Name: fixed volume or fixed pressure: a murine model of hemorrhagic shock
Uploaded: 2011-06-06T12:00:00
Duration: 16 min 31 s
Description: Watch this Scientific Journal Video about Fixed Volume or Fixed Pressure: A Murine Model of Hemorrhagic Shock at JoVE.com

Funding Source/Number Molecular Biology of Hemorrhagic Shock GM053789

1. Instrument and Surgical Field Preparation:
1. Instrument Preparation.
All surgical procedures are performed using aseptic techniques. A surgical blue pad and sterile field dressing are used. All materials and instruments are sterilized before use. 6-0 Suture, cotton tip applicators, gauze, male-male 3-way stopcocks, and instruments are autoclave sterilized. Transducers, PE-50, and PE-10 tubing are Ethylene oxide sterilized. All 3-way stopcocks, syringes and needles are received sterile.
6-0 Suture is cut into 1-inch pieces (6 pieces / animal) and put into small 
sterilization pouches. Cotton-tip applicators, 4x4 gauze squares, and male-male 3-way stopcocks are put into either small or medium sized sterilization pouches and autoclaved. Our surgical instruments are autoclave sterilized each evening. They are washed after surgery using antibacterial soap and tap water. They are allowed to dry on a clean surgical blue pad. They are then carefully placed into a sterilization pouch and sterilized for use the next day.
Since the transducers and tubing have plastic components, they must be 
sterilized using gas, i.e. Ethylene Oxide. PE-10 tubing is cut into 5-inch pieces and placed into a small sterilization pouch. PE-50 tubing is cut into 18-inch pieces and placed into a medium sterilization pouch. 

2. Surgical Field.
To set-up the surgical field, first, turn on the hot bead sterilizer to ensure it reaches the appropriate temperature&#x2013; 300-350&deg;F before starting surgeries. Then proceed to the next steps by placing surgical blue pads down on an alcohol wiped benchtop. One pad goes under the microscope and the other goes on the circulating heating pad where the BP analyzers are located. Place a sterile field dressing over both surgical blue pads. Fill a stainless steel instrument tray 1/3 of the way with 70% alcohol. Use enough 70% EtOH to cover all surgical instruments. Use a separate sterile field dressing and place it next to the microscope. Place all sterile instruments, suture, gauze, and catheters on this sterile field. Be careful when opening sterile instruments and suture Not to contaminate them by touching them. It is best to use sterile gloves when doing this setup procedure.

2. Mechanical Set-up and Procedures:
1. Catheter Set-up. 
To set-up the right leg murine catheter used to measure BP, first, put on sterile gloves. Then get the sterile PE-10 tubing from the autoclaved pouch. Grab the middle of the tubing with pointer finger and thumb leaving about an inch between them. Stretch this section of the tubing just a bit to make it thinner in diameter to help with catheter insertion. After stretching the tubing cut it in half using sterile scissors. The 5-inch tubing should now be two pieces approximately 2 &frac12;-inches in length. Make sure to bevel the stretched end. 
*Do NOT angle the beveled edge too much as this could increase the chances of exiting the lumen by poking through the underside of the vessel wall.
Insert a 30G needle into the blunt unstretched end of the tubing. Get a sterile 1cc syringe and a 3-way stopcock. Use an alcohol wipe to sterilize the top of the 10cc sterile vial containing the heparinized saline solution (0.1ml Heparin/9.9ml Saline). Fill the syringe with 0.6-0.7cc of the heparin solution. Attach the 30G needle and catheter to the end of the 3-way that is directly across from the male end. Fill the stopcock, 30G needle, and PE-10 tubing with the heparin solution. Make sure to get ALL the AIR BUBBLES OUT of the system. The most effective way to remove all the air bubbles is to use gravity. Point the needle toward the ground while letting the tubing dangle to the benchtop. Give the needle hub a flick of the fingers and the bubbles will float to the top of the heparin solution. Remove the 30G needle from the 3-way and remove the bubbles. Withdraw fluid in the 3-way back into the 1cc syringe to remove any bubbles that are trapped in the 3-way and reattach the tubing to the 3-way. Approximately 1cc of the mixture should remain in the syringe. The mouse will receive about 0.05cc of this mixture (as a result of flushing the catheter to maintain patency upon insertion) equaling about 1U heparin/mouse. Place this completed catheter onto the sterile field dressing with the surgical instruments.
To set-up the left leg murine catheter follow the same procedure as described above with the exception of the 3-way stopcock. The left leg is used to draw the blood and the 3-way stopcock is not required. Fill another sterile 1cc syringe with 0.15-0.2cc of the heparinized saline mixture. Hook up the 30G needle and PE-10 tubing directly to the 1cc syringe. Fill this left leg catheter system with the solution. Remove the bubbles from this system, too. Place the completed catheter onto the sterile field dressing with the sterile instruments.
2. Transducer Set-up.
Hook up a sterile transducer to the digi-med BPA 400 analyzer according to the micro-med specifications. Attach a 3-way stopcock to both ends of the transducer. Fill a 10cc syringe with Lactated Ringer solution (LR) and attach it to the 3-way so the transducer will lie flat on the benchtop. Insert a 23G needle into both ends of the piece of precut presterilized 18-inches PE-50 tubing. Attach one end of the PE-50 tubing to the 3-way with the 10cc syringe attached. Fill the 3-way and PE-50 set-up with LR. Make sure to get ALL the AIR BUBBLES OUT of the system as described in the previous section. Reattach the 3-way to the transducer and fill the transducer and 2nd 3-way with LR. Finally, attach the metal male-male leur-lock stopcock to the 23G needle of the PE-50 tubing for attachment to the right leg murine catheter. 
*It is critical that fluid remain in the transducer when in operation. 
*Follow calibration and zero procedures according to Micro-med protocol.
3. Surgical and Experimental Procedures:
1. Surgical Procedures.
Begin by administering an intraperitoneal injection of Pentobarbitol Sodium (Nembutol) (70mg/kg @ 1:10 dilution). This procedure is accomplished by, first, picking the mouse up from its cage using the base (most proximal end) of its tail. Next, place the animal on top of the cage while still holding its tail. Grab the neck scruff of the mouse with the thumb and middle finger on either side of the mouse just behind the forepaws. The index finger is used to pull the skin on the head/neck region back toward the scruff to immobilize the head. The mouse&#x2019;s tail is then wrapped and held between the little finger and the ring finger while the ring finger is pressed into the lumbar region of the mouse&#x2019;s spine. The mouse should be asleep within 5 minutes. After the animal is anesthetized place them onto the metal surgical plate in the supine position. The loose loop tape technique is used to immobilize the animals by taping their extremities. The loose loop technique entails simply cutting thin strips of tape and wrapping the tape loosely around each of the fore limbs inferior to the paw and around each of the hind limbs inferior to the paw. The tape is then stuck back to itself and the left over tape is attached to the board. This allows the extremities of the mice to assume a more natural anatomical position. The animal&#x2019;s abdominal and inguinal areas are then shaved using Oster A5 clippers size 40 blade. A 4x4 gauze is doused with betadine and the surgical area is then wiped for sterility. 
After immobilization and sterilization, a nose cone with 1cc of isoflurane is placed over the mouse&#x2019;s nose for a few seconds before making the initial incision. The nose cone consists of a 50cc conical tube filled with gauze. Half of the bottom of the tube is cut out creating a space for the mouse&#x2019;s nose to rest inside without contact. A cap (bottom of a tissue storage container) is placed onto the end of the 50cc conical to ensure the isoflurane vapors don&#x2019;t escape. Once the animal&#x2019;s respirations begin to slow, a small 4-5mm incision is made in the skin parallel to the left internal oblique muscle of the abdomen and the left transverse abdominus muscle. Dissection of the femoral vein and artery follows. Make sure not to damage the surrounding muscles or touch nerves. To begin this dissection, separate the adipose tissue from the oblique and transverse abdominal muscles by grabbing the adipose tissue with the dumonts at the abdominal connection. Pull this tissue away from the muscle wall. Then, blunt dissect along the abdominal muscles teasing away fascia and adipose tissue using the other pair of dumonts. Just beneath this adipose tissue lie the femoral vein and artery along with the femoral nerve.
*Be sure not to damage the vastus intermedius, medialis, and lateralis muscle of the quadriceps femoris or the rectus femoris. There is really no need to grab or even touch these muscles. 
*DO NOT TOUCH the FEMORAL NERVE
Dissect away the nerve by grabbing the adipose tissue that lies next to it. Pull this tissue laterally from the vein and artery and the nerve will follow as it is embedded in this tissue. As the nerve is pulled laterally, blunt dissect the fascia by placing the other dumonts, point down, against the artery and opening and closing them. The vessels are very superficial so be sure not to dig into the underlying muscles. After the nerve is separated, use the dumonts to separate the fascia holding the vessels to the muscles. Keep the dumonts closed and slip the dumonts dorsal to the vessels. As the tip of the dumonts appears on the other side of the vein, open them to blunt dissect the fascia. Keep the dumonts dorsal to the vessel and grab the first suture. Put the suture into the dumonts and pull the suture back through the opening made between the vessels and the underlying muscles. Again, there is no need to damage any surrounding muscle.
Put a total of 3, 6-0 sutures around the vein and artery. Suture 1 is the most proximal to the abdominal muscles. Tie a knot but leave it loose and hemostat it off. The concave edge of the hemostat should rest on the animal&#x2019;s abdominal cavity. Suture 2 is the most distal in location. This suture can be tied off immediately ligating the vessels and hemostated, again concave side down. The distal and proximal sutures are used to pull the vessels taut (to prevent blood loss) and lift them a bit to aid in catheter insertion. Suture 3 is a catheter support suture. Place this suture between the distal and proximal sutures. 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. It is very white. Make a small incision on the top of the artery using the microscissors. Make this hole close to the distal suture so there is an ample amount of artery for the initial catheter insertion. Use the dumonts to open the hole by placing one end of the dumonts into the arterial vessel lumen and closing them over the vessel wall. Be sure the middle suture is proximal to the arterial hole so it can be used to hold the catheter in place after initial insertion. 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. Release the proximal hemostat. This release will 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 dumont and use the other to push the catheter into the vessel about 4-5mm. Holding the vessel around the catheter helps to prevent tearing of the artery. The tip of the catheter should rest just beneath the internal oblique and transverse abdominus muscles. For blood clot prevention in the arterial line, withdraw blood into the catheter and push it back into the mouse several times to infuse heparinized fluid. Repeat this procedure for the other leg for bilateral femoral artery cannulation. Hook the animal up to the physiological parameter monitor, i.e. the BPA-400 analyzer and flush the arterial lines. Put 1 or 2 drops of sterile saline into the surgical opening to keep the surrounding tissue moist. Be sure to keep this area saturated throughout the procedure. Place a sterile field dressing over the animal throughout the procedure to help maintain sterility. 
Place surgical instruments into 70% alcohol and wipe them with sterile gauze. Put them into the hot bead sterilizer for &tilde;20 seconds for sterilization between animals. Remove the surgical instruments and spray them with 70% alcohol to help them cool off. Place them onto the sterile pad. Make sure there is no alcohol left on the instruments that will drip back into the next animals.
2. Hemorrhagic Shock.
Over a 15min. timeframe, approximately 1/2 of the mouse&#x2019;s blood volume is withdrawn achieving a mean arterial pressure of 28-32mm Hg.
*For a 25-27g mouse the initial volume of blood withdrawn to achieve the desired pressure is approximately 0.6cc
This procedure is a fixed pressure method as opposed to a fixed volume. These procedures can, however be followed for both fixed pressure and fixed volume hemorrhage. While being consistently monitored, the animal will remain in hemorrhagic shock for 1.5-3hours. As the animal attempts to compensate and the mean arterial pressure begins to rise again slightly (visible via the BP/HR analyzer) withdraw more blood to achieve the desired pressure. Although supplements (0.05cc Nembutol IP) are rarely ever needed during the HS procedure, animal respiration, whisker movement, reflex tests, and the digital BP/HR reading will help determine when an animal needs a supplement of anesthesia. Animals are kept under lamp and on a circulating heating pad to help maintain a temperature of 36-37&deg;C through the Hemorrhagic Shock procedure. Temperature is checked via a rectal probe. 
3. Resuscitation.
After the shock time elapses, the animal is resuscitated (R) using Lactated Ringers (LR) solution at a fixed volume of 3x each animal's shed blood volume. The LR volume is administered via a syringe pump set to dispense, at a constant rate, the volume over a 15min. interval. Remove the catheter and ligate the vessels using the 3 sutures. Pull the catheter just past the proximal suture and tie this suture completely off. That will prevent any blood loss. Collateral flow prevents the hind limbs from becoming ischemic. 
4. Post Rescitation.
After R both hind limb openings are sewn up using sterile 4-0 PDSII suture. The loose loop tape is removed and the animals are placed into a clean cage which is kept on a circulating heating pad for several hours post recovery. Analgesic must be administered as the animals begin to waken from the anesthesia to properly manage pain. Buprenorphine (0.1mg/kg) is injected subcutaneously as the animals begin physical activity, but not before, so as not to compromise respiratory function. Use of aggressive post operative monitoring as warranted by IACUC approved protocol, individual animal behaviors, and current medical conditions is recommended. 
4.	Secrets for Success:
<ol>
<li>When dissecting the vessels, Be sure NOT to TOUCH the Femoral Nerve </li>
<li>Be sure not to damage the rectus femoris, rectus lateralis, and vastus medialis muscles. There is really no need to grab or even touch these muscles. </li>
<li>Do Not make the angle of the bevel of the catheter too sharp as this will lead to the catheter exiting the lumen and uncontrolled bleeding </li>
<li>Make sure to take the extra effort to remain as sterile as possible. </li>
<li>Make sure there are NO air bubbles in any of the catheter or transducer systems.</li>
<li>Make sure to have LR fluid in transducer while it is in operation. A dry run could damage your BP system.</li>
<li>Make the arterial hole small.</li>
<li>Make the arterial hole close to the distal suture.</li>
<li>Push catheter while simultaneously pulling vessel over it for catheter insertion in order to avoid ripping the artery.</li>
<li>Be sure to keep the surgical openings in the legs saturated with sterile saline to prevent uncontrolled tissue damage or fluid loss.</li>
<li>Patience and finesse are critical.</li>
<li>Pay close attention to the animal&#x2019;s physiology throughout the experiment.</li>
</ol>
5.	Post Operative Concerns:
<ol>
<li>Check for ischemia in leg. Although collateral flow should prevent this.</li>
<li>Animal could have trouble using hind limbs as a result of surgical manipulation and associated inflammation. Manage pain appropriately.</li> 
<li>Touching and subsequently damaging the nerve could lead to inability of the animal to mobilize.</li>
</ol>

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+silico+and+in+vivo+approach+to+elucidate+the+inflammatory+complexity+of+CD14-deficient+mice.">In silico and in vivo approach to elucidate the inflammatory complexity of CD14-deficient mice.</a> Mol Med. 12, 88-96 (2006).</li><li>Kobbe, P., Stoffels, B., Schmidt, J., Tsukamoto, T., Gutkin, D. W., Bauer, A. J., 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=IL-10+deficiency+augments+acute+lung+but+not+liver+injury+in+hemorrhagic+shock.">IL-10 deficiency augments acute lung but not liver injury in hemorrhagic shock.</a> Cytokine. 45, 26-31 (2009).</li><li>Prince, J. M., Ming, M. J., Levy, R. M., Liu, S., Pinsky, D. J., Vodovotz, Y., 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=Early+Growth+Response+1+mediates+the+systemic+and+hepatic+inflammatory+response+initiated+by+hemorrhagic+shock.">Early Growth Response 1 mediates the systemic and hepatic inflammatory response initiated by hemorrhagic shock.</a> Shock. 27, 157-164 (2007).</li><li>Matsutani, T., Samy, A. n. a. n. t. h. a., Kang, T. S., Bland, S. -. C., Chaudry, K. I., H, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+genetic+background+influences+severity+of+immune+responses+following+trauma-hemorrhage.">Mouse genetic background influences severity of immune responses following trauma-hemorrhage.</a> Cytokine. 30, 168-176 (2005).</li><li>Rushing, G. D., Britt, L. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reperfusion+injury+after+hemorrhage:+a+collective+review.">Reperfusion injury after hemorrhage: a collective review.</a> Ann Surg. 247, 929-937 (2008).</li><li>Makley, A. T., Goodman, M. D., Friend, L. W., Bailey, S. R., Lentsch, A. B., Pritts, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Damage+control+resuscitation+with+fresh+blood+products+attenuates+the+systemic+inflammatory+response+following+hemorrhagic+shock.">Damage control resuscitation with fresh blood products attenuates the systemic inflammatory response following hemorrhagic shock.</a> J Surg Res. 158, 423-423 (2010).</li><li>Stansbury, L. G., Dutton, R. P., Stein, D. M., Bochicchio, G. V., Scalea, T. M., Hess, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Controversy+in+trauma+resuscitation:+do+ratios+of+plasma+to+red+blood+cells+matter?.">Controversy in trauma resuscitation: do ratios of plasma to red blood cells matter?.</a> Transfus Med Rev. 23, 255-265 (2009).</li><li>Gao, J., Zhao, W. X., Xue, F. S., Zhou, L. J., Yu, Y. H., Zhou, H. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+different+resuscitation+fluids+on+acute+lung+injury+in+a+rat+model+of+uncontrolled+hemorrhagic+shock+and+infection.">Effects of different resuscitation fluids on acute lung injury in a rat model of uncontrolled hemorrhagic shock and infection.</a> J Trauma. 6, 1213-1219 (2009).</li><li>Maier, S., Holz-Holzl, C., Pajk, W., Ulmer, H., Hengl, C., Dunser, M., Haas, T., Velik-Salchner, C., Fries, D., Greiner, A., Hasibeder, W., Knotzer, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microcirculatory+parameters+after+isotonic+and+hypertonic+colloidal+fluid+resuscitation+in+acute+hemorrhagic+shock.">Microcirculatory parameters after isotonic and hypertonic colloidal fluid resuscitation in acute hemorrhagic shock.</a> J Trauma. 66, 33-45 (2009).</li><li>Stahel, P. F., Smith, W. R., Moore, E. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Current+trends+in+resuscitation+strategy+for+the+multiply+injured+patient.">Current trends in resuscitation strategy for the multiply injured patient.</a> Injury. 40, 27-35 (2009).</li><li>Dar, D. E., Soustiel, J. F., Zaaroor, M., Em, B., L, A., Shapira, Y., Semenikhina, L., Solopov, A., Krausz, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Moderate+lactated+ringer's+resuscitation+yields+best+neurological+outcome+in+controlled+hemorrhagic+shock+combined+with+brain+injury+in+rats.">Moderate lactated ringer's resuscitation yields best neurological outcome in controlled hemorrhagic shock combined with brain injury in rats.</a> Shock. , (2009).</li><li>Moon, P. F., Hollyfield-Gilbert, M. A., Myers, T. L., Uchida, T., Kramer, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluid+compartment+in+hemorrhaged+rats+after+hyperosmotic+crystalloid+and+hyperoncotic+crystalloid+resuscitation.">Fluid compartment in hemorrhaged rats after hyperosmotic crystalloid and hyperoncotic crystalloid resuscitation.</a> Am J Physiol. 270, F1-F8 (1996).</li><li>Perel, P., Roberts, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Colloids+vs+crystalloids+for+fluid+resuscitation+in+critically+ill+patients.">Colloids vs crystalloids for fluid resuscitation in critically ill patients.</a> Cochrane Database System Rev. 4, CD000567-CD000567 (2007).</li></ol>

Experiments on animals were overseen and performed in accordance with the guidelines and regulations set forth by the Institutional Animal Care and Use Committee and the Research Conduct and Compliance Office of the University of Pittsburgh, a fully accredited AALAS/AALAC institution. Animal sources include Jackson Laboratories and Charles Rivers Laboratories. All animals undergo extensive health assurance through each vendor as well as the University of Pittsburgh&#x2019;s internal animal health monitoring programs. This research is conducted in accordance with the US government principals for Use of vertebrate animals. The program is registered with the USDA, and has a letter of assurance with the Public Health Service Office of Laboratory Animal Welfare.

<table border="1">
	<tbody>
 	<tr>
 		<th>Instrument</th>
 	<th>Manufacturer</th>
 	<th>Catalogue Number</th>
 </tr>
 <tr>
 	<td> Dumonts(SS/45&deg; angle 0.2x0.12mm-tip, 13.5cm length)</td>
 <td>Fine Science Tools</td>
 <td>11203-25</td>
 
 </tr>
 <tr>
 	<td>Surgical scissors (straight-12cm)</td>
 <td>Fine Science Tools</td>
 <td>14068-12</td>
 </tr>
 <tr>
 	<td>Hemostats (curved-12.5cm)</td>
 <td>Fine Science Tools</td>
 <td>13009-12</td>
 </tr>
 <tr>
 	<td>Microscissors (spring scissors- straight-8cm)</td>
 <td>Fine Science Tools</td>
 <td>15000-00</td>
 </tr>

 <tr>
 	<td>Forceps (0.8mm-tip, curved-10cm)</td>
 <td>Fine Science Tools</td>
 <td>11050-10</td>
 </tr>

 <tr>
 	<td>suture reel 6-0</td>
 <td>Fine Science Tools</td>
 <td>18020-60</td>
 </tr>

 <tr>
 	<td>suture 4-0 PDSII </td>
 <td>Penn Vet</td>
 <td>ETHZ304H</td>
 </tr>

 <tr>
 	<td>gauze 4x4</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>cotton-tip applicator</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>30G needle</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>23G needle</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>3cc syringe</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>5cc syringe</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>10cc syringe</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</td>
 </tr>

 <tr>
 	<td>50cc conical tube</td>
 <td>can be purchased through any global vendor</td>
 <td>&nbsp;</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"</td>
 <td>Corporate Express</td>
 <td>MMM26001</td>
 </tr>
		<tr>
 	<td>
straight side wide mouth jars (used as cap for nose cone)
</td>
 <td>
VWR
</td>
 <td>
159000-058
</td>
 </tr>
 <tr>
 	<td>stainless steel tray 8" x 11"</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"x8"</td>
 <td>VWR</td>
 <td>24008</td>
 </tr>
		<tr>
 	<td>sterilization pouch 5"x10"</td>
 <td>VWR</td>
 <td>24010</td>
 </tr>
 
		<tr>
 	<td>Wild M650 microscope w/ boom stand</td>
 <td>Leica</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Leica IC D digital camera/live image </td>
 <td>Leica</td>
 <td>&nbsp;</td>
 </tr>
		<tr>
 	<td>Digi-Med BPA-400 analyzer &amp; systems integrator</td>
 <td>Micro-Med</td>
 <td>SYS-400</td>
 </tr>
 <tr>
 	<td>TXD-310 (Digi-Med Transducer) </td>
 <td>Micro-Med</td>
 <td>TXD-300</td>
 </tr>
		<tr>
 	<td>Computer</td>
 <td>Dell</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td> Hot bead instrument sterilizer</td>
 <td>VWR</td>
 <td>18000-45</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>

<table border="1">
	<tbody>
 	<tr>
 	<th>Reagent</th>
 <th>Concentration</th>
 <th>Manufacturer</th>
 <th>Catalogue Number</th>
 </tr>
 <tr>
 	<td>Pentobarbital Sodium (Nembutol Sodium Solution)</td>
 <td>70mg/kg</td>
 <td>Ovation </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Buprenorphine HCl</td>
 <td>0.1mg/kg</td>
 <td>Bedford Laboratories</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Lactated Ringers Injection 250cc IV bag</td>
 <td>Baxter </td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Aerrane (Isoflurane)</td>
 <td>99.9%</td>
 <td>NLS Animal Health </td>
 <td>105996</td>
 </tr>
 <tr>
 	<td>Heparin Sodium </td>
 <td>1000U</td>
 <td>Abraxis</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Bacteriostatic Sodium Chloride</td>
 <td>0.9%</td>
 <td>Hospira</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Ethyl Alcohol</td>
 <td>70%</td>
 <td>Pharmaco-AAPER</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Triadine Povidone Iodine (Betadine)</td>
 <td>&nbsp;</td>
 <td>Triad disposables</td>
 <td>&nbsp;</td>
 </tr>
 

 
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Fixed Volume or Fixed Pressure: A Murine Model of Hemorrhagic Shock at JoVE.com

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

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, ...

fixed-volume-or-fixed-pressure-a-murine-model-of-hemorrhagic-shock

Research

JoVE Journal

Medicine

Measurement of the Pressure-volume Curve in Mouse Lungs

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential phenotypic changes are best assessed by measurement of the changes in lung elasticity. To best understand specific mechanisms underlying such pathologies in mice, it is essential to make functional measurements that can reflect the developing pathology. Although there are many ways to measure elasticity, the classical method is that of the total lung pressure-volume (PV) curve done over the whole range of lung volumes. This measurement has been made on adult lungs from nearly all mammalian species dating back almost 100 years, and such PV curves also played a major role in the discovery and understanding of the function of pulmonary surfactant in fetal lung development. Unfortunately, such total PV curves have not been widely reported in the mouse, despite the fact that they can provide useful information on the macroscopic effects of structural changes in the lung. Although partial PV curves measuring just the changes in lung volume are sometimes reported, without a measure of absolute volume, the nonlinear nature of the total PV curve makes these partial ones very difficult to interpret. In the present study, we describe a standardized way to measure the total PV curve. We have then tested the ability of these curves to detect changes in mouse lung structure in two common lung pathologies, emphysema and fibrosis. Results showed significant changes in several variables consistent with expected structural changes with these pathologies. This measurement of the lung PV curve in mice thus provides a straightforward means to monitor the progression of the pathophysiologic changes over time and the potential effect of therapeutic procedures.

Here we present a protocol to simply and reliably measure the lung pressure-volume curve in mice, showing that it is sufficiently sensitive to detect phenotypic parenchymal changes in two common lung pathologies, pulmonary fibrosis and emphysema. This metric provides a means to quantify the lung&#8217;s structural changes with developing pathology.

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential ...

Scleral Cross-linking Using Riboflavin and Ultraviolet-A Radiation for Prevention of Axial Myopia in a Rabbit Model

Myopic individuals, especially those with severe myopia, are at higher-than-normal risk of cataract, glaucoma, retinal detachment and chorioretinal abnormalities. In addition, pathological myopia is a common irreversible cause of visual impairment and blindness1-3. Our study demonstrates the effect of scleral crosslinking using riboflavin and ultraviolet-A radiation on the development of axial myopia in a rabbit model. The axial length of the eyeball was measured by A-scan ultrasound in New Zealand white rabbits aged 13 days (male and female). The eye then underwent 360&#176; conjunctival peritomy with scleral crosslinking, followed by tarsorrhaphy. Axial elongation was induced in 13 day-old New Zealand rabbits by suturing their right eye eyelids (tarsorrhaphy). The eyes were divided into quadrants, and every quadrant had two scleral irradiation zones, each with an area of 0.2 cm&#178; and a radius of 4 mm. Crosslinking was performed by dropping 0.1% dextran-free riboflavin-5-phosphate onto the irradiation zones 20 sec before ultraviolet-A irradiation and every 20 sec during the 200 sec irradiation time. UVA radiation (370 nm) was applied perpendicular to the sclera at 57 mW/cm&#178; (total UVA light dose, 57 J/cm&#178;). Tarsorrhaphies were removed on day 55, followed by repeated axial length measurements. This study demonstrates that scleral crosslinking with riboflavin and ultraviolet-A radiation effectively prevents occlusion-induced axial elongation in a rabbit model.

We demonstrate the effect of scleral crosslinking with riboflavin and UVA on an axial elongation rabbit eye. Axial elongation was induced in 13 day-old New Zealand rabbits (male and female) by suturing their right eye eyelids (tarsorrhaphy).

Myopic individuals, especially those with severe myopia, are at higher-than-normal risk of cataract, glaucoma, retinal detachment and chorioretinal ...

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.

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 ...

Surgical Training for the Implantation of Neocortical Microelectrode Arrays Using a Formaldehyde-fixed Human Cadaver Model

This protocol describes a procedure to assist surgeons in training for the implantation of microelectrode arrays into the neocortex of the human brain. Recent technological progress has enabled the fabrication of microelectrode arrays that allow recording the activity of multiple individual neurons in the neocortex of the human brain. These arrays have the potential to bring unique insight onto the neuronal correlates of cerebral function in health and disease. Furthermore, the identification and decoding of volitional neuronal activity opens the possibility to establish brain-computer interfaces, and thus might help restore lost neurological functions. The implantation of neocortical microelectrode arrays is an invasive procedure requiring a supra-centimetric craniotomy and the exposure of the cortical surface; thus, the procedure must be performed by an adequately trained neurosurgeon. In order to provide an opportunity for surgical training, we designed a procedure based on a human cadaver model. The use of a formaldehyde-fixed human cadaver bypasses the practical, ethical and financial difficulties of surgical practice on animals (especially non-human primates) while preserving the macroscopic structure of the head, skull, meninges and cerebral surface and allowing realistic, operating room-like positioning and instrumentation. Furthermore, the use of a human cadaver is closer to clinical daily practice than any non-human model. The major drawbacks of the cadaveric simulation are the absence of cerebral pulsation and of blood and cerebrospinal fluid circulation. We suggest that a formaldehyde-fixed human cadaver model is an adequate, practical and cost-effective approach to ensure proper surgical training before implanting microelectrode arrays in the living human neocortex.

We designed a procedure in which a formaldehyde-fixed human cadaver is used to assist neurosurgeons in training for the implantation of microelectrode arrays into the neocortex of the human brain.

This protocol describes a procedure to assist surgeons in training for the implantation of microelectrode arrays into the neocortex of the human brain. ...

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 ...

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 ...

Closed Chest Biventricular Pressure-Volume Loop Recordings with Admittance Catheters in a Porcine Model

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular evaluation is often performed in preclinical research. However, the right and left ventricles exert functional interdependence due to their parallel and serial connections, encouraging simultaneous evaluation of both ventricles. Furthermore, various pharmacological interventions may affect the ventricles and their preloads and afterloads differently. 
We describe our closed chest approach to admittance-based bi-ventricular PV loop recordings in a porcine model of acute right ventricular (RV) overload. We utilize minimally invasive techniques with all vascular accesses guided by ultrasound. PV catheters are positioned, under fluoroscopic guidance, to avoid thoracotomy in animals, as the closed chest approach maintains the relevant cardiopulmonary physiology. The admittance technology provides real-time PV loop recordings without the need for post-hoc processing. Furthermore, we explain some essential troubleshooting steps during critical timepoints of the presented procedure. 
The presented protocol is a reproducible and physiologically relevant approach to obtain a bi-ventricular cardiac PV loop recording in a large animal model. This can be applied to a large variety of cardiovascular animal research.

Here we present a closed chest approach to admittance-based bi-ventricular pressure-volume loop recordings in pigs with acute right ventricular dysfunction.

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular ...

Complete and Partial Resuscitative Endovascular Balloon Occlusion of the Aorta for Hemorrhagic Shock

Resuscitative endovascular balloon occlusion of the aorta (REBOA) devices grew out of a military-civilian partnership to develop new capabilities for hemorrhage control. With the advent of purpose-built devices, REBOA has become increasingly common in civilian trauma and acute care settings. Currently available REBOA catheters were designed as complete aortic occlusion devices. However, the therapeutic window for complete aortic occlusion is time-limited due to ischemia-reperfusion injury. The partial procedure allows blood flow past the level of occlusion while maintaining targeted proximal pressure, which has been shown to reduce distal ischemia and adjunctive resuscitation requirements in preclinical studies with prolonged occlusion times as compared to traditional complete occlusion.
pREBOA-PRO is the first catheter designed to enable partial and complete aortic occlusion and is currently in limited market release at seven Level I trauma centers in North America. This paper will focus on procedural considerations for REBOA, including patient selection criteria and a comparison of complete and partial aortic occlusion in a simulator, along with highlighting critical steps to improve clinical outcomes. Additionally, this paper reviews a contrast-enhanced CT scan from a trauma patient that shows distal perfusion after 2 h of partial aortic occlusion using this newly designed catheter and discusses representative results from the limited market release to highlight the profound effect of technological innovation on outcomes in vascular emergencies.

A commercial catheter was designed to facilitate true partial resuscitative endovascular balloon occlusion of the aorta (REBOA) and address the complications associated with complete aortic occlusion. Initial clinical reports indicate that partial REBOA improves transition to reperfusion, reduction of distal ischemia, and extension of safe occlusion time compared to complete occlusion.

Resuscitative endovascular balloon occlusion of the aorta (REBOA) devices grew out of a military-civilian partnership to develop new capabilities for ...

Description of a Swine Infant Model of Volume-Controlled Hemorrhagic Shock

Hemorrhagic shock is a leading cause of morbidity and mortality in pediatric patients. Interpretation of the clinical indicators validated in adults to guide resuscitation and comparison between different therapies is difficult in children due to the inherent heterogeneity of this population. As a result, compared to adults, appropriate management of pediatric hemorrhagic shock is still not well established. In addition, the scarcity of pediatric patients with hemorrhagic shock precludes the development of clinically relevant studies. For this reason, an experimental pediatric animal model is necessary to study the effects of hemorrhage in children as well as their response to different therapies. We present an infant animal model of volume-controlled hemorrhagic shock in anesthetized young pigs. Hemorrhage is induced by withdrawing a previously calculated blood volume, and the pig is subsequently monitored and resuscitated with different therapies. Here, we describe a precise and highly reproducible model of hemorrhagic shock in immature swine. The model yields hemodynamic data that characterizes compensatory mechanisms that are activated in response to severe hemorrhage.

This article aims to provide researchers with a detailed and accessible guide to set up an infant porcine model of hemorrhagic shock.

Hemorrhagic shock is a leading cause of morbidity and mortality in pediatric patients. Interpretation of the clinical indicators validated in adults to ...

Dosage-Dependent Induction of Endotoxemia in Pigs via LPS Infusion

Lipopolysaccharide Infusion as a Porcine Endotoxemic Shock Model

Sepsis and septic shock are frequently encountered in patients treated in intensive care units (ICUs) and are among the leading causes of death in these patients. It is caused by a dysregulated immune response to an infection. Even with optimized treatment, mortality rates remain high, which makes further insights into the pathophysiology and new treatment options necessary. Lipopolysaccharide (LPS) is a component of the cell membrane of gram-negative bacteria, which are often responsible for infections causing sepsis and septic shock. 
The severity and high mortality of sepsis and septic shock make standardized experimental studies in humans impossible. Thus, an animal model is needed for further studies. The pig is especially well suited for this purpose as it closely resembles humans in anatomy, physiology, and size. 
This protocol provides an experimental model for endotoxemic shock in pigs by LPS infusion. We were able to reliably induce changes frequently observed in septic shock patients, including hemodynamic instability, respiratory failure, and acidosis. This will allow researchers to gain valuable insight into this highly relevant condition and evaluate new therapeutic approaches in an experimental setting.

Author Spotlight: Induction of Experimental Endotoxemic Shock in Pigs for Studying Hemodynamic and Respiratory Failure

We provide a protocol for an experimental endotoxemic shock model in pigs by infusion of lipopolysaccharide.

Sepsis and septic shock are frequently encountered in patients treated in intensive care units (ICUs) and are among the leading causes of death in these ...