The work of this manuscript was supported by funding to Dr. Neal by the Vascular Medicine Institute Pilot Project Program in Hemostasis and Vascular Biology (P3HVB) and the AAST Research Fellowship. This work is supported by U.S. National Institutes of Health grants 1 R35 GM119526-01 and UM1HL120877-01.

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
<ol>
	<li>Sterilize all surgical instruments, suture, gauze, cotton tip applicators, tubing, and tubing connectors prior to the procedure.
	<ol>
		<li>Sterilize surgical instruments, suture, gauze, and cotton tip applicators in an autoclave. Sterilize tubing and tubing connectors with ethylene oxide.</li>
	</ol>
	</li>
	<li>Surgical field
	<ol>
		<li>Turn on a water-circulating heating pad and set to 37 &#176;C. Place a surgical blue pad on top of it and then a sterile drape on top of the surgical blue pad.</li>
		<li>Open all sterilized instruments onto the sterile drape. Use sterile gloves to avoid breaking sterility during this step.</li>
		<li>Fill a stainless steel bowl with 70% ethanol and set aside. This will be used to clean instruments between animals.</li>
		<li>Turn on microbead sterilizer and allow to heat to 150 &#176;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.</li>
	</ol>
	</li>
	<li>Transducer Set-up
	<ol>
		<li>Connect a sterile transducer, PE-50 tubing, two 23G needles, and male-male luer, and a three-way stopcock.6</li>
		<li>Calibrate and zero the transducer per manufacturer instructions.</li>
	</ol>
	</li>
</ol>
2. Liver Laceration Surgical Procedure
<ol>
	<li>Anesthesia Induction and Positioning
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Shave the abdomen and bilateral groins with a razor.</li>
		<li>Soak sterile gauze with betadine and apply to the abdomen and bilateral groins for surgery.&#160;For survival experiments, prep the abdomen and groins with betadine followed by ethanol for a total of three prep cycles.</li>
		<li>Insert a rectal temperature probe to monitor core temperature throughout the procedure. Keep the core temperature between 35-37 &#176;C.</li>
	</ol>
	</li>
	<li>Femoral artery and venous cannulation
	<ol>
		<li>For venous catheter set-up: fill PE-10 tubing, a 30 G needle, and a three-way stopcock with Lactated Ringer&#39;s solution from an IV bag.</li>
		<li>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.</li>
		<li>Place the mouse under a dissecting microscope.</li>
		<li>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.</li>
		<li>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.
		<ol>
			<li>Do not grab the nerve during this part of the dissection.</li>
		</ol>
		</li>
		<li>Loop three 6-0 silk sutures around the artery and vein proximal to the profunda femoris take off.
		<ol>
			<li>Place suture 1 most proximally and leave loose.</li>
			<li>Place suture 2 most distally and tie immediately.</li>
			<li>Place suture 3 between Suture 1 and 2 and leave loose.</li>
		</ol>
		</li>
		<li>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.</li>
		<li>Insert the three-way catheter into the artery. Immediately tie off Suture 1 and 2 to secure the catheter in place.</li>
		<li>Connect the three-way catheter to the transducer and collect baseline blood pressure data.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Liver Laceration
	<ol>
		<li>Pre-weigh a tube containing 0.5 mL of PBS, three absorption triangles, and one weigh boat per mouse.</li>
		<li>Make a ventral midline laparotomy incision starting at the xiphoid process and extending caudally to allow exposure of the liver completely.</li>
		<li>Insert one absorption triangle in the abdomen against the right abdominal wall. Repeat on the left side.
		<ol>
			<li>Make sure the absorption triangle is away from the liver to avoid a packing hemostatic effect after the liver is lacerated.</li>
		</ol>
		</li>
		<li>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.</li>
		<li>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.&#160;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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Weigh absorption filters to calculate total blood loss.</li>
	</ol>
	</li>
	<li>Post-operative Care
	<ol>
		<li>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.&#160;Mice are euthanized with a combination of cardiac puncture and an overdose of inhaled isoflurane.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Allow mice free access to food and water after they are returned to their normal cages post-operatively.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Chang, R., Cardenas, J. C., Wade, C. E., Holcomb, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+the+understanding+of+trauma-induced+coagulopathy.">Advances in the understanding of trauma-induced coagulopathy.</a> Blood. 128 (8), 1043-1049 (2016).</li><li>Kutcher, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+paradigm+shift+in+trauma+resuscitation:+evaluation+of+evolving+massive+transfusion+practices.">A paradigm shift in trauma resuscitation: evaluation of evolving massive transfusion practices.</a> JAMA surgery. 148 (9), 834-840 (2013).</li><li>Tsukamoto, T., Pape, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+Models+for+Trauma+Research.">Animal Models for Trauma Research.</a> Shock. 31 (1), 3-10 (2009).</li><li>Darwiche, S. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pseudofracture:+an+acute+peripheral+tissue+trauma+model.">Pseudofracture: an acute peripheral tissue trauma model.</a> J Vis Exp. (50), (2011).</li><li>Lomas-Niera, J. L., Perl, M., Chung, C. -. S., Ayala, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shock+and+Hemorrhage:+an+Overview+of+Animal+Models.">Shock and Hemorrhage: an Overview of Animal Models.</a> Shock. 24, 33-39 (2005).</li><li>Kohut, L. K., Darwiche, S. S., Brumfield, J. M., Frank, A. M., Billiar, T. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fixed+volume+or+fixed+pressure:+a+murine+model+of+hemorrhagic+shock.">Fixed volume or fixed pressure: a murine model of hemorrhagic shock.</a> J Vis Exp. (52), (2011).</li><li>Matsuoka, T., Hildreth, J., Wisner, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Liver+injury+as+a+model+of+uncontrolled+hemorrhagic+shock:+resuscitation+with+different+hypertonic+regimens.">Liver injury as a model of uncontrolled hemorrhagic shock: resuscitation with different hypertonic regimens.</a> J Trauma. 39 (4), 674-680 (1995).</li><li>Komachi, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adhesive+and+Robust+Multilayered+Poly(lactic+acid)+Nanosheets+for+Hemostatic+Dressing+in+Liver+Injury+Model.">Adhesive and Robust Multilayered Poly(lactic acid) Nanosheets for Hemostatic Dressing in Liver Injury Model.</a> J. Biomed. Mater. Res. Part B Appl. Biomater. , (2016).</li><li>Orfanos, N. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+antioxidants+on+a+porcine+model+of+liver+hemorrhage.">The effects of antioxidants on a porcine model of liver hemorrhage.</a> J Trauma Acute Care Surg. 80 (6), 964-971 (2016).</li><li>Morgan, C. E., Prakash, V. S., Vercammen, J. M., Pritts, T., Kibbe, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+validation+of+4+different+rat+models+of+uncontrolled+hemorrhage.">Development and validation of 4 different rat models of uncontrolled hemorrhage.</a> JAMA Surgery. 150 (4), 316-324 (2015).</li><li>Rosselli, D. D., Brainard, B. M., Schmiedt, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficacy+of+a+topical+bovine-derived+thrombin+solution+as+a+hemostatic+agent+in+a+rodent+model+of+hepatic+injury.">Efficacy of a topical bovine-derived thrombin solution as a hemostatic agent in a rodent model of hepatic injury.</a> Can J Vet Res. 14 (14), 303-308 (2015).</li><li>Sheppard, F. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+Nonhuman+Primate+(Rhesus+Macaque)+Model+of+Uncontrolled+Traumatic+Liver+Hemorrhage.">Development of a Nonhuman Primate (Rhesus Macaque) Model of Uncontrolled Traumatic Liver Hemorrhage.</a> Shock. 44, 114-122 (2015).</li><li>Nemzek-Hamlin, J. A., Hwang, H., Hampel, J. A., Yu, B., Raghavendran, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+murine+model+of+blunt+hepatic+trauma.">Development of a murine model of blunt hepatic trauma.</a> Comp Med. 63 (5), 398-408 (2013).</li><li>Vogel, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Platelet-derived+HMGB1+is+a+critical+mediator+of+thrombosis.">Platelet-derived HMGB1 is a critical mediator of thrombosis.</a> J Clin Invest. 125 (12), (2015).</li><li>Modery-Pawlowski, C. L., Tian, L. L., Ravikumar, M., Wong, T. L., Sen Gupta, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+and+in+vivo+hemostatic+capabilities+of+a+functionally+integrated+platelet-mimetic+liposomal+nanoconstruct.">In vitro and in vivo hemostatic capabilities of a functionally integrated platelet-mimetic liposomal nanoconstruct.</a> Biomaterials. 34 (12), 3031-3041 (2013).</li></ol>

The authors have no financial competing interests to declare.

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...

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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>SS/45 dumonts</td>
			<td>Fine Science Tools</td>
			<td>11203-25</td>
		</tr>
		<tr>
			<td>surgical scissors</td>
			<td>Fine Science Tools</td>
			<td>14068-12</td>
		</tr>
		<tr>
			<td>hemostats</td>
			<td>Fine Science Tools</td>
			<td>13009-12</td>
		</tr>
		<tr>
			<td>microscissors</td>
			<td>Fine Science Tools</td>
			<td>15000-08</td>
		</tr>
		<tr>
			<td>0.8mm curved forceps</td>
			<td>Fine Science Tools</td>
			<td>11009-13</td>
		</tr>
		<tr>
			<td>suture reel 6-0</td>
			<td>Fine Science Tools</td>
			<td>18020-60</td>
		</tr>
		<tr>
			<td>suture 4-0 silk w/ needle</td>
			<td>Owens Minor</td>
			<td>K188H</td>
		</tr>
		<tr>
			<td>gauze 4x4</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>cotton-tip applicator</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>30G needle</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>23G needle</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>10cc syringe</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>50cc conical tube</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>1cc syringe w/ 25G needle</td>
			<td>Fisher Scientific</td>
			<td>14-826-88</td>
		</tr>
		<tr>
			<td>Polyethylene 10 tubing 100`(PE-10)</td>
			<td>Fisher Scientific</td>
			<td>14-170-12P</td>
		</tr>
		<tr>
			<td>Polyethylene 50 tubing 100`(PE-50)</td>
			<td>Fisher Scientific</td>
			<td>14-170-12B</td>
		</tr>
		<tr>
			<td>3-way stopcock</td>
			<td>Fisher Scientific</td>
			<td>NC9779127</td>
		</tr>
		<tr>
			<td>surgical blue pad</td>
			<td>Fisher Scientific</td>
			<td>50-7105</td>
		</tr>
		<tr>
			<td>Sterile Field dressings</td>
			<td>Fisher Scientific</td>
			<td>NC9517505</td>
		</tr>
		<tr>
			<td>tape rolls 1&#34;</td>
			<td>Corporate Express</td>
			<td>MMM26001</td>
		</tr>
		<tr>
			<td>straight side wide mouth jars</td>
			<td>VWR</td>
			<td>159000-058</td>
		</tr>
		<tr>
			<td>stainless steel tray 8&#34; x 11&#34;</td>
			<td>VWR</td>
			<td>62687-049</td>
		</tr>
		<tr>
			<td>male-male leur lock 3-way</td>
			<td>VWR</td>
			<td>20068-909</td>
		</tr>
		<tr>
			<td>sterilization pouch 3&#34;x8&#34;</td>
			<td>VWR</td>
			<td>24008</td>
		</tr>
		<tr>
			<td>sterilization pouch 5&#34;x10&#34;</td>
			<td>VWR</td>
			<td>24010</td>
		</tr>
		<tr>
			<td>absorption triangles</td>
			<td>Fine Science Tools</td>
			<td>18105-03</td>
		</tr>
		<tr>
			<td>7mm wound clip applier</td>
			<td>Fisher Scientific</td>
			<td>E0522687</td>
		</tr>
		<tr>
			<td>1000 7mm wound clips</td>
			<td>Fisher Scientific</td>
			<td>E0522687</td>
		</tr>
		<tr>
			<td>betadine (4oz)</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>sterile gloves</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>eppendorfs&#160;</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>1/2cc Lo-Dose insulin syringe</td>
			<td>Fisher Scientific</td>
			<td>12-826-79</td>
		</tr>
		<tr>
			<td>small weigh boat</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>lactated ringers</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>hepranized saline solution (.1&#181; hep + 9.9&#181;NaCl)</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>phosphate buffered saline&#160;</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>pentobarbital&#160;</td>
			<td colspan="2">can be purchased through any global vendor</td>
		</tr>
		<tr>
			<td>Wild M650 microscope w/ boom stand</td>
			<td>Leica</td>
			<td></td>
		</tr>
		<tr>
			<td>Digi-Med BPA-400 analyzer &#38; systems integrator</td>
			<td>Micro-Med</td>
			<td>SYS-400</td>
		</tr>
		<tr>
			<td>TXD-310 (Digi-Med Transducer)&#160;</td>
			<td>Micro-Med</td>
			<td>TXD-300</td>
		</tr>
		<tr>
			<td>Computer</td>
			<td>Dell</td>
			<td></td>
		</tr>
		<tr>
			<td>microbead instrument sterilizer</td>
			<td>VWR</td>
			<td>11156-002</td>
		</tr>
		<tr>
			<td>Oster A5 clippers w. size 40 blade</td>
			<td>VWR</td>
			<td>10749-020</td>
		</tr>
		<tr>
			<td>circulating heating pad 18x26</td>
			<td>Harvard</td>
			<td>py872-5272</td>
		</tr>
		<tr>
			<td>rectal thermometer</td>
			<td>Kent Scientific</td>
			<td>RET-3</td>
		</tr>
	</tbody>
</table>

uncontrolled hemorrhagic shock modeled via liver laceration in mice with real time hemodynamic monitoring

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.

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...

Watch this Scientific Journal Video about Uncontrolled Hemorrhagic Shock Modeled via Liver Laceration in Mice with Real Time Hemodynamic Monitoring at JoVE.com

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

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.

Uncontrolled hemorrhage is an important cause of preventable deaths among trauma patients. We have developed a murine model of uncontrolled hemorrhage via ...

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Medicine

8.5K Views.  University of Pittsburgh. 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.

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

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

It is common knowledge that severe blood loss and traumatic injury can lead to a cascade of detrimental signaling events often resulting in mortality. 1, 2, 3, 4, 5 These signaling events can also lead to sepsis and/or multiple organ dysfunction (MOD). 6, 7, 8, 9 It is critical then to investigate the causes of suppressed immune function and detrimental signaling cascades in order to develop more effective ways to help patients who suffer from traumatic injuries. 10 This fixed pressure Hemorrhagic Shock (HS) procedure, although technically challenging, is an excellent resource for investigation of these pathophysiologic conditions. 11, 12, 13 Advances in the assessment of biological systems, i.e. Systems Biology have enabled the scientific community to further understand complex physiologic networks and cellular communication patterns. 14 Hemorrhagic Shock has proven to be a vital tool for unveiling these cellular communication patterns as they relate to immune function. 15, 16, 17, 18 This procedure can be mastered! This procedure can also be used as either a fixed volume or fixed pressure approach. We adapted this technique in the murine model to enhance research in innate and adaptive immune function. 19, 20, 21 Due to their small size HS in mice presents unique challenges. However due to the many available mouse strains, this species represents an unparalleled resource for the study of the biologic responses. The HS model is an important model for studying cellular communication patterns and the responses of systems such as hormonal and inflammatory mediator systems, and danger signals, i.e. DAMP and PAMP upregulation as it elicits distinct responses that differ from other forms of shock. 22, 23, 24, 25 The development of transgenic murine strains and the induction of biologic agents to inhibit specific signaling have presented valuable opportunities to further elucidate our understanding of the up and down regulation of signal transduction after severe blood loss, i.e. HS and trauma 26, 27, 28, 29, 30. 
There are numerous resuscitation methods (R) in association with HS and trauma. 31, 32, 33, 34 A fixed volume resuscitation method of solely lactated ringer solution (LR), equal to three times the shed blood volume, is used in this model to study endogenous mechanisms such as remote organ injury and systemic inflammation. 35, 36, 38 This method of resuscitation is proven to be effective in evaluating the effects of HS and trauma 38, 39.

The Hemorrhagic Shock model has been a reliable and reproducible resource facilitating the identification and understanding of signaling cascades associated with inflammation and end-organ damage after trauma. This article provides a step-by-step description of surgical and mechanical aspects associated with the Hemorrhagic Shock experimental procedure in mice.

It is common knowledge that severe blood loss and traumatic injury can lead to a cascade of detrimental signaling events often resulting in mortality. 1, ...

Controlled Cervical Laceration Injury in Mice

Use of genetically modified mice enhances our understanding of molecular mechanisms underlying several neurological disorders such as a spinal cord injury (SCI). Freehand manual control used to produce a laceration model of SCI creates inconsistent injuries often associated with a crush or contusion component and, therefore, a novel technique was developed. Our model of cervical laceration SCI has resolved inherent difficulties with the freehand method by incorporating 1) cervical vertebral stabilization by vertebral facet fixation, 2) enhanced spinal cord exposure, and 3) creation of a reproducible laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm in depth without associated contusion. Compared to the standard methods of creating a SCI laceration such as freehand use of a scalpel or scissors, our method has produced a consistent lesion. This method is useful for studies on axonal regeneration of corticospinal, rubrospinal, and dorsal ascending tracts.

A novel technique to create a reproducible in vivo model of cervical spinal cord laceration injury in the mouse is described. This technique is based on spine stabilization by fixation of the cervical facets and laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm.

Use of genetically modified mice enhances our  understanding of molecular mechanisms underlying several neurological disorders  such as a spinal cord ...

Real-time Cytotoxicity Assays in Human Whole Blood

A live cell-based whole blood cytotoxicity assay (WCA) that allows access to temporal information of the overall cell cytotoxicity is developed with high-throughput cell positioning technology. The targeted tumor cell populations are first preprogrammed to immobilization into an array format, and labeled with green fluorescent cytosolic dyes. Following the cell array formation, antibody drugs are added in combination with human whole blood. Propidium iodide (PI) is then added to assess cell death. The cell array is analyzed with an automatic imaging system. While cytosolic dye labels the targeted tumor cell populations, PI labels the dead tumor cell populations. Thus, the percentage of target cancer cell killing can be quantified by calculating the number of surviving targeted cells to the number of dead targeted cells. With this method, researchers are able to access time-dependent and dose-dependent cell cytotoxicity information. Remarkably, no hazardous radiochemicals are used. The WCA presented here has been tested with lymphoma, leukemia, and solid tumor cell lines. Therefore, WCA allows researchers to assess drug efficacy in a highly relevant ex&#160;vivo condition.

The whole blood cytotoxicity assay (WCA) is a cytotoxicity assay developed by incorporating high-throughput cell positioning technology with fluorescence microscopy and automated image processing. Here, we describe how lymphoma cells treated with an anti-CD20 antibody can be analyzed real-time in human whole blood to provide quantitative cellular cytotoxicity analysis.

A live cell-based whole blood cytotoxicity assay (WCA) that allows access to temporal information of the overall cell cytotoxicity is developed with ...

Monitoring of Systemic and Hepatic Hemodynamic Parameters in Mice

The use of mouse models in experimental research is of enormous importance for the study of hepatic physiology and pathophysiological disturbances. However, due to the small size of the mouse, technical details of the intraoperative monitoring procedure suitable for the mouse were rarely described. Previously we have reported a monitoring procedure to obtain hemodynamic parameters for rats. Now, we adapted the procedure to acquire systemic and hepatic hemodynamic parameters in mice, a species ten-fold smaller than rats. This film demonstrates the instrumentation of the animals as well as the data acquisition process needed to assess systemic and hepatic hemodynamics in mice. Vital parameters, including body temperature, respiratory&#160;rate&#160;and heart&#160;rate were recorded throughout the whole procedure. Systemic hemodynamic parameters consist of carotid artery pressure (CAP) and central venous pressure (CVP). Hepatic perfusion parameters include portal vein pressure (PVP), portal flow rate as well as the flow rate of the common hepatic artery (table 1). Instrumentation and data acquisition to record the normal values was completed within 1.5 h. Systemic and hepatic hemodynamic parameters remained within normal ranges during this procedure.
This procedure is challenging but feasible. We have already applied this procedure to assess hepatic hemodynamics in normal mice as well as during 70% partial hepatectomy and in liver lobe clamping experiments. Mean PVP after resection (n= 20), was 11.41&#177;2.94 cmH2O which was significantly higher (P&#60;0.05) than before resection (6.87&#177;2.39 cmH2O). The results of liver lobe clamping experiment indicated that this monitoring procedure is sensitive and suitable for detecting small changes in portal pressure and portal flow rate. In conclusion, this procedure is reliable in the hands of an experienced micro-surgeon but should be limited to experiments where mice are absolutely needed.

This film demonstrates how to acquire systemic and hepatic hemodynamics in mice. The whole monitoring includes acquisition of vital parameters, systemic blood pressure, central venous pressure, common hepatic artery flow rate, and portal vein pressure as well as the portal flow rate in mice.

The use of mouse models in experimental research is of enormous importance for the study of hepatic physiology and pathophysiological disturbances. ...

Real-time Pressure-volume Analysis of Acute Myocardial Infarction in Mice

Acute myocardial infarction can lead to acute heart failure and cardiogenic shock. The evaluation of hemodynamics is critical for the evaluation of any potential therapeutic approach directed against acute left ventricular (LV) dysfunction. Current imaging modalities (e.g., echocardiography and magnetic resonance imaging) have several limitations since data on LV pressure cannot directly be measured. LV catheterization in mice undergoing coronary artery occlusion could serve as a novel method for a real-time evaluation of LV function.
At the beginning of the procedure, mice were anesthetized followed by endotracheal intubation. For LV catheterization, the right carotid artery was exposed via middle-neck incision. The catheter was introduced and placed into the LV cavity. Left thoracotomy was conducted and the left main coronary artery (LCA) was ligated. To induce reperfusion, the suture was released after 45 min. Pressure-volume data was recorded at all times.
Ligation of the LCA caused a decrease in LV systolic function as evidenced by a 30% reduction in stroke volume, LV ejection fraction (EF), and cardiac output. Maximum dP/dt as a parameter for LV contractility was also significantly reduced and diastolic function was severely impaired (minimum dP/dt -40%). Reperfusion over a period of 20 min did not lead to a complete recovery of LV function.
Real-time pressure-volume analysis served as a valid procedure for monitoring cardiac function during acute myocardial infarction in mice. Maintaining stable anesthesia and a standardized surgical approach was crucial to ensure valid results. As the early phase of acute myocardial infarction is critical for morbidity and mortality, the delineated method could be beneficial for preclinical evaluation of new strategies for cardioprotection.

Acute myocardial infarction in mice induces acute but incompletely characterized changes in left ventricular (LV) function. LV catheterization in mice undergoing coronary artery occlusion serves as a novel method for a real-time evaluation of LV function.

Acute myocardial infarction can lead to acute heart failure and cardiogenic shock. The evaluation of hemodynamics is critical for the evaluation of any ...

Complete and Partial Aortic Occlusion for the Treatment of Hemorrhagic Shock in Swine

Hemorrhage remains the leading cause of preventable deaths in trauma. Endovascular management of non-compressible torso hemorrhage has been at the forefront of trauma care in recent years. Since complete aortic occlusion presents serious concerns, the concept of partial aortic occlusion has gained a growing attention. Here, we present a large animal model of hemorrhagic shock to investigate the effects of a novel partial aortic balloon occlusion catheter and compare it with a catheter that works on the principles of complete aortic occlusion. Swine are anesthetized and instrumented in order to conduct controlled fixed-volume hemorrhage, and hemodynamic and physiological parameters are monitored. Following hemorrhage, aortic balloon occlusion catheters are inserted and inflated in the supraceliac aorta for 60 min, during which the animals receive whole-blood resuscitation as 20% of the total blood volume (TBV). Following balloon deflation, the animals are monitored in a critical care setting for 4 h, during which they receive fluid resuscitation and vasopressors as needed. The partial aortic balloon occlusion demonstrated improved distal mean arterial pressures (MAPs) during the balloon inflation, decreased markers of ischemia, and decreased fluid resuscitation and vasopressor use. As swine physiology and homeostatic responses following hemorrhage have been well-documented and are like those in humans, a swine hemorrhagic shock model can be used to test various treatment strategies. In addition to treating hemorrhage, aortic balloon occlusion catheters have become popular for their role in cardiac arrest, cardiac and vascular surgery, and other high-risk elective surgical procedures.

Here, we present a protocol demonstrating a hemorrhagic shock model in swine that uses aortic occlusion as a bridge to definitive care in trauma. This model has application in testing a wide range of surgical and pharmacological therapeutic strategies.

Hemorrhage remains the leading cause of preventable deaths in trauma. Endovascular management of non-compressible torso hemorrhage has been at the ...

Measurement of the Hepatic Venous Pressure Gradient and Transjugular Liver Biopsy

Here we provide a detailed protocol describing the clinical procedure of hepatic venous pressure gradient (HVPG) measurement in patients with advanced chronic liver disease followed by an instruction for transjugular biopsy. Under local anesthesia and ultrasound guidance, a catheter introducer sheath is placed in the right internal jugular vein. Using&#160;fluoroscopic guidance, a balloon catheter is advanced into the inferior vena cava (IVC) and inserted into a large hepatic vein. Correct and sufficient wedge position of the catheter is ensured by injecting contrast media while the balloon is blocking the outflow of the cannulated hepatic vein. After calibrating the external pressure transducer, continuous pressure recordings are obtained with triplicate recordings of the wedged hepatic venous pressure (WHVP) and free hepatic venous pressure (FHVP). The difference between FHVP and WHVP is referred to as HVPG, with values &#8805;10 mm Hg indicating clinically significant portal hypertension (CSPH). Before removing the catheter, pressure readings obtained in the IVC at the same level, as well as the right atrial pressure are recorded.
Finally, a transjugular liver biopsy can be obtained via the same vascular route. Different systems are available; however, core biopsy needles are preferred over aspiration needles, especially for cirrhotic livers. Again, under fluoroscopic guidance a biopsy needle introducer sheath is advanced into an hepatic vein. Next, the transjugular biopsy needle is gently advanced through the introducer sheath: (i) in case of aspiration biopsy, the needle is advanced into the liver parenchyma under aspiration and then removed quickly, or (ii) in case of a core biopsy, the cutting-mechanism is triggered inside the parenchyma. Several separate passages can be safely performed to obtain sufficient liver specimens via transjugular biopsy. In experienced hands, the combination of these procedures takes about 30-45 min.

Here, we present a protocol for measurement of hepatic venous pressure gradient (HVPG),the gold standard to diagnose clinically significant portal hypertension. Moreover, we describe how to perform a transjugular liver biopsy within the same session.

Here we provide a detailed protocol describing the clinical procedure of hepatic venous pressure gradient (HVPG) measurement in patients with advanced ...

Standardized Hemorrhagic Shock Induction Guided by Cerebral Oximetry and Extended Hemodynamic Monitoring in Pigs

Hemorrhagic shock ranks among the main reasons for severe injury-related death. The loss of circulatory volume and oxygen carriers can lead to an insufficient oxygen supply and irreversible organ failure. The brain exerts only limited compensation capacities and is particularly at high risk of severe hypoxic damage.This article demonstrates the reproducible induction of life-threatening hemorrhagic shock in a porcine model by means of calculated blood withdrawal. We titrate shock induction guided by near-infrared spectroscopy&#160;and extended hemodynamic monitoring to display systemic circulatory failure, as well as cerebral microcirculatory oxygen depletion. In comparison to similar models that primarily focus on predefined removal volumes for shock induction, this approach highlights a titration by means of the resulting failure of macro- and microcirculation.

Hemorrhagic shock is a severe complication in seriously injured patients, which leads to life-threatening oxygen undersupply. We present a standardized method to induce hemorrhagic shock via blood withdrawal in pigs that is guided by hemodynamics and microcirculatory cerebral oxygenation.

Hemorrhagic shock ranks among the main reasons for severe injury-related death. The loss of circulatory volume and oxygen carriers can lead to an ...

Real-Time Monitoring of Neurocritical Patients with Diffuse Optical Spectroscopies

Neurophysiological monitoring is an important goal in the treatment of neurocritical patients, as it may prevent secondary damage and directly impact morbidity and mortality rates. However, there is currently a lack of suitable non-invasive, real-time technologies for continuous monitoring of cerebral physiology at the bedside. Diffuse optical techniques have been proposed as a potential tool for bedside measurements of cerebral blood flow and cerebral oxygenation in case of neurocritical patients. Diffuse optical spectroscopies have been previously explored to monitor patients in several clinical scenarios ranging from neonatal monitoring to cerebrovascular interventions in adults. However, the feasibility of the technique to aid clinicians by providing real-time information at the bedside remains largely unaddressed. Here, we report the translation of a diffuse optical system for continuous real-time monitoring of cerebral blood flow, cerebral oxygenation, and cerebral oxygen metabolism during intensive care. The real-time feature of the instrument could enable treatment strategies based on patient-specific cerebral physiology rather than relying on surrogate metrics, such as arterial blood pressure. By providing real-time information on the cerebral circulation at different time scales with relatively cheap and portable instrumentation, this approach may be especially useful in low-budget hospitals, in remote areas and for&#160;monitoring in open fields (e.g., defense and sports).

Presented here is a protocol for non-invasively monitoring cerebral hemodynamics of neurocritical patients in real-time and at the bedside using diffuse optics. Specifically, the proposed protocol uses a hybrid diffuse optical systems to detect and display real-time information on cerebral oxygenation, cerebral blood flow and cerebral metabolism.

Neurophysiological monitoring is an important goal in the treatment of neurocritical patients, as it may prevent secondary damage and directly impact ...

Organ Ischemia-Reperfusion Injury by Simulating Hemodynamic Changes in Rat Liver Transplant Model

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, including ischemia-reperfusion injury (IRI) of extrahepatic organs. This model requires numerous experiments and devices. The duration of anhepatic phase is closely related to the time to develop IRI after transplantation. In this experiment, we used hemodynamic changes to induce extrahepatic organ damage in rats and determined the maximum tolerance time. The time until the most severe organ injury varied for different organs. This method can easily be replicated and can also be used to study IRI of the extrahepatic organs after liver transplantation.

This paper provides a detailed description of how to build an animal model of the anhepatic phase (liver ischemia) in rats to facilitate basic research into ischemia-reperfusion injury after liver transplantation.

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, ...