Name: noninvasive and invasive renal hypoxia monitoring in a porcine model of hemorrhagic shock
Uploaded: 2022-10-28T14:20:00.000+00:00
Duration: 7 min 48 s
Description: Watch this Scientific Journal Video about Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock at JoVE.com

The work in this grant is funded by the University of Utah Clinical and Translational Science Institute through the Translational and Clinical Studies Pilot Program and the Department of Defense office of&#160;the Congressionally Directed Medical Research Programs (PR192745).

The Institutional Animal Care and Use Committee of the University of Utah approved all experimental protocols described here. Prior to the experiment, a total of 12 castrated male or non-pregnant female Yorkshire swine weighing 50-75 kg and between 6-8 months old were acclimated in their enclosures for at least 7 days. During this period, all care is directed by a veterinarian and in accordance with the Guide for the Care and Use of Laboratory Animals and the Animal Welfare Act Regulations and Standards. The animals are fasted overnight prior to induction of anesthesia but are allowed free access to water.
1. Sensor assembly
<ol>
	<li>Cut a 6 cm piece of 3/8 in thermoplastic elastomer (TPE) tubing, 25 mm pieces of 1/8 in and 3/16 in PVC tubing, and 31 mm pieces of 1/8 in and 3/16 in PVC tubing.</li>
	<li>Drill a hole in the top of the non-vented cap to fit the exposed tip of the temperature probe; start with a 3/32 in drill bit, then use a 1/8 in drill bit.</li>
	<li>Use a 5/32 in drill bit to drill out the top part of the T-connector to fit the oxygen sensor.</li>
	<li>Slide the shorter piece of the 1/8 in PVC tubing over the inlet side of the flow sensor. Slide the longer 1/8 in piece of PVC tubing over the outlet side (as designated by the arrow on the flow sensor itself) of the flow sensor. Slide the shorter and longer 3/16 in pieces of PVC tubing over the corresponding lengths of the 1/8 in PVC tubing. Insert the barbed end of the male luer lock connector into the open end of the 1/8 in PVC tubing. 
	NOTE: If necessary, use a heat gun to heat the tubing prior to sliding over barbed fittings. It is also possible to use isopropyl alcohol to lubricate the barbed end to make it easier to slide the tubing over the barbed connector.</li>
	<li>Mix the biocompatible glue.</li>
	<li>Expose the tip of the temperature probe by removing any protective sheathing or tubing. Fill the inside of the tubing of the thermistor with biocompatible glue but do not cover the exposed tip.</li>
	<li>Assemble the parts as shown in Figure 1. Use the glue to secure each luer lock connection, when inserting the thermistor in the non-vented cap, and prior to sliding the 3/8 in TPE tubing over the barbed end.</li>
	<li>Prior to sterilization, ensure the blue cap on the oxygen stick is not twisted too tightly, or it will be difficult to undo after sterilization. 
	NOTE: An image of an assembled device is shown in Figure 1 for reference. For this experiment, the fiber optic cable was connected to an electro-optical module that contains software that is designed to work with the specific oxygen sensors used in the device. Any luminescence quenching-based oxygen sensor and compatible data collection device will work. In addition, a custom module and a printed circuit board were designed to connect the flow sensor and temperature probe. Custom software was used to collect and display data in real-time.</li>
</ol>
2. Experimental procedure
<ol>
	<li>Induction of anesthesia and monitoring.
	<ol>
		<li>Sedate the animal with a combined intramuscular injection of Ketamine (2.2 mg/kg) and Xylazine (2.2 mg/kg) &#160;and Telazol (4.4 mg/kg).</li>
		<li>Depending on the size of the animal, place an appropriately sized (most likely between 7 mm and 8 mm) cuffed endotracheal tube with the assistance of a laryngoscope.</li>
		<li>Apply eye lubricant to both eyes.</li>
		<li>Following induction, mechanically ventilate the animal with the maintenance of anesthesia with 1.5%-3.0% gaseous isoflurane mixed in oxygen. Set the fraction of inspired oxygen between 40%-100%, the positive end-expiratory pressure to 4 cm H2O, the tidal volume to 6-8 mL/kg, and adjust the respiratory rate and tidal volume to maintain end-tidal CO2 of 35-45 mmHg.</li>
		<li>Monitor and confirm the proper depth of anesthesia by assessing jaw tone, palpebral reflex approximately every 15 min, and absence of spontaneous movement throughout the experiment. Additionally, monitor clinical parameters of tissue perfusion (mucous membrane color, capillary refill time, heart rate), pulse oximetry, end-tidal CO2, core body temperature, and electrocardiogram.</li>
		<li>Position the animal in dorsal recumbency on a warming blanket and secure each leg to the table.</li>
		<li>The protocol is a non-survival procedure with euthanasia of the animal at the end of the experiment, as described in section 5.</li>
	</ol>
	</li>
	<li>Prepare the animal for the experiment.
	<ol>
		<li>Prepare all puncture sites (which are listed in steps 2.2.3-2.2.7) by scrubbing the skin with three alternating scrubs of chlorhexidine followed by alcohol. After the third scrub, apply chlorhexidine and allow to dry completely, then drape the surgical site in a sterile fashion.</li>
		<li>Locally infiltrate all puncture and incision sites with 2% lidocaine for local pain relief.</li>
		<li>Using ultrasound guidance and the Seldinger technique, place a 9 Fr catheter in the right external jugular vein for medication infusion and central venous pressure monitoring and a 7 Fr catheter in the right femoral vein for resuscitation.</li>
		<li>Under ultrasound guidance, place a 7 Fr sheath in the right brachial artery.</li>
		<li>Under ultrasound guidance, place a 7 Fr sheath in the right femoral artery.</li>
		<li>Under ultrasound guidance, place a 7 Fr sheath in the left femoral artery.</li>
		<li>Under ultrasound guidance, place a 5 Fr sheath in the right or left carotid artery.</li>
		<li>Monitor the pressure distal to the balloon of the resuscitative endovascular occlusion of the aorta (REBOA) catheter via the left femoral artery sheath.
		<ol>
			<li>Connect a disposable pressure transducer to the arterial catheter that is distal to the REBOA balloon.</li>
		</ol>
		</li>
		<li>Monitor the pressure proximal to the balloon of the REBOA catheter via the carotid artery sheath.
		<ol>
			<li>Connect a disposable pressure transducer to the arterial catheter that is proximal to the REBOA balloon.</li>
		</ol>
		</li>
		<li>Perform a midline laparotomy by making an incision along the midline of the abdomen, starting at the inferior part of the sternum and ending at the pubis.</li>
		<li>With the abdomen open, identify the bladder and perform a cystotomy, or make a small incision, to insert the tip of a 20 Fr urinary catheter in the bladder. Close the cystotomy with the urinary catheter in place using a purse string suture. After the catheter is in place, secure it to the skin with sutures.</li>
		<li>Prior to connecting the outlet of the catheter to the urinary collection bag, insert the cone-shaped end of the noninvasive PuO2 monitor into the outlet of the catheter.</li>
		<li>Place the open tubing at the end of the novel PuO2 monitor over the cone-shaped connector on the tubing that is connected to the urine collection bag.</li>
		<li>Remove the spleen to eliminate hemorrhage-induced auto-transfusion.
		<ol>
			<li>Locate the spleen. Identify the hilum of the spleen or the site where the splenic artery and vein enter the spleen. Clamp and transect each vessel.</li>
			<li>After transection, ligate each vessel using modified Miller knots using 2-0 sutures.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Place the instrument to measure bladder PuO2 and tissue oxygenation.
	<ol>
		<li>Measure PuO2&#160;at the outlet of&#160;the bladder.
		<ol>
			<li>Identify the balloon on the catheter. Just below the balloon make an incision along the long axis of the catheter, ensuring that you do not cut the lumen that connects to the balloon.</li>
			<li>After making the incision, insert a t-connector that contains the sensing material into the incision.</li>
			<li>Use tissue glue to secure the t-connector in place.</li>
			<li>Connect the fiber optic cable from the bladder data collection device to the connector that contains the sensing material.</li>
			<li>Create a new file on the data collection device and note the time difference between the stand-alone collection device and other devices used in the experiment.
			<ol>
				<li>For the data collection device used in this study: push the back arrow to reach the main menu.</li>
				<li>Go to measurement settings and click on Ok. Use the arrows to highlight the measurement browser box and push Ok.</li>
				<li>Push the right arrow to create a new file. Type in the name of the new file and select Done.</li>
				<li>Highlight the new file name and select Ok. Navigate to the measurement screen and click on Ok to start recording.</li>
			</ol>
			</li>
		</ol>
		</li>
		<li>Measure medullary renal tissue oxygenation.
		<ol>
			<li>Identify the location of the kidney internally.</li>
			<li>Move the bowel so that you have a clear line of site and access to the entire kidney.&#160;</li>
			<li>Insert the sensor into 2&#34; 18 gauge catheter. Adjust the luer lock connector on the sensor so that the tip of the sensor is exposed. Remove the catheter and place it over an 18 gauge needle.</li>
			<li>Place the 18 gauge needle and 2 in catheter into the renal medulla under ultrasound guidance.</li>
			<li>Remove the needle, keeping the catheter in place. Thread the tissue sensor through the catheter and use the luer lock to connect the sensor to the catheter.&#160;</li>
			<li>Use tissue glue to secure the catheter in place.&#160;</li>
			<li>Connect the tissue sensor to the data collection box.</li>
			<li>Wait for 10 min before beginning the experimental protocol after preparing the instrumentation and animal. This will be considered a baseline period.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Experimental protocol
	<ol>
		<li>Prior to starting the experimental procedure, ensure the mean arterial pressure (MAP) is &#8805;65 mmHg. If MAP is below the threshold, give up to two 5 mL/kg boluses of isotonic crystalloid solution. If MAP remains below 65 mmHg, infuse norepinephrine (0.02 &#181;g/kg/min) until target MAP is achieved.</li>
		<li>Induce hemorrhagic shock.
		<ol>
			<li>Remove 25% (estimated as 60 mL/kg) of the animal&#39;s estimated blood volume through the right brachial artery sheath over 30 min into gently agitated citrated blood collection bags. Mark the beginning of blood removal as t = 0 min.</li>
			<li>Store the removed blood in a warm water bath at 37 &#176;C.</li>
			<li>Then perform randomization to assign animals to either the REBOA with whole blood or REBOA with crystalloids group (n = 6 for each group).</li>
		</ol>
		</li>
		<li>Place the REBOA catheter.
		<ol>
			<li>Insert a 7 Fr REBOA catheter in the right femoral artery sheath. Place the balloon of the catheter immediately superior to the diaphragm and confirm the location using fluoroscopy.</li>
			<li>At t = 30 min, inflate the REBOA balloon and completely occlude the aorta for 45 min.</li>
		</ol>
		</li>
		<li>Initiate resuscitation and administer critical care.
		<ol>
			<li>At t = 70 min, transfuse each animal with their shed blood over 15 min.</li>
			<li>Infuse intravenous calcium over 10 min to prevent citrate-induced hypocalcemia.</li>
			<li>At t = 75 min, deflate the REBOA balloon over 10 min.</li>
			<li>Until t = 360 min, resuscitate the animal with fluids and norepinephrine to maintain a MAP &#62; 65 mmHg.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>End of experiment and euthanasia
	<ol>
		<li>Collect any remaining blood or urine samples.</li>
		<li>Euthanize the animal by injecting a combination of Pentobarbital Sodium (390 mg) and Phenytoin Sodium (50 mg) (1 mL/10 lbs).</li>
	</ol>
	</li>
</ol>
3. Data processing
<ol>
	<li>Time-sync all data files.
	<ol>
		<li>Based on the times that were noted on each device relative to each other and the start of the experiment, align all data files such that t = 0 indicates the start of the experiment.</li>
	</ol>
	</li>
	<li>Remove any data points associated with error flags from the flow sensor. 
	NOTE: The error types are High Flow Rate and Air-in-Line. The High Flow Rate error indicates the flow rate exceeded the sensor&#39;s output limit. The Air-in-Line error flag is raised when the sensor detects air in the flow channel.</li>
	<li>Discard the data associated with the negative flow.
	<ol>
		<li>Once flow becomes negative, track the volume that flows past the sensor in the backward direction.</li>
		<li>After the flow becomes positive, track the volume and compare it to the volume of negative flow to only include measurements from recently voided urine.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Gomes, E., Antunes, R., Dias, C., Ara&#250;jo, R., Costa-Pereira, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+kidney+injury+in+severe+trauma+assessed+by+RIFLE+criteria:+a+common+feature+without+implications+on+mortality.">Acute kidney injury in severe trauma assessed by RIFLE criteria: a common feature without implications on mortality.</a> Scandinavian Journal of Trauma, Resuscitation and Emergency Medicine. 18, 1 (2010).</li><li>Bihorac, A., 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=Incidence,+clinical+predictors,+genomics,+and+outcome+of+acute+kidney+injury+among+trauma+patients.">Incidence, clinical predictors, genomics, and outcome of acute kidney injury among trauma patients.</a> Annals of Surgery. 252 (1), 158-165 (2010).</li><li>Perkins, Z. B., 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=Trauma+induced+acute+kidney+injury.">Trauma induced acute kidney injury.</a> Plos One. 14 (1), 0211001 (2019).</li><li>Lai, W. H., 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=Post-traumatic+acute+kidney+injury:+a+cross-sectional+study+of+trauma+patients.">Post-traumatic acute kidney injury: a cross-sectional study of trauma patients.</a> Scandinavian Journal of Trauma, Resuscitation and Emergency Medicine. 24 (1), 136 (2016).</li><li>Khwaja, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=KDIGO+clinical+practice+guidelines+for+acute+kidney+injury.">KDIGO clinical practice guidelines for acute kidney injury.</a> Nephron Clinical Practice. 120 (4), 179-184 (2012).</li><li>Saour, M., 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=Assessment+of+modification+of+diet+in+renal+disease+equation+to+predict+reference+serum+creatinine+value+in+severe+trauma+patients:+Lessons+from+an+observational+study+of+775+cases.">Assessment of modification of diet in renal disease equation to predict reference serum creatinine value in severe trauma patients: Lessons from an observational study of 775 cases.</a> Annals of Surgery. 263 (4), 814-820 (2016).</li><li>Ostermann, M., Joannidis, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+kidney+injury+2016:+diagnosis+and+diagnostic+workup.">Acute kidney injury 2016: diagnosis and diagnostic workup.</a> Critical Care. 20 (1), 299 (2016).</li><li>Koeze, J., 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=Incidence,+timing+and+outcome+of+AKI+in+critically+ill+patients+varies+with+the+definition+used+and+the+addition+of+urine+output+criteria.">Incidence, timing and outcome of AKI in critically ill patients varies with the definition used and the addition of urine output criteria.</a> BMC Nephrology. 18 (1), 70 (2017).</li><li>Ralib, A., Pickering, J. W., Shaw, G. M., Endre, Z. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+urine+output+definition+of+acute+kidney+injury+is+too+liberal.">The urine output definition of acute kidney injury is too liberal.</a> Critical Care. 17 (3), 112 (2013).</li><li>Ostermann, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diagnosis+of+acute+kidney+injury:+Kidney+disease+improving+global+outcomes+criteria+and+beyond.">Diagnosis of acute kidney injury: Kidney disease improving global outcomes criteria and beyond.</a> Current Opinion Critical Care. 20 (6), 581-587 (2014).</li><li>Harrois, A., Libert, N., Duranteau, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+kidney+injury+in+trauma+patients.">Acute kidney injury in trauma patients.</a> Current Opinion Critical Care. 23 (6), 447-456 (2017).</li><li>Ow, C. P. C., Ngo, J. P., Ullah, M. M., Hilliard, L. M., Evans, R. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+hypoxia+in+kidney+disease:+Cause+or+consequence.">Renal hypoxia in kidney disease: Cause or consequence.</a> Acta Physiologica. 222 (4), 12999 (2018).</li><li>Leonhardt, K. O., Landes, R. R., McCauley, R. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anatomy+and+physiology+of+intrarenal+oxygen+tension:+Preliminary+study+of+the+effets+of+anesthetics.">Anatomy and physiology of intrarenal oxygen tension: Preliminary study of the effets of anesthetics.</a> Anesthesiology. 26 (5), 648-658 (1965).</li><li>Stafford-Smith, M., Grocott, H. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+medullary+hypoxia+during+experimental+cardiopulmonary+bypass:+a+pilot+study.">Renal medullary hypoxia during experimental cardiopulmonary bypass: a pilot study.</a> Perfusion. 20 (1), 53-58 (2005).</li><li>Kitashiro, 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=Monitoring+urine+oxygen+tension+during+acute+change+in+cardiac+output+in+dogs.">Monitoring urine oxygen tension during acute change in cardiac output in dogs.</a> Journal of Applied Physiology. 79 (1), 202-204 (1995).</li><li>Sgouralis, I., 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=Bladder+urine+oxygen+tension+for+assessing+renal+medullary+oxygenation+in+rabbits:+experimental+and+modeling+studies.">Bladder urine oxygen tension for assessing renal medullary oxygenation in rabbits: experimental and modeling studies.</a> American Journal of Physiology. Regulatory, Integrative, and Comparative Physiology. 311 (3), 532-544 (2016).</li><li>Kainuma, M., Kimura, N., Shimada, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+acute+changes+in+renal+arterial+blood+flow+on+urine+oxygen+tension+in+dogs.">Effect of acute changes in renal arterial blood flow on urine oxygen tension in dogs.</a> Critical Care Medicine. 18 (3), 309-312 (1990).</li><li>Zhu, M. Z. L., 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=Urinary+hypoxia:+an+intraoperative+marker+of+risk+of+cardiac+surgery-associated+acute+kidney+injury.">Urinary hypoxia: an intraoperative marker of risk of cardiac surgery-associated acute kidney injury.</a> Nephrology Dialysis Transplantation. 33 (12), 2191-2201 (2018).</li><li>Silverton, N. A., 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=Noninvasive+urine+oxygen+monitoring+and+the+risk+of+acute+kidney+injury+in+cardiac+surgery.">Noninvasive urine oxygen monitoring and the risk of acute kidney injury in cardiac surgery.</a> Anesthesiology. 135 (3), 406-418 (2021).</li><li>Lankadeva, Y. 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=Intrarenal+and+urinary+oxygenation+during+norepinephrine+resuscitation+in+ovine+septic+acute+kidney+injury.">Intrarenal and urinary oxygenation during norepinephrine resuscitation in ovine septic acute kidney injury.</a> Kidney International. 90 (1), 100-108 (2016).</li><li>Evans, R. G., 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=Renal+hemodynamics+and+oxygenation+during+experimental+cardiopulmonary+bypass+in+sheep+under+total+intravenous+anesthesia.">Renal hemodynamics and oxygenation during experimental cardiopulmonary bypass in sheep under total intravenous anesthesia.</a> American Journal of Physiology. Regulatory, Integrative, and Comparative Physiology. 318 (2), 206-213 (2020).</li><li>Sgouralis, I., Evans, R. G., Layton, A. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+medullary+and+urinary+oxygen+tension+during+cardiopulmonary+bypass+in+the+rat.">Renal medullary and urinary oxygen tension during cardiopulmonary bypass in the rat.</a> Mathematical Medicine and Biology. 34 (3), 313-333 (2017).</li><li>Lankadeva, Y. R., Kosaka, J., Evans, R. G., Bellomo, R., May, C. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Urinary+oxygenation+as+a+surrogate+measure+of+medullary+oxygenation+during+angiotensin+II+therapy+in+septic+acute+kidney+injury.">Urinary oxygenation as a surrogate measure of medullary oxygenation during angiotensin II therapy in septic acute kidney injury.</a> Critical Care Medicine. 46 (1), 41-48 (2018).</li><li>Ngo, J. P., 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=Factors+that+confound+the+prediction+of+renal+medullary+oxygenation+and+risk+of+acute+kidney+injury+from+measurement+of+bladder+urine+oxygen+tension.">Factors that confound the prediction of renal medullary oxygenation and risk of acute kidney injury from measurement of bladder urine oxygen tension.</a> Acta Physiologica. 227 (1), 13294 (2019).</li><li>Spahn, D. 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=The+European+guideline+on+management+of+major+bleeding+and+coagulopathy+following+trauma:+fifth+edition.">The European guideline on management of major bleeding and coagulopathy following trauma: fifth edition.</a> Critical Care. 23 (1), 98 (2019).</li><li>Legrand, M., 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=Fluid+resuscitation+does+not+improve+renal+oxygenation+during+hemorrhagic+shock+in+rats.">Fluid resuscitation does not improve renal oxygenation during hemorrhagic shock in rats.</a> Anesthesiology. 112 (1), 119-127 (2010).</li><li>Badin, J., 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=Relation+between+mean+arterial+pressure+and+renal+function+in+the+early+phase+of+shock:+a+prospective,+explorative+cohort+study.">Relation between mean arterial pressure and renal function in the early phase of shock: a prospective, explorative cohort study.</a> Critical Care. 15 (3), 135 (2011).</li><li>Ribeiro Junio, M. A. 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+complications+associated+with+resuscitative+endovascular+balloon+occlusion+of+the+aorta+(REBOA).">The complications associated with resuscitative endovascular balloon occlusion of the aorta (REBOA).</a> World Journal of Emergency Surgery. 13, 20 (2018).</li></ol>

N. Silverton, K. Kuck, and L. Lofgren are inventors of a patent and patent application surrounding the noninvasive monitor used in this study. This prototype is under development for commercial consideration by N. Silverton and K. Kuck, but as of yet, no commercial activity has occurred. The other authors declare no competing interests. The interpretation and reporting of these data are the responsibility of the authors alone.

AKI is a common complication in patients with trauma, and currently, there is no validated bedside monitor for kidney tissue oxygenation, which could enable earlier AKI detection and guide potential interventions. This manuscript describes the use and instrumentation of a porcine hemorrhagic shock model to establish noninvasive PuO2 as an early indicator of AKI and a novel resuscitation endpoint in trauma settings.
One of the distinct advantages of this porcine model is the ability ...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/64461">Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock</a>. The Protocol&#160;section was updated.
Step 2.3.1&#160;-&#160;2.3.1.2&#160;of the Protocol was updated from:
<ol>
	<li>Measure PuO2&#160;in the bladder.
	<ol>
		<li>Remove all air from the bladder by slowly squeezing the bladder while ensuring urine does not leak out.</li>
		<li>Place the tip of the luminescence quenching-based PuO2&#160;sensor in the bladder&#160;via&#160;a cystotomy, similar to the catheter.</li>
		<li>Connect the fiber optic cable from the bladder sensor to the data collection device.</li>
	</ol>
	</li>
</ol>
to:
<ol>
	<li>&#160;Measure PuO2 at the outlet of the bladder
	<ol>
		<li>Identify the balloon on the catheter. Just below the balloon make an incision along the long axis of the catheter, ensuring that you do not cut the lumen that connects to the balloon.&#160;</li>
		<li>After making the incision, insert a t-connector that contains the sensing material into the incision.&#160;</li>
		<li>Use tissue glue to secure the t-connector in place.&#160;</li>
		<li>Connect the fiber optic cable from the bladder data collection device to the connector that contains the sensing material.&#160;</li>
	</ol>
	</li>
</ol>
Step 2.3.2.2&#160;-&#160;2.3.2.7&#160;of the Protocol was updated from:
<ol start="2">
	<li>Make a flank incision large enough to expose the kidney (approx. 2-3 in) on the side of the pig at approximately the same location where the kidney was identified.</li>
	<li>With the tips of a retractor together, introduce the retractor into the incision and then spread the tips of the retractor to expose the kidney.</li>
	<li>Use a micro-manipulator or similar tool to hold the oxygen probe steady. If possible, attach this tool to the end of an articulating arm.</li>
	<li>Attach the other end of the articulating arm to the surgical table so that the other end that will hold the oxygen probe is near the opened incision. If the tool that is used to hold the oxygen probe is not connected to an articulating arm, position the tool so the oxygen sensor is near the opened incision and is stable.</li>
	<li>Unlock all articulating joints of the arm. Using ultrasound, place the tip of the oxygen probe in the medulla region of the kidney. Lock all articulating joints on the arm.</li>
	<li>After confirming placement of the tip of the sensor in the medulla with ultrasound, use the micromanipulator to retract the needle housing the luminescence-based oxygen sensor. Connect the other end of the sensor to the data collection device connected to the computer running the data collection software. Start recording.</li>
</ol>
to:
<ol start="2">
	<li>Move the bowel so that you have a clear line of site and access to the entire kidney.&#160;</li>
	<li>Insert the sensor into 2&#34; 18 gauge catheter. Adjust the luer lock connector on the sensor so that the tip of the sensor is exposed. Remove the catheter and place it over an 18 gauge needle.</li>
	<li>Place the 18 gauge needle and 2 in catheter into the renal medulla under ultrasound guidance.</li>
	<li>Remove the needle, keeping the catheter in place. Thread the tissue sensor through the catheter and use the luer lock to connect the sensor to the catheter.&#160;</li>
	<li>Use tissue glue to secure the catheter in place.&#160;</li>
	<li>Connect the tissue sensor to the data collection box.</li>
</ol>
The Table of Materials was updated from:
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>1/8&#34; PVC tubing</td>
			<td>Qosina</td>
			<td>SKU: T4307</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>3/16&#34; PVC tubing</td>
			<td>Qosina</td>
			<td>SKU: T4310</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>3/32&#34; (1), 1/8&#34; (1), 5/32&#34; (1) drill bit</td>
			<td>Dewalt</td>
			<td>N/A</td>
			<td>For building noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>3/8&#34; TPE tubing&#160;</td>
			<td>Qosina</td>
			<td>SKU: T2204</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Biocompatible Glue</td>
			<td>Masterbond</td>
			<td>EP30MED</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Bladder oxygen measurement device</td>
			<td>Presens</td>
			<td>Fibox 4</td>
			<td>Stand-alone fiber optic oxygen meter</td>
		</tr>
		<tr>
			<td>Bladder PuO2 sensor</td>
			<td>Presens</td>
			<td>DP-PSt3</td>
			<td>Oxygen dipping probe</td>
		</tr>
		<tr>
			<td>Chlorhexidine 4% scrub</td>
			<td>Vetone</td>
			<td>N/A</td>
			<td>For scrubbing insertion or puncture sites</td>
		</tr>
		<tr>
			<td>Conical connector with female luer lock</td>
			<td>Qosina</td>
			<td>SKU: 51500</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Cuffed endotracheal tube</td>
			<td>Vetone</td>
			<td>600508</td>
			<td>For sedating the subject and providing respiratory support</td>
		</tr>
		<tr>
			<td>Euthanasia solution (pentobarbital sodium|pheyntoin sodium)</td>
			<td>Vetone</td>
			<td>11168</td>
			<td>For euthanasia after completion of experiment</td>
		</tr>
		<tr>
			<td>General purpose temperature probe, 400 series thermistor</td>
			<td>Novamed</td>
			<td>10-1610-040</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Hemmtop Magic Arm 11 inch</td>
			<td>Amazon</td>
			<td>B08JTZRKYN</td>
			<td>Holding invasive oxygen sensor in place</td>
		</tr>
		<tr>
			<td>HotDog veterinary warming system</td>
			<td>HotDog</td>
			<td>V106</td>
			<td>For controlling subject temperature during experiment</td>
		</tr>
		<tr>
			<td>Invasive tissue oxygen measurement device</td>
			<td>Presens</td>
			<td>Oxy-1 ST&#160;</td>
			<td>Compact oxygen transmitter</td>
		</tr>
		<tr>
			<td>Invasive tissue oxygen sensor</td>
			<td>Presens</td>
			<td>PM-PSt7</td>
			<td>Profiling oxygen microsensor</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Vetone</td>
			<td>501017</td>
			<td>To maintain sedation throughout the experiment</td>
		</tr>
		<tr>
			<td>Isotonic crystalloid solution</td>
			<td>HenrySchein</td>
			<td>1537930 or 1534612</td>
			<td>Used during resuscitation in the critical care period</td>
		</tr>
		<tr>
			<td>Liquid flow sensor</td>
			<td>Sensirion</td>
			<td>LD20-2600B</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Male luer lock to barb connector</td>
			<td>Qosina</td>
			<td>SKU: 11549</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Male to male luer connector</td>
			<td>Qosina</td>
			<td>SKU: 20024</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Noninvasive oxygen measurement device</td>
			<td>Presens</td>
			<td>EOM-O2-mini</td>
			<td>Electro optical module transmitter for contactless oxygen measurements</td>
		</tr>
		<tr>
			<td>Non-vented male luer lock cap</td>
			<td>Qosina</td>
			<td>SKU: 65418</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Norepinephrine</td>
			<td>HenrySchein</td>
			<td>AIN00610</td>
			<td>Infusion during resuscitation</td>
		</tr>
		<tr>
			<td>O2 sensor stick</td>
			<td>Presens</td>
			<td>SST-PSt3-YOP</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>PowerLab data acquisition platform</td>
			<td>AD Instruments</td>
			<td>N/A</td>
			<td>For data collection</td>
		</tr>
		<tr>
			<td>REBOA catheter</td>
			<td>Certus Critical Care</td>
			<td>N/A</td>
			<td>Used in experimental protocol</td>
		</tr>
		<tr>
			<td>Super Sheath arterial catheters (5 Fr, 7 Fr, 9 Fr)</td>
			<td>Boston Scientific</td>
			<td>C1894</td>
			<td>For intravascular access</td>
		</tr>
		<tr>
			<td>Suture</td>
			<td>Ethicon</td>
			<td>C013D</td>
			<td>For securing catheter to skin and closing incisions</td>
		</tr>
		<tr>
			<td>T connector, all female luer locks</td>
			<td>Qosina</td>
			<td>SKU: 88214</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
	</tbody>
</table>
to:
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>1/8&#34; PVC tubing</td>
			<td>Qosina</td>
			<td>SKU: T4307</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>3/16&#34; PVC tubing</td>
			<td>Qosina</td>
			<td>SKU: T4310</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>3/8&#34; TPE tubing&#160;</td>
			<td>Qosina</td>
			<td>SKU: T2204</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>3/32&#34; (1), 1/8&#34; (1), 5/32&#34; (1) drill bit</td>
			<td>Dewalt</td>
			<td>N/A</td>
			<td>For building noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Biocompatible Glue</td>
			<td>Masterbond</td>
			<td>EP30MED</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Bladder PuO2 sensor</td>
			<td>Presens</td>
			<td>DP-PSt3</td>
			<td>Oxygen dipping probe</td>
		</tr>
		<tr>
			<td>Bladder oxygen measurement device</td>
			<td>Presens</td>
			<td>Fibox 4</td>
			<td>Stand-alone fiber optic oxygen meter</td>
		</tr>
		<tr>
			<td>Chlorhexidine 4% scrub</td>
			<td>Vetone</td>
			<td>N/A</td>
			<td>For scrubbing insertion or puncture sites</td>
		</tr>
		<tr>
			<td>Conical connector with female luer lock</td>
			<td>Qosina</td>
			<td>SKU: 51500</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Cuffed endotracheal tube</td>
			<td>Vetone</td>
			<td>600508</td>
			<td>For sedating the subject and providing respiratory support</td>
		</tr>
		<tr>
			<td>Euthanasia solution (pentobarbital sodium|pheyntoin sodium)</td>
			<td>Vetone</td>
			<td>11168</td>
			<td>For euthanasia after completion of experiment</td>
		</tr>
		<tr>
			<td>General purpose temperature probe, 400 series thermistor</td>
			<td>Novamed</td>
			<td>10-1610-040</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>HotDog veterinary warming system</td>
			<td>HotDog</td>
			<td>V106</td>
			<td>For controlling subject temperature during experiment</td>
		</tr>
		<tr>
			<td>Invasive tissue oxygen measurement device</td>
			<td>Optronix</td>
			<td>N/A</td>
			<td>OxyLite&#8482; oxygen monitors</td>
		</tr>
		<tr>
			<td>Invasive tissue oxygen sensor</td>
			<td>Optronix</td>
			<td>NX-BF/OT/E</td>
			<td>Oxygen/Temperature bare-fibre sensor</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Vetone</td>
			<td>501017</td>
			<td>To maintain sedation throughout the experiment</td>
		</tr>
		<tr>
			<td>Isotonic crystalloid solution</td>
			<td>HenrySchein</td>
			<td>1537930 or 1534612</td>
			<td>Used during resuscitation in the critical care period</td>
		</tr>
		<tr>
			<td>Liquid flow sensor</td>
			<td>Sensirion</td>
			<td>LD20-2600B</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Male luer lock to barb connector</td>
			<td>Qosina</td>
			<td>SKU: 11549</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Male to male luer connector</td>
			<td>Qosina</td>
			<td>SKU: 20024</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>Norepinephrine</td>
			<td>HenrySchein</td>
			<td>AIN00610</td>
			<td>Infusion during resuscitation</td>
		</tr>
		<tr>
			<td>Noninvasive oxygen measurement device</td>
			<td>Presens</td>
			<td>EOM-O2-mini</td>
			<td>Electro optical module transmitter for contactless oxygen measurements</td>
		</tr>
		<tr>
			<td>Non-vented male luer lock cap&#160;</td>
			<td>Qosina</td>
			<td>SKU: 65418</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>O2 sensor stick</td>
			<td>Presens</td>
			<td>SST-PSt3-YOP</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
		<tr>
			<td>PowerLab data acquisition platform</td>
			<td>AD Instruments</td>
			<td>N/A</td>
			<td>For data collection</td>
		</tr>
		<tr>
			<td>REBOA catheter</td>
			<td>Certus Critical Care</td>
			<td>N/A</td>
			<td>Used in experimental protocol</td>
		</tr>
		<tr>
			<td>Super Sheath arterial catheters (5 Fr, 7 Fr, 9 Fr)</td>
			<td>Boston Scientific</td>
			<td>C1894</td>
			<td>for intravascular access</td>
		</tr>
		<tr>
			<td>Suture</td>
			<td>Ethicon</td>
			<td>C013D</td>
			<td>For securing catheter to skin and closing incisions</td>
		</tr>
		<tr>
			<td>T connector, all female luer locks</td>
			<td>Qosina</td>
			<td>SKU: 88214</td>
			<td>Part of noninvasive PuO2 monitor</td>
		</tr>
	</tbody>
</table>

An erratum was issued for:&#160;<a href="https://www.jove.com/t/64461">Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock</a>. The Protocol and Table of Materials were&#160;updated.

Erratum: Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock

Acute kidney injury (AKI) affects up to 50% of patients with trauma admitted to the intensive care unit1. Patients who develop AKI tend to have longer hospital and intensive care unit lengths of stay and a threefold greater risk of mortality2,3,4. Currently, AKI is most commonly defined by the Kidney Disease Improving Global Outcomes (KDIGO) guidelines, which are based on changes in serum creatinine concentration from baseline or periods of prolonged oliguria5. Baseline creatinine concentration data are unavailable in most patients with trauma, and estimation equations are unreliable and have not been validated in patients with trauma6. In addition, serum creatinine concentration may not change until at least 24 h after the injury, precluding early identification and intervention7. While research suggests that urine output is an earlier indicator of AKI than serum creatinine concentration, the KDIGO criteria require a minimum of 6 h of oliguria, which precludes interventions targeting injury prevention8. The optimal hourly urine output threshold and appropriate duration of oliguria for defining AKI are also debated, which limit its effectiveness as an early marker of the disease9,10. Thus, current diagnostic measures for AKI are not useful in trauma settings, lead to delayed diagnosis of AKI, and do not provide real-time information regarding a patient&#39;s risk status for developing AKI.
While the development of AKI in a trauma setting is complex and likely associated with several causes such as poor renal perfusion due to hypovolemia, reduced renal blood flow due to vasoconstriction, trauma-related inflammation, or ischemia-reperfusion injury, renal hypoxia is a common factor among most forms of AKI11,12. In particular, the medulla region of the kidney is highly susceptible to an imbalance between oxygen demand and supply in the trauma setting due to reduced oxygen delivery and high metabolic activity associated with sodium reabsorption. Thus, if it were possible to measure renal medulla oxygenation, it may be possible to monitor a patient&#39;s risk status for developing AKI. While this is not clinically feasible, urinary partial pressure of oxygen (PuO2) at the outlet of the kidney strongly correlates with medullary tissue oxygenation13,14. Other studies have shown that it is possible to measure bladder PuO2 and that it changes in response to stimuli that alter medullary oxygen and renal pelvis PuO2 levels, such as a decrease in renal blood flow15,16,17. These studies suggest that PuO2 may indicate end-organ perfusion and could be useful for monitoring the impact of interventions in trauma settings on renal function.
To monitor PuO2 noninvasively, a noninvasive PuO2 monitor was developed that can easily connect to the end of a urinary catheter outside the body. The noninvasive PuO2 monitor consists of three main components: a temperature sensor, a luminescence quenching oxygen sensor, and a thermal-based flow sensor. Since each oxygen sensor is optically based and relies on the Stern-Volmer relationship to quantify the relationship between luminescence and oxygen concentration, a temperature sensor is necessary to offset any potential confounding effects of changes in temperature. The flow sensor is important to quantify urine output and to determine the direction and magnitude of urine flow. All three components are connected by a combination of male, female, and t-shaped luer lock connectors and poly-vinyl chloride (PVC) flexible tubing. The end with the conical connector connects to the outlet of the urinary catheter, and the end with tubing over the conical connector connects slides over the connector on the urine collection bag.
Despite measuring distally to the bladder, a recent study showed that low urinary PuO2 during cardiac surgery is associated with an increased risk of developing AKI18,19. Similarly, current animal models have primarily focused on the early detection of AKI during cardiac surgery and sepsis14,20,21,22. Thus, questions remain about the use of this novel device in settings of trauma. The aim of this research is to establish PuO2 as an early marker of AKI and investigate its use as a resuscitative endpoint in patients with trauma. This manuscript describes a porcine model of hemorrhagic shock that includes the placement of the noninvasive PuO2 monitor, a bladder PuO2 sensor, and a tissue oxygen sensor in the renal medulla. Data from the noninvasive monitor will be compared to bladder PuO2 and invasive tissue oxygen measurements. The noninvasive monitor also includes a flow sensor which will be useful for understanding the relationship between urine flow rate and oxygen ingress, which reduces the ability to infer renal medullar tissue oxygenation from noninvasive PuO2 as urine traverses the urinary tract. Additionally, data from the three oxygen sensors will be compared to systemic vital signs, such as mean arterial pressure. The central hypothesis is that noninvasive PuO2 data will strongly correlate with invasive medullary oxygen content and will reflect medullary hypoxia during resuscitation. Noninvasive PuO2 monitoring has the potential to improve trauma-related outcomes by identifying AKI earlier and serving as a novel resuscitative endpoint after hemorrhage that is indicative of end-organ rather than systemic oxygenation.

<table><tbody><tr><td>1/8&quot; PVC tubing</td><td>Qosina</td><td>SKU: T4307</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>3/16&quot; PVC tubing</td><td>Qosina</td><td>SKU: T4310</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>3/8&quot; TPE tubing&amp;nbsp;</td><td>Qosina</td><td>SKU: T2204</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>3/32&quot; (1), 1/8&quot; (1), 5/32&quot; (1) drill bit</td><td>Dewalt</td><td>N/A</td><td>For building noninvasive PuO2 monitor</td></tr><tr><td>Biocompatible Glue</td><td>Masterbond</td><td>EP30MED</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>Bladder PuO&lt;sub&gt;2 &lt;/sub&gt;sensor</td><td>Presens</td><td>DP-PSt3</td><td>Oxygen dipping probe</td></tr><tr><td>Bladder oxygen measurement device</td><td>Presens</td><td>Fibox 4</td><td>Stand-alone fiber optic oxygen meter</td></tr><tr><td>Chlorhexidine 4% scrub</td><td>Vetone</td><td>N/A</td><td>For scrubbing insertion or puncture sites</td></tr><tr><td>Conical connector with female luer lock</td><td>Qosina</td><td>SKU: 51500</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>Cuffed endotracheal tube</td><td>Vetone</td><td>600508</td><td>For sedating the subject and providing respiratory support</td></tr><tr><td>Euthanasia solution (pentobarbital sodium|pheyntoin sodium)</td><td>Vetone</td><td>11168</td><td>For euthanasia after completion of experiment</td></tr><tr><td>General purpose temperature probe, 400 series thermistor</td><td>Novamed</td><td>10-1610-040</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>HotDog veterinary warming system</td><td>HotDog</td><td>V106</td><td>For controlling subject temperature during experiment</td></tr><tr><td>Invasive tissue oxygen measurement device</td><td>Optronix</td><td>N/A</td><td>OxyLite&amp;trade; oxygen monitors</td></tr><tr><td>Invasive tissue oxygen sensor</td><td>Optronix</td><td>NX-BF/OT/E</td><td>Oxygen/Temperature bare-fibre sensor</td></tr><tr><td>Isoflurane</td><td>Vetone</td><td>501017</td><td>To maintain sedation throughout the experiment</td></tr><tr><td>Isotonic crystalloid solution</td><td>HenrySchein</td><td>1537930 or 1534612</td><td>Used during resuscitation in the critical care period</td></tr><tr><td>Liquid flow sensor</td><td>Sensirion</td><td>LD20-2600B</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>Male luer lock to barb connector</td><td>Qosina</td><td>SKU: 11549</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>Male to male luer connector</td><td>Qosina</td><td>SKU: 20024</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>Norepinephrine</td><td>HenrySchein</td><td>AIN00610</td><td>Infusion during resuscitation</td></tr><tr><td>Noninvasive oxygen measurement device</td><td>Presens</td><td>EOM-O2-mini</td><td>Electro optical module transmitter for contactless oxygen measurements</td></tr><tr><td>Non-vented male luer lock cap&amp;nbsp;</td><td>Qosina</td><td>SKU: 65418</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>O&lt;sub&gt;2&lt;/sub&gt; sensor stick</td><td>Presens</td><td>SST-PSt3-YOP</td><td>Part of noninvasive PuO2 monitor</td></tr><tr><td>PowerLab data acquisition platform</td><td>AD Instruments</td><td>N/A</td><td>For data collection</td></tr><tr><td>REBOA catheter</td><td>Certus Critical Care</td><td>N/A</td><td>Used in experimental protocol</td></tr><tr><td>Super Sheath arterial catheters (5 Fr, 7 Fr, 9 Fr)</td><td>Boston Scientific</td><td>C1894</td><td>for intravascular access</td></tr><tr><td>Suture</td><td>Ethicon</td><td>C013D</td><td>For securing catheter to skin and closing incisions</td></tr><tr><td>T connector, all female luer locks</td><td>Qosina</td><td>SKU: 88214</td><td>Part of noninvasive PuO2 monitor</td></tr></tbody></table>

noninvasive and invasive renal hypoxia monitoring in a porcine model of hemorrhagic shock

Up to 50% of patients with trauma develop acute kidney injury (AKI), in part due to poor renal perfusion after severe blood loss. AKI is currently diagnosed based on a change in serum creatinine concentration from baseline or prolonged periods of decreased urine output. Unfortunately, baseline serum creatinine concentration data is unavailable in most patients with trauma, and current estimation methods are inaccurate. In addition, serum creatinine concentration may not change until 24-48 h after the injury. Lastly, oliguria must persist for a minimum of 6 h to diagnose AKI, making it impractical for early diagnosis. AKI diagnostic approaches available today are not useful for predicting risk during the resuscitation of patients with trauma. Studies suggest that urinary partial pressure of oxygen (PuO2) may be useful for assessing renal hypoxia. A monitor that connects the urinary catheter and the urine collection bag was developed to measure PuO2 noninvasively. The device incorporates an optical oxygen sensor that estimates PuO2 based on luminescence quenching principles. In addition, the device measures urinary flow and temperature, the latter to adjust for confounding effects of temperature changes. Urinary flow is measured to compensate for the effects of oxygen ingress during periods of low urine flow. This article describes a porcine model of hemorrhagic shock to study the relationship between noninvasive PuO2, renal hypoxia, and AKI development. A key element of the model is the ultrasound-guided surgical placement in the renal medulla of an oxygen probe, which is based on an unsheathed optical microfiber. PuO2 will also be measured in the bladder and compared to the kidney and noninvasive PuO2 measurements. This model can be used to test PuO2 as an early marker of AKI and assess PuO2 as a resuscitative endpoint after hemorrhage that is indicative of end-organ rather than systemic oxygenation.

Figure 1 shows an image of the noninvasive PuO2 monitor described in this manuscript. Figure 2 shows a plot of MAP and noninvasive PuO2 measurements in a single subject during an experiment similar to the described porcine hemorrhage model. At the start of the experiment, as hemorrhage was initiated, there was a drop in MAP and PuO2. Following the initial decline in PuO2 it gradually increased until after the REBOA bal...

Watch this Scientific Journal Video about Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock at JoVE.com

Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock

Presented here is a protocol to measure renal oxygenation in the medulla and noninvasive urine oxygen partial pressure in a hemorrhagic shock porcine model to establish urine oxygen partial pressure as an early indicator of acute kidney injury (AKI) and a novel resuscitative endpoint.

Up to 50% of patients with trauma develop acute kidney injury (AKI), in part due to poor renal perfusion after severe blood loss. AKI is currently ...

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Research

JoVE Journal

Bioengineering

1.1K Views.  University of Utah. Presented here is a protocol to measure renal oxygenation in the medulla and noninvasive urine oxygen partial pressure in a hemorrhagic shock porcine model to establish urine oxygen partial pressure as an early indicator of acute kidney injury (AKI) and a novel resuscitative endpoint. 

Video: Noninvasive and Invasive Renal Hypoxia Monitoring in a Porcine Model of Hemorrhagic Shock

A Swine Model of Neonatal Asphyxia

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, perfusion deficit, hypoxic-ischemic encephalopathy, pulmonary hypertension, vasculopathic enterocolitis, renal failure and thrombo-embolic complications. Animal models are developed to help us understand the patho-physiology and pharmacology of neonatal asphyxia. In comparison to rodents and newborn lambs, the newborn piglet has been proven to be a valuable model. The newborn piglet has several advantages including similar development as that of 36-38 weeks human fetus with comparable body systems, large body size (&tilde;1.5-2 kg at birth) that allows the instrumentation and monitoring of the animal and controls the confounding variables of hypoxia and hemodynamic derangements.
We here describe an experimental protocol to simulate neonatal asphyxia and allow us to examine the systemic and regional hemodynamic changes during the asphyxiating and reoxygenation process as well as the respective effects of interventions. Further, the model has the advantage of studying multi-organ failure or dysfunction simultaneously and the interaction with various body systems. The experimental model is a non-survival procedure that involves the surgical instrumentation of newborn piglets (1-3 day-old and 1.5-2.5 kg weight, mixed breed) to allow the establishment of mechanical ventilation, vascular (arterial and central venous) access and the placement of catheters and flow probes (Transonic Inc.) for the continuously monitoring of intra-vascular pressure and blood flow across different arteries including main pulmonary, common carotid, superior mesenteric and left renal arteries. Using these surgically instrumented piglets, after stabilization for 30-60 minutes as defined by Z&lt;10% variation in hemodynamic parameters and normal blood gases, we commence an experimental protocol of severe hypoxemia which is induced via normocapnic alveolar hypoxia. The piglet is ventilated with 10-15% oxygen by increasing the inhaled concentration of nitrogen gas for 2h, aiming for arterial oxygen saturations of 30-40%. This degree of hypoxemia will produce clinical asphyxia with severe metabolic acidosis, systemic hypotension and cardiogenic shock with hypoperfusion to vital organs. The hypoxia is followed by reoxygenation with 100% oxygen for 0.5h and then 21% oxygen for 3.5h. Pharmacologic interventions can be introduced in due course and their effects investigated in a blinded, block-randomized fashion.

Large animal models have good translational values in the examination of physiology and pharmacology of neonatal asphyxia. Using newborn piglets, we develop an experimental protocol to simulate neonatal asphyxia which has advantages of studying the systemic and regional hemodynamics, oxygen transport with biochemical and pathologic pathways and correlations.

Annually more than 1 million neonates die worldwide as related to asphyxia. Asphyxiated neonates commonly have multi-organ failure including hypotension, ...

Medicine

Invasive Hemodynamic Monitoring of Aortic and Pulmonary Artery Hemodynamics in a Large Animal Model of ARDS

One of the leading causes of morbidity and mortality in patients with heart failure is right ventricular (RV) dysfunction, especially if it is due to pulmonary hypertension. For a better understanding and treatment of this disease, precise hemodynamic monitoring of left and right ventricular parameters is important. For this reason, it is essential to establish experimental pig models of cardiac hemodynamics and measurements for research purpose. 
This article shows the induction of ARDS by using oleic acid (OA) and consequent right ventricular dysfunction, as well as the instrumentation of the pigs and the data acquisition process that is needed to assess hemodynamic parameters. To achieve right ventricular dysfunction, we used oleic acid (OA) to cause ARDS and accompanied this with pulmonary artery hypertension (PAH). With this model of PAH and consecutive right ventricular dysfunction, many hemodynamic parameters can be measured, and right ventricular volume load can be detected. 
All vital parameters, including respiratory rate (RR), heart rate (HR) and body temperature were recorded throughout the whole experiment. Hemodynamic parameters including femoral artery pressure (FAP), aortic pressure (AP), right ventricular pressure (peak systolic, end systolic and end diastolic right ventricular pressure), central venous pressure (CVP), pulmonary artery pressure (PAP) and left arterial pressure (LAP) were measured as well as perfusion parameters including ascending aortic flow (AAF) and pulmonary artery flow (PAF). Hemodynamic measurements were performed using transcardiopulmonary thermodilution to provide cardiac output (CO). Furthermore, the PiCCO2 system (Pulse Contour Cardiac Output System 2) was used to receive parameters such as stroke volume variance (SVV), pulse pressure variance (PPV), as well as extravascular lung water (EVLW) and global end-diastolic volume (GEDV). Our monitoring procedure is suitable for detecting right ventricular dysfunction and monitoring hemodynamic findings before and after volume administration.

We present a protocol of creating right ventricular dysfunction in a pig model by inducing ARDS. We demonstrate invasive monitoring of left and right ventricular cardiac output using flow probes around the aorta and the pulmonary artery, as well as blood pressure measurements in the aorta and pulmonary artery.

One of the leading causes of morbidity and mortality in patients with heart failure is right ventricular (RV) dysfunction, especially if it is due to ...

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 ...

Invasive Hemodynamic Assessment for the Right Ventricular System and Hypoxia-Induced Pulmonary Arterial Hypertension in Mice

Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder. Mice are a popular animal model used to mimic this disease. However, the evaluation of right ventricular pressure (RVP) and pulmonary artery pressure (PAP) remains technically challenging in mice. RVP and PAP are more difficult to measure than left ventricular pressure because of the anatomical differences between the left and right heart systems. In this paper, we describe a stable right heart hemodynamic measurement method and its validation using healthy and PAH mice. This method is based on open-chest surgery and mechanical ventilation support. It is a complicated procedure compared to closed chest procedures. While a well-trained surgeon is required for this surgery, the advantage of this procedure is that it can generate both RVP and PAP parameters at the same time, so it is a preferable procedure for the evaluation of PAH models.

Here, we present a protocol to perform an invasive hemodynamic assessment of the right ventricle and pulmonary artery in mice using an open-chest surgery approach.

Pulmonary arterial hypertension (PAH) is a chronic and severe cardiopulmonary disorder. Mice are a popular animal model used to mimic this disease. ...

Orthotopic Kidney Auto-Transplantation in a Porcine Model Using 24 Hours Organ Preservation And Continuous Telemetry

In the present era of organ transplantation with critical organ shortage, various strategies are employed to expand the pool of available allografts for kidney transplantation (KT). Even though, the use of allografts from extended criteria donors (ECD) could partially ease the shortage of organ donors, ECD organs carry a potentially higher risk for inferior outcomes and postoperative complications. Dynamic organ preservation techniques, modulation of ischemia-reperfusion and preservation injury, and allograft therapies are in the spotlight of scientific interest in an effort to improve allograft utilization and patient outcomes in KT.
Preclinical animal experiments are playing an essential role in translational research, especially in the medical device and drug development. The major advantage of the porcine orthotopic auto-transplantation model over ex vivo or small animal studies lies within the surgical-anatomical and physiological similarities to the clinical setting. This allows the investigation of new therapeutic methods and techniques and ensures a facilitated clinical translation of the findings. This protocol provides a comprehensive and problem-oriented description of the porcine orthotopic kidney auto-transplantation model, using a preservation time of 24 hours and telemetry monitoring. The combination of sophisticated surgical techniques with highly standardized and state-of-the-art methods of anesthesia, animal housing, perioperative follow up, and monitoring ensure the reproducibility and success of this model.

Large animal models play an essential role in preclinical transplantation research. Due to its similarities to the clinical setup, the porcine model of orthotopic kidney auto-transplantation described in this article provides an excellent in vivo setting for the testing of organ preservation techniques and therapeutic interventions.

In the present era of organ transplantation with critical organ shortage, various strategies are employed to expand the pool of available allografts for ...

Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its incidence is still considerably high and therefore, influences patient outcome. Microcirculation plays a key role in tissue perfusion and oxygen supply and is often dissociated from macrohemodynamics. Thus, direct evaluation of spinal cord microcirculation is essential for the development of microcirculation-targeted therapies and the evaluation of existing approaches in regard to spinal cord microcirculation. However, most of the methods do not provide real-time assessment of spinal cord microcirculation. The aim of this study is to describe a standardized protocol for real-time spinal cord microcirculatory evaluation using laser-Doppler needle probes directly inserted in the spinal cord. We used a porcine model of ischemia/reperfusion to induce deterioration of the spinal cord microcirculation. In addition, a fluorescent microsphere injection technique was used. Initially, animals were anesthetized and mechanically ventilated. Thereafter, laser-Doppler needle probe insertion was performed, followed by the placement of cerebrospinal fluid drainage. A median sternotomy was performed for exposure of the descending aorta to perform aortic cross-clamping. Ischemia/reperfusion was induced by supra-celiac aortic cross-clamping for a total of 48 min, followed by reperfusion and hemodynamic stabilization. Laser-Doppler Flux was performed in parallel with macrohemodynamic evaluation. In addition, automated cerebrospinal fluid drainage was used to maintain a stable cerebrospinal pressure. After completion of the protocol, animals were sacrificed, and the spinal cord was harvested for histopathological and microsphere analysis. The protocol reveals the feasibility of spinal cord microperfusion measurements using laser-Doppler probes and shows a marked decrease during ischemia as well as recovery after reperfusion. Results showed comparable behavior to fluorescent microsphere evaluation. In conclusion, this new protocol might provide a useful large animal model for future studies using real-time spinal cord microperfusion assessment in ischemia/reperfusion conditions.

Spinal cord microcirculation plays a pivotal role in spinal cord injury. Most methods do not allow real-time assessment of spinal cord microcirculation, which is essential for the development of microcirculation-targeted therapies. Here, we propose a protocol using Laser-Doppler-Flow Needle probes in a large animal model of ischemia/reperfusion.

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its ...

A Large Animal Model for Acute Kidney Injury by Temporary Bilateral Renal Artery Occlusion

Acute kidney injury (AKI) is associated with higher risk for morbidity and mortality post-operatively. Ischemia-reperfusion injury (IRI) is the most common cause of AKI. To mimic this clinical scenario, this study presents a highly reproducible large animal model of renal IRI in swine using temporary percutaneous bilateral balloon-catheter occlusion of the renal arteries. The renal arteries are occluded for 60 min by introducing the balloon-catheters through the femoral and carotid artery and advancing them into the proximal portion of the arteries. Iodinated contrast is injected in the aorta to assess any opacification of the kidney vessels and confirm the success of the artery occlusion. This is furtherly confirmed by the flattening of the pulse waveform at the tip of the balloon catheters. The balloons are deflated and removed after 60 min of bilateral renal artery occlusion, and the animals are allowed to recover for 24 h. At the end of the study, plasma creatinine and blood urea nitrogen significantly increase, while eGFR and urine output significantly decrease. The need for iodinated contrast is minimal and does not affect renal function. Bilateral renal artery occlusion better mimics the clinical scenario of perioperative renal hypoperfusion, and the percutaneous approach minimizes the impact of the inflammatory response and the risk of infection seen with an open approach, such as a laparotomy. The ability to create and reproduce this clinically relevant swine model eases the clinical translation to humans.

This study presents a highly reproducible large animal model of renal ischemia-reperfusion injury in swine using temporary percutaneous bilateral balloon-catheter occlusion of the renal arteries for 60 min and reperfusion for 24 h.

Acute kidney injury (AKI) is associated with higher risk for morbidity and mortality post-operatively. Ischemia-reperfusion injury (IRI) is the most ...

Multiple Intravenous Bolus Dosing and Invasive Hemodynamic Assessment in a Hypoxia-Induced Mouse Pulmonary Artery Hypertension Model

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, there is no cure for PAH. It is important to discover new compounds that can be used to treat PAH. The mouse hypoxia-induced PAH model is a widely used model for PAH research. This model recapitulates human clinical manifestations of PAH Group 3 disease and is an important research tool to evaluate the effectiveness of new experimental therapies for PAH. Research using this model often requires the administration of compounds in mice. For a compound that needs to be given directly into the bloodstream, optimizing intravenous (IV) administration is a key part of the experimental procedures. Ideally, the IV injection system should permit multiple injections over a set time course. Although the mouse hypoxia-induced PAH model is very popular in many laboratories, it is technically challenging to perform multiple IV bolus dosing and invasive hemodynamic assessment in this model. In this protocol, we present step-by-step instructions on how to carry out multiple IV bolus dosing via mouse jugular vein and perform arterial and right ventricle catheterization for hemodynamic assessment in mouse hypoxia-induced PAH model.

This protocol provides a step-by-step procedure for executing multiple intravenous bolus dose administration and invasive hemodynamic monitoring in mice. Investigators can use this protocol for future therapeutic compound screening for pulmonary artery hypertension.

Pulmonary arterial hypertension (PAH) is a progressive life-threatening disease, primarily affecting small pulmonary arterioles of the lung. Currently, ...

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 ...