced molecular techniques.

Protocol

Mice were housed in accordance with University of Pittsburgh (Pittsburgh, PA, USA) and National Institutes of Health (NIH; Bethesda, MD, USA) animal care guidelines in specific pathogen-free conditions with 12 h light-dark cycles and free access to standard feed and water. All animal experiments were approved and conducted in accordance with the guidelines set forth by the Animal Research and Care Committee at the University of Pittsburgh.

1. Surgical Field and Instrument Setup

1. Sterilize all surgical instruments, suture, gauze, cotton tip applicators, tubing, and tubing connectors prior to the procedure.
   1. Sterilize surgical instruments, suture, gauze, and cotton tip applicators in an autoclave. Sterilize tubing and tubing connectors with ethylene oxide.
2. Surgical field
   1. Turn on a water-circulating heating pad and set to 37 °C. Place a surgical blue pad on top of it and then a sterile drape on top of the surgical blue pad.
   2. Open all sterilized instruments onto the sterile drape. Use sterile gloves to avoid breaking sterility during this step.
   3. Fill a stainless steel bowl with 70% ethanol and set aside. This will be used to clean instruments between animals.
   4. Turn on microbead sterilizer and allow to heat to 150 °C. This will also be used to clean instruments between animals. If performing surgery on more than 5 mice, be sure to change instruments to a new sterile set.
3. Transducer Set-up
   1. Connect a sterile transducer, PE-50 tubing, two 23G needles, and male-male luer, and a three-way stopcock.⁶
   2. Calibrate and zero the transducer per manufacturer instructions.

2. Liver Laceration Surgical Procedure

1. Anesthesia Induction and Positioning
   1. Inject sodium pentobarbital intraperitoneally at dose of 70 mg/kg. Anesthesia should take effect between 5-10 min; assess depth of anesthesia with a toe pinch. If the mouse has response to the toe pinch, additional time or anesthesia is needed. If additional anesthesia is needed during the procedure, supplement sodium pentobarbital. Do not give supplements in larger quantities than 0.05 mL to prevent overdose.
   2. After the mouse is completely under anesthesia, position the mouse supine on a surgical board. Secure all four limbs of the mouse to the board with tape.
   3. Shave the abdomen and bilateral groins with a razor.
   4. Soak sterile gauze with betadine and apply to the abdomen and bilateral groins for surgery. For survival experiments, prep the abdomen and groins with betadine followed by ethanol for a total of three prep cycles.
   5. Insert a rectal temperature probe to monitor core temperature throughout the procedure. Keep the core temperature between 35-37 °C.
2. Femoral artery and venous cannulation
   1. For venous catheter set-up: fill PE-10 tubing, a 30 G needle, and a three-way stopcock with Lactated Ringer's solution from an IV bag.
   2. For arterial catheter set-up: fill PE-10 tubing and 30G needle with heparinized saline (1:10 dilution of 1,000 U heparin). Heparin-saline is required to prevent clotting.
   3. Place the mouse under a dissecting microscope.
   4. Make a 4-5 mm longitudinal incision over the groin muscle using surgical Iris scissors. Using Dumont forceps grab the adipose tissue connected to the adductor muscle and pull laterally for a clean exposure of the femoral bundle. Do not dissect through the adipose tissue as this will lead to vascular injury.
   5. Carefully dissect the nerve away from the artery and vein with the Dumont forceps. There is a fat pad adjacent to the nerve. Grab this with one Dumont forcep and pull laterally; this pulls the nerve away from the artery creating a plane for dissection. With other Dumont forcep bluntly dissect the connective tissue between the nerve and the artery.
      1. Do not grab the nerve during this part of the dissection.
   6. Loop three 6-0 silk sutures around the artery and vein proximal to the profunda femoris take off.
      1. Place suture 1 most proximally and leave loose.
      2. Place suture 2 most distally and tie immediately.
      3. Place suture 3 between Suture 1 and 2 and leave loose.
   7. Make an arteriotomy on the ventral surface of the vessel. Use of microvascular scissors is recommended to make the arteriotomy to avoid transection of the vessel.
   8. Insert the three-way catheter into the artery. Immediately tie off Suture 1 and 2 to secure the catheter in place.
   9. Connect the three-way catheter to the transducer and collect baseline blood pressure data.
   10. Repeat steps 2.2.4 - 2.2.6 on the opposite groin. Cannulate the femoral vein in a similar manner to the artery. Perform a venotomy on the ventral surface of the vessel followed by catheter insertion. This catheter can be utilized for fluid or drug administration.
3. Liver Laceration
   1. Pre-weigh a tube containing 0.5 mL of PBS, three absorption triangles, and one weigh boat per mouse.
   2. Make a ventral midline laparotomy incision starting at the xiphoid process and extending caudally to allow exposure of the liver completely.
   3. Insert one absorption triangle in the abdomen against the right abdominal wall. Repeat on the left side.
      1. Make sure the absorption triangle is away from the liver to avoid a packing hemostatic effect after the liver is lacerated.
   4. Carefully grab the left-middle lobe of the liver and lacerate 75% of the lobe with surgical Iris scissors. Place the lacerated segment in a tube containing PBS.
   5. Close the abdominal wall with staples via a staple applicator. Grasp the skin and muscle together and fire the staple. Do this as quickly as possible after the liver laceration to avoid blood loss outside of the abdomen. In survival experiments, the abdomen is closed in two layers. A running absorbable suture for the muscle followed by a running layer of non-absorbable suture for the skin provides adequate closure.
   6. For mice that are for survival timepoints longer than 30 min the femoral catheters should be removed, the artery and vein tied with suture 3 from step 2.2.6. Bilateral groins are then closed in two layers as described in the previous step.
   7. Following a specified time of interest for hemorrhage (30 min up to 72 h), remove the staples. Remove the absorption triangles and put into corresponding pre-weighed weigh boats. Use additional absorption triangles to soak up any unabsorbed blood.
   8. Weigh absorption filters to calculate total blood loss.
4. Post-operative Care
   1. Leave mice that are to be sacrificed at 30 min on the surgical board and under constant monitoring and under full anesthesia until the time of sacrifice. Mice are euthanized with a combination of cardiac puncture and an overdose of inhaled isoflurane.
   2. Place mice that are for longer survival time points in a recovery cage on top of a water-circulating heating pad. Constantly monitor the mice during recovery and do not leave unattended until they regain consciousness to maintain sternal recumbency. Return the mouse to cage space with other mice only once it has recovered from anesthesia.
   3. Administer post-operative analgesia with 0.1 mg/kg buprenex via subcutaneous injection once awoken from anesthesia and every 12 h after until the time of sacrifice.
   4. Allow mice free access to food and water after they are returned to their normal cages post-operatively.
   5. At the time of sacrifice for survival mice, anesthesia is accomplished with inhaled isoflurane. Once under anesthesia blood is collected via a right heart cardiac puncture, blood loss is recorded as described above and finally euthanasia is insured with an overdose of isoflurane.

Results

The liver laceration model results in reproducible and consistent blood loss in mice. Figure 1A demonstrates the consistent weight of lacerated liver that can be obtained with a standard deviation of only 0.02 g. This consistency in lacerated liver weight allows the ability to reproduce the model between mice and in different experimental set-ups such as different resuscitative protocols. As well, the reproducible weight of the lacerated liver, with a standard error of on...

Discussion

The murine liver laceration model described here provides a reliable, consistent model of uncontrolled hemorrhage. This model is straightforward to perform but there are important steps that require meticulous consideration. The most technically challenging part of the model is cannulation of the femoral vessels for hemodynamic monitoring and fluid/drug administration. Care must be taken during the dissection of the nerve and the arteriotomy/venotomy. It is important to not touch the nerve during the dissection of the ve...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
SS/45 dumonts	Fine Science Tools	11203-25
surgical scissors	Fine Science Tools	14068-12
hemostats	Fine Science Tools	13009-12
microscissors	Fine Science Tools	15000-08
0.8 mm curved forceps	Fine Science Tools	11009-13
suture reel 6-0	Fine Science Tools	18020-60
suture 4-0 silk w/ needle	Owens Minor	K188H
gauze 4 x 4	can be purchased through any global vendor
cotton-tip applicator	can be purchased through any global vendor
30 G needle	can be purchased through any global vendor
23 G needle	can be purchased through any global vendor
10 cc syringe	can be purchased through any global vendor
50 cc conical tube	can be purchased through any global vendor
1 cc syringe w/ 25G needle	Fisher Scientific	14-826-88
Polyethylene 10 tubing 100`(PE-10)	Fisher Scientific	14-170-12P
Polyethylene 50 tubing 100`(PE-50)	Fisher Scientific	14-170-12B
3-way stopcock	Fisher Scientific	NC9779127
surgical blue pad	Fisher Scientific	50-7105
Sterile Field dressings	Fisher Scientific	NC9517505
tape rolls 1"	Corporate Express	MMM26001
straight side wide mouth jars	VWR	159000-058
stainless steel tray 8" x 11"	VWR	62687-049
male-male leur lock 3-way	VWR	20068-909
sterilization pouch 3" x 8"	VWR	24008
sterilization pouch 5" x 10"	VWR	24010
absorption triangles	Fine Science Tools	18105-03
7 mm wound clip applier	Fisher Scientific	E0522687
1,000 7 mm wound clips	Fisher Scientific	E0522687
betadine (4 oz)	can be purchased through any global vendor
sterile gloves	can be purchased through any global vendor
eppendorfs	can be purchased through any global vendor
1/2 cc Lo-Dose insulin syringe	Fisher Scientific	12-826-79
small weigh boat	can be purchased through any global vendor
lactated ringers	can be purchased through any global vendor
hepranized saline solution (.1 µL; hep + 9.9 µL; NaCl)	can be purchased through any global vendor
phosphate buffered saline	can be purchased through any global vendor
pentobarbital	can be purchased through any global vendor
Wild M650 microscope w/ boom stand	Leica
Digi-Med BPA-400 analyzer & systems integrator	Micro-Med	SYS-400
TXD-310 (Digi-Med Transducer)	Micro-Med	TXD-300
Computer	Dell
microbead instrument sterilizer	VWR	11156-002
Oster A5 clippers w. size 40 blade	VWR	10749-020
circulating heating pad 18 x 26	Harvard	py872-5272
rectal thermometer	Kent Scientific	RET-3