This work was supported by &#34;Soci&#233;t&#233; Fran&#231;aise d&#39;Anesth&#233;sie et de R&#233;animation&#34; (Paris, France), &#34;Fondation d&#39;entreprises Genavie&#34; (Nantes, France), &#34;F&#233;d&#233;ration fran&#231;aise de cardiologie&#34; (France), &#34;Agence nationale de la recherche&#34; (20-ASTC-0032-01-hErOiSmE) (Paris, France) and &#34;Direction G&#233;n&#233;rale de l&#39;Armement&#34; (Paris, France). Thomas Dupas was supported by grants from Direction G&#233;n&#233;rale de l&#39;Armement (DGA), France and R&#233;gion des Pays de la Loire during his PhD. Antoine Persello was supported by grants from InFlectis BioScience, France during his PhD. We thank the &#34;Agence Nationale de la Recherche&#34; (Paris, France), &#34;Direction G&#233;n&#233;rale de l&#39;Armement&#34; (Paris, France), and &#34;Sauve ton coeur&#34; association (France) to support this work. We thank the UTE IRS-UN core facility (SFR Bonamy, Nantes Universit&#233;, Nantes, France) and the IBISA core facility Therassay (Nantes, France) for their assistance and technical support.

All the procedures were approved and performed in compliance with the regional ethics committee (protocol (#17858 and #32499, CEEA-Pays de la Loire, France) according to Directive 2010/63/EU of the European Union. The reporting is in accordance with current ARRIVE guidelines and the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals (NIH Pub. No. 85-23, revised 2011).
1. Ethical status and general information on rats
<ol>
	<li>Male Wistar Han rats (Charles River, Saint-Germain-Nuelles, France) were delivered to the Unit&#233; Th&#233;rapeutique Exp&#233;rimentale at 10 weeks of age. House the rats for at least 1 week under standard temperature (21-24 &#176;C), humidity (40%-60%), and 12 h light/dark cycle with a light period starting at 07:30 a.m.</li>
	<li>Provide food and water ad libitum. Use 11-13 weeks-old rats weighing 300-400 g for the protocol.</li>
</ol>
2. Room set-up and preparation steps
<ol>
	<li>Switch on the anesthesia station, set the heating mat to 37.5 &#176;C, and cover it with a non-sterile drape.</li>
	<li>Sterilize the surgical instruments required for the protocol: DeBakey atraumatic forceps (1), fine and sharp scissors or scalpel (1), standard pattern forceps (2), needle holders (1), Vannas micro dissecting scissors (1), clamp (1).</li>
	<li>Prepare catheters for the jugular vein and femoral artery.
	<ol>
		<li>For the jugular vein catheter, mount Polyethylene (PE) tubing (PE50) on a 23 G needle with the tip cut off at the end.</li>
		<li>For the femoral artery catheter, prepare the one described in step 2.3.1 and attach a PE10 tubing to the end of the PE50 tubing.</li>
	</ol>
	</li>
	<li>Prepare the pressure transducer and catheters for the femoral artery and jugular vein by filling it with lactated Ringer&#39;s solution heparinized at 100 UI/mL. Place a 2 mL syringe containing lactated Ringer&#39;s solution heparinized at 100 UI/mL at the end of the catheters and the pressure transducer. Be careful not to get any bubbles, as these cause signal defects.</li>
	<li>Switch on the software associated with the pressure transducer and calibrate it according to the supplier&#39;s instructions.</li>
</ol>
3. Preparing the rat for surgery
<ol>
	<li>Anesthetize the rat with an induction box (parameters: sevoflurane 8%, air flow rate: 1 L/min). After checking the depth of anesthesia via pedal reflex, maintain anesthesia at sevoflurane 4%, with an airflow rate of 0.6 L/min. The use of analgesics preoperatively was avoided because of the haemodynamic effects of these molecules. 
	NOTE: From this step, the depth of anesthesia is assessed every 20 min via pedal reflex.</li>
	<li>Weigh the rat, place it in the dorsal decubitus position on a heating mat, and depilate it at the inguinal and neck regions.</li>
	<li>Disinfect depilated areas with alternating 10% and 4% povidone-iodine solutions (3 times each) using nonsterile gauze pads.</li>
	<li>Apply a drop of ophthalmic ointment to the animal&#39;s eyes to prevent drying.</li>
	<li>Place the rat on a heating mat on the surgical table (dorsal decubitus). Place the lubricated rectal probe to control the rat&#39;s temperature. Maintain anesthesia at sevoflurane 4%, with an airflow rate of 0.6 L/min.</li>
	<li>Change gloves for sterile ones, place the sterile drape on the rat, and cut it at the inguinal and neck regions.</li>
</ol>
4. Jugular vein cannulation
<ol>
	<li>Locate the jugular region in the lower right neck region, above the clavicle, by visible pulsation. Utilizing the DeBakey atraumatic forceps, gently grasp the skin, then make a precise incision using fine, sharp scissors or a scalpel.</li>
	<li>Apply a drop of local anesthetic (lidocaine 2%) to the incised area. Gently slash tissue with standard pattern forceps.</li>
	<li>Locate the jugular vein and gently liberate it from its muscular compartment with standard pattern forceps. Place a 4/0 silk suture thread and securely ligate the distal side (toward the head). Use the suture with the needle holders to tension the jugular vein.</li>
	<li>Place a 4/0 suture on the proximal side (toward the heart) and prepare a surgeon knot without tightening it. Make a small incision in the jugular vein using Vannas micro dissecting scissors.</li>
	<li>After carefully grasping the vein wall with small forceps, insert the PE50 catheter in the jugular vein by hand or with a standard pattern forceps. At this stage, use a sterile gauze pad to wipe eventual drops of blood, which may leak from the vein.</li>
	<li>Advance the catheter slightly (0.5 cm) and verify its correct placement in the vein by drawing a small amount of blood and then re-injecting it (blood in the cannula indicates it is in the vein). Tighten the previously prepared knot to secure the catheter.</li>
	<li>Leave the catheter in place, attached to a syringe pre-filled with lactated Ringer&#39;s solution, and cover the incised area with a moistened sterile gauze pad.</li>
</ol>
5. Femoral artery cannulation
<ol>
	<li>Use DeBakey atraumatic forceps to grasp the skin of the Scarpa triangle of the left leg and make an incision with fine and sharp scissors or a scalpel. Apply a drop of local anesthetic (lidocaine 2%) to the incised area. Gently dilacerate tissue with standard pattern forceps.</li>
	<li>Locate the femoral triad (artery, vein, and nerve). Pass one forceps under the triad and, using the second standard pattern forceps, very gently separate the artery from the nerve and femoral vein. 
	NOTE: The artery is smaller, pink, and pulsating. This is a critical step; the vein and artery are fragile and can easily rupture.</li>
	<li>Place a 4/0 suture and ligate the femoral artery distally (leg side). Take the suture with the needle holders to put in tension the femoral artery. Place a 4/0 suture on the proximal side (heart side) and prepare a knot without closing it. Clamp the artery (place the clamp upstream of the proximal side suture).</li>
	<li>Incise the femoral artery transversely with Vannas micro dissecting scissors (a small volume of blood should flow from the artery). Using standard pattern forceps, gently grasp the artery wall and cannulate the femoral artery with the catheter (PE10 ended), holding it with forceps.</li>
	<li>Gently unclamp to check for leaks and then check the pressure signal (validates that the catheter is in the femoral artery), and check that blood is not flowing back into the catheter (sign of leakage from the valves/pressure transducer).</li>
	<li>If the signal is good (expected values: systolic blood pressure: 120 mmHg, diastolic blood pressure: 80 mmHg) and there are no signs of leakage, advance the catheter slightly (0.5 cm) and tighten the previously prepared surgeon knot.</li>
	<li>Heparinize the animal at 100 UI/kg and apply a moistened sterile gauze pad over the incised area. After surgery, maintain anesthesia at sevoflurane 3% at an air flow rate of 0.6 L/min.</li>
</ol>
6. Hemorrhagic shock protocol (Figure 1)
<ol>
	<li>Phase 1: Stabilization (5 min).
	<ol>
		<li>After heparinization, wait 10 min for pressure values to stabilize. Make a 5 min recording for basal hemodynamic values.</li>
	</ol>
	</li>
	<li>Phase 2: Exsanguination (5 min): 
	NOTE: This phase corresponds to the fixed-volume phase of the model.
	<ol>
		<li>Draw 5 mL of blood from the femoral artery over 5 min (500 &#181;L/30 s) using a 1 mL syringe (expected values: systolic blood pressure: 45 mmHg, diastolic blood pressure: 30 mmHg).</li>
		<li>Prepare a mixture of 50% lactated Ringer&#39;s solution with 50% collected blood at room temperature. Place the mixture in a 2 mL syringe and place it at the end of the jugular vein cannula.</li>
	</ol>
	</li>
	<li>Phase 3: Hemorrhagic shock (75 min) 
	NOTE: This phase is the fixed-pressure phase of the model.
	<ol>
		<li>Maintain mean arterial pressure at an average of 35 mmHg. When mean arterial pressure is over or equal to 38 mmHg, withdraw 200 &#181;L of blood via the femoral artery (mean on n = 12 rats: 10.2 mL).</li>
		<li>If the mean arterial pressure drops below 32 mmHg, inject 200 &#181;L of a 50% blood-50% lactated Ringer&#39;s solution mixture via the jugular vein (mean on n = 12 rats: 0.90 mL).</li>
		<li>Measure peripheral blood lactate on a blood drop from the end of the tail using the tip of a needle (26 G) after sterile preparation of the tail with a gauze pad at the end of the hemorrhagic shock phase.</li>
	</ol>
	</li>
	<li>Phase 4: Intravenous fluid resuscitation (20 min)
	<ol>
		<li>Resuscitate with a lactated Ringer&#39;s solution at room temperature (RT) at 10 mL/kg over 20 min (flow rate of 10.5 mL/h for a 350 g rat) with a 20 mL syringe through the jugular catheter with a syringe pump.</li>
	</ol>
	</li>
</ol>
7. End of surgery and recovery and post-surgical follow-up
<ol>
	<li>Clamp the jugular vein and the femoral artery and remove the catheter. Ligate vessels. Check carefully that no blood is leaking out.</li>
	<li>Add a drop of local anesthetic to the incised areas. Suture incised areas with subcutaneous and cutaneous stitches using 5-0 sterile suture. Disinfect with 10% povidone-iodine solution.</li>
	<li>Subcutaneously inject buprenorphine (0.05 mg/kg, 0.3 mL/kg) using a 1 mL syringe with a 26 G needle.</li>
	<li>Wait for the rat to wean from anesthesia while monitoring temperature. When it shows signs of waking up (vibrissae moving, paw movements), remove the rectal probe and place the rat in its cage on a heating mat. 10 min later, return the rat to his accommodation room.</li>
	<li>Administer buprenorphine subcutaneously (0.05 mg/kg, 0.3 mL/kg) every 8 h.</li>
	<li>Evaluate respiration rate, behavior, temperature, and blood lactate 16 h after hemorrhagic shock induction, as described in step 6.3.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66523/66523fig01.jpg" /> 
Figure 1: Model of mixed rat hemorrhagic shock. Created with BioRender.com <a href="https://www.jove.com/files/ftp_upload/66523/66523fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
8. 24 h after hemorrhagic shock induction
<ol>
	<li>Anesthetize the animal as described in step 3.1. Place the animal on the surgical table and insert the rectal probe as described in step 3.5. Prepare the catheter for carotid artery cannulation as described for the jugular vein in steps 2.3 and 2.4.</li>
	<li>Use DeBakey atraumatic forceps to grasp the skin and make an incision in the middle of the neck with fine and sharp scissors or a scalpel. Apply a drop of local anesthetic (lidocaine 2%).</li>
	<li>Gently dilacerate tissue with standard pattern forceps. Separate the salivary glands, then open the tracheal muscle to reveal the tracheal rings.</li>
	<li>Take the left carotid artery and separate it from the nerve with standard pattern forceps (it is highly likely that the animal&#39;s breathing will accelerate). Ligate the carotid artery distally (head side) with 4/0 silk thread.</li>
	<li>Place a suture thread on the proximal side (heart side) and prepare a surgeon knot without closing it. Clamp the artery (place the clamp upstream of the proximal side suture).</li>
	<li>Incise the carotid artery with Vannas micro dissecting scissors (a small volume of blood should flow from the artery). Using standard pattern forceps, gently grasp the artery wall to enlarge the opening and cannulate the carotid artery with the catheter provided, holding it with forceps.</li>
	<li>Unclamp and check the pressure signal to confirm catheter placement, and check that blood is not flowing back into the catheter (a sign of leakage from the valves/pressure transducer).</li>
	<li>If the signal is good and there is no leakage, advance the catheter slightly (0.5cm) and tighten and secure the previously prepared surgeon knot.</li>
	<li>Heparinize the animal at 100 UI/kg and apply a moistened sterile compress over the incision.</li>
	<li>After surgery, maintain anesthesia using sevoflurane 3% at an airflow rate of 0.6 L/min.</li>
	<li>Wait 10 min to record 24 h hemodynamic values (expected values: systolic blood pressure: 120 mmHg, diastolic blood pressure: 60 mmHg).</li>
	<li>To assess plasmatic markers of organ damage, draw 1 mL of blood from the carotid artery and centrifuge at 1600 x g for 10 min. Aliquot and save plasma for further analysis.</li>
	<li>Sacrifice the rat immediately after measuring hemodynamic values at 24 h and collect blood. 
	NOTE: The method of euthanasia should be adapted to the specific parameters or samples that will be collected afterward. The sham group undergoes surgery only (sections 1-5, 7, and 8) without the procedure of hemorrhagic shock (section 6). Animals were randomly assigned by the experimenter between the sham and hemorrhagic shock groups. Only the experimenter was aware of the group allocation at the different stages of the experiment.</li>
</ol>

Vascular Cannulation with Hemorrhagic Shock Induction in Rats

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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=Inhibition+of+Bruton's+tyrosine+kinase+activity+attenuates+hemorrhagic+shock-induced+multiple+organ+dysfunction+in+rats.">Inhibition of Bruton's tyrosine kinase activity attenuates hemorrhagic shock-induced multiple organ dysfunction in rats.</a> Ann Surg. 277 (3), e624-e633 (2023).</li><li>Curcio, L., D'Orsi, L., De Gaetano, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Seven+mathematical+models+of+hemorrhagic+shock.">Seven mathematical models of hemorrhagic shock.</a> Comput Math Methods Med. 2021, e6640638 (2021).</li><li>R&#246;nn, T., Lendemans, S., de Groot, H., Petrat, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+model+of+severe+hemorrhagic+shock+in+rats.">A new model of severe hemorrhagic shock in rats.</a> Comp Med. 61 (5), 419-426 (2011).</li><li>Hussmann, B., Lendemans, S., de Groot, H., Rohrig, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Volume+replacement+with+Ringer-lactate+is+detrimental+in+severe+hemorrhagic+shock+but+protective+in+moderate+hemorrhagic+shock:+studies+in+a+rat+model.">Volume replacement with Ringer-lactate is detrimental in severe hemorrhagic shock but protective in moderate hemorrhagic shock: studies in a rat model.</a> Crit Care. 18 (1), 5 (2014).</li><li>Mihara, R., Takasu, A., Maemura, K., Minami, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prolonged+severe+hemorrhagic+shock+at+a+mean+arterial+pressure+of+40+mmHg+does+not+lead+to+brain+damage+in+rats.">Prolonged severe hemorrhagic shock at a mean arterial pressure of 40 mmHg does not lead to brain damage in rats.</a> Acute Med Surg. 5 (4), 350-357 (2018).</li><li>Vincent, J. -. L., De Backer, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circulatory+shock.">Circulatory shock.</a> N Engl J Med. 369 (18), 1726-1734 (2013).</li><li>Halbgebauer, 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=Hemorrhagic+shock+drives+glycocalyx,+barrier+and+organ+dysfunction+early+after+polytrauma.">Hemorrhagic shock drives glycocalyx, barrier and organ dysfunction early after polytrauma.</a> J Crit Care. 44, 229-237 (2018).</li><li>Chipman, A. M., Jenne, C., Wu, F., Kozar, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contemporary+resuscitation+of+hemorrhagic+shock:+What+will+the+future+hold.">Contemporary resuscitation of hemorrhagic shock: What will the future hold.</a> Am J Surg. 220, 580-588 (2020).</li></ol>

The authors declare that they have no known competing financial interests or personal relationships that could have influenced the work reported in this paper. The data supporting this study's findings are available from corresponding authors upon reasonable request.

In this paper, we described for the first time a representative rat model of hemorrhagic shock based on a mix between the fixed pressure and fixed volume models. We demonstrated that 24 h after the shock induction, our model is associated with an alteration of hemodynamic parameters and metabolism.
Due to its complex pathophysiology, the study of hemorrhagic shock requires the utilization of integrated animal models. Indeed, in vitro approaches cannot mimic all the pathways involved in this disease. Awakening the animals after the hemorrhagic shock protocol is a step that ensures a better replication of the clinical situation. Because of the difficulty involved in waking up the animals, very few studies have included this stage. The rare studies that wake animals up sacrifice them at short times (2 h or 6 h), which does not fully reflect what is happening for patients16,18,23,24. Despite the development of hemorrhagic shock models, only a few studies have evaluated parameters (inflammation, apoptosis, organ dysfunction) 24 h after the shock induction, thus highlighting the difficulty of this kind of protocol25,26,27. The development of computer and mathematical models has revolutionized research. Numerous mathematical models of hemorrhagic shock have been developed, but most of these models do not take into account the full range of body fluid exchanges during hemorrhagic shock and require improvement before potential clinical applicability28. To date, one of the main challenges is the development of an animal model that mimics the pathology in humans as closely as possible.
A large number of hemorrhagic shock models are described in the literature and differ via vascular approaches, volumes of blood drawn, or the targeted pressure13. More generally, the hemorrhagic shock models can be classified into 3 groups: fixed-volume hemorrhage, fixed-pressure hemorrhage, and uncontrolled hemorrhage. The standardization and reproducibility with the fixed-volume hemorrhage are difficult and explained by the blood volume/body weight ratio, which decreases linearly with the rat&#39;s weight. The fixed-pressure hemorrhage is widely used, thus explaining that the settings (targeted pressure, duration of shock) are very variable from one study to another, making it difficult to transpose results from one model to another. It is also important to point out that hemodynamic impairment, which plays a pivotal role in the pathophysiology of hemorrhagic shock, is not systematically assessed, which could increase the discrepancy in results between studies. Finally, the uncontrolled hemorrhage model, although clinically relevant, raises questions of reproducibility and ethics. In order to reconcile clinical relevance, standardization, and reproducibility as much as possible, we have developed a mixed model with both fixed-volume and fixed-pressure phases.
In the model described here, the temperature and the respiration rate are not modified 24 hours post-surgery. This can be explained by the fact that surgery is performed under sterile conditions, thus limiting the pro-inflammatory response. Hemorrhagic shock is defined as an acute circulatory failure due to blood loss associated with a drop in blood pressure. As in humans, this model of hemorrhagic shock causes a decrease in the mean arterial pressure, notably due to a decrease in diastolic blood pressure. Interestingly, and as previously described, the heart rate is unchanged after the resuscitation phase in this model of hemorrhagic shock29,30,31. The drop in mean arterial pressure is probably associated with reduced organ perfusion, leading to multivisceral dysfunction, which can be illustrated by the increase in various plasmatic markers in our model (creatininemia, cardiac troponin T, ASAT, and ALAT). The disruption in oxygen supply leads to anaerobic metabolism, which causes an increase in lactatemia32. As previously described, this model of hemorrhagic shock leads to an increase in blood lactate levels30. This increase could be associated with the ischemia caused at the level of the femoral artery. However, considering that the animals in the sham group have physiological lactatemia and underwent the same surgical procedure as the hemorrhagic shock group, it would seem that this increase is linked to hemorrhagic shock protocol. Taken together, all these data confirm that the protocol described in this study allows the development of a new relevant model of hemorrhagic shock in the rat.
The limitation of this model is the use of heparin, which is essential to reduce the natural coagulation of blood when it comes into contact with plastic materials such as cannulas. However, the use of heparin can impact the coagulopathy associated with traumatic hemorrhagic shock33. This study involves healthy male animals aged 11-13 weeks old. Considering that sex, age, and comorbidities (hypertension, diabetes, etc.) can impact the results, it would be relevant to evaluate their impact in our model. In the protocol, the resuscitation step is performed via an injection of Ringer Lactate, a crystalloid that could promote coagulopathy and tissue edema34. Although the use of blood products is optimal, these are scarce and perishable, and it could be difficult to have a sufficient stock of rat blood for the entire protocol. Blood product and crystalloids/colloids-based resuscitation hemorrhagic shock models are two complementary approaches.
The strengths of this model are 1) its high reproducibility (illustrated by the low variability in the results), 2) its ease of application (most of the instruments are classical and vascular approaches are known) and 3) its clinical relevance, notably due the animal awakening and multi-visceral dysfunction. Based on the behavioral score described in Supplementary File 1, limit points have been set up. The sacrifice will be discussed if a score above 9 is reached, according to the attached table. If a score of 11 is reached, the animal will be systematically euthanized. In this study, none of the animals reached a score higher than 8, and therefore, none was excluded from the study. This may explain why the model described here is associated with a mortality rate 3 times lower than that of the other 24 h study (16% vs. 47%)25.
The critical step of the model is the hemorrhagic shock phase. It is important to respect the pressure range of 32-38 mmHg. In fact, we observed that maintaining mean arterial pressures below 32 mmHg resulted in a rapid and abrupt drop in pressure. Conversely, maintaining a pressure above 38 mmHg does not provide a model that is sufficiently close to clinical reality. These observations are in accordance with the interval of mean arterial pressure targeted in other models13.
In conclusion, we demonstrated that the rat hemorrhagic shock model detailed in this study is clinically relevant and could be useful both in understanding pathophysiological mechanisms by identifying new biological actors/pathways and in identifying new therapeutic strategies by testing different candidate molecules.

Hemorrhagic shock (HS) is a state of shock characterized by significant blood volume loss, resulting in tissue dysoxia. HS is a complex pathology that associates hemodynamic and metabolic changes along with pro- and anti-inflammatory responses. Approximately 1.9 million deaths worldwide are attributed to the hemorrhage and its consequences each year1. Current guidelines for care primarily involve intravenous fluid administration (supplemented or not with vasoactive molecules) and oxygen therapy. However, these treatments are symptomatic and can be ineffective, which explains why the HS-associated mortality remains high2. This justifies the importance of identifying new molecular and cellular mechanisms and, thus, treatments to reduce mortality.
Animal models allow for deciphering the pathophysiological mechanisms involved in diseases and testing new therapeutic strategies. Numerous animal models of hemorrhagic shock exist in the literature. These models differ not only in the species used but also in the means of inducing HS (e.g., fixed pressure vs. fixed volume) (Table 1, Table 2)3,4,5,6,7,8,9,10,11,12,13. Also, protocols vary within the same type of model (e.g., time of hemorrhage, targeted mean arterial pressure) (Table 3)14,15,16,17,18,19,20. Considering the wide variety of existing hemorrhagic shock models and the complexity of replicating the clinical situation, the preclinical study of this pathology remains limited. The development of a reproducible, standardizable, and easy-to-implement hemorrhagic shock model is in everyone&#39;s interest. This would facilitate comparison between the various studies and thus unravel the complex pathophysiology of the hemorrhagic shock. The aim of this protocol was to develop a new clinically relevant model of hemorrhagic shock in rats using two successive phases of hemorrhage with fixed volume followed by a fixed low blood pressure phase.
Table 1: Species used as a model for hemorrhagic shock3,4,5,6,7,8,9,10,11,12,13. <a href="https://www.jove.com/files/ftp_upload/66523/Table 1.xlsx" target="_blank">Please click here to download this Table.</a>
Table 2: The different types of hemorrhagic shock13. <a href="https://www.jove.com/files/ftp_upload/66523/Table 2.xlsx" target="_blank">Please click here to download this Table.</a>
Table 3: Example of the diversity of experimental models of hemorrhagic shock in rats induced by a fixed pressure protocol. Summary of parameters for different experimental models of hemorrhagic shock. Vessels shown in red are arteries, and those shown in blue are veins. For resuscitation, the volume of blood sampled is used as a reference (blood: resuscitation with a volume identical to that of blood sampled during shock; x2: resuscitation with a volume twice that of blood sampled during shock; x4: resuscitation with a volume four times that of blood sampled during shock). MAP: Mean Arterial Pressure; RL: Ringer Lactate14,15,16,17,18,19,20. <a href="https://www.jove.com/files/ftp_upload/66523/Table 3.xlsx" target="_blank">Please click here to download this Table.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td>1 mL syringe</td>
			<td>TERUMO</td>
			<td>MDSS01SE</td>
			<td></td>
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		<tr>
			<td>2.5 mL syringe</td>
			<td>TERUMO</td>
			<td>SS*02SE1</td>
			<td></td>
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		<tr>
			<td>20 mL syringe</td>
			<td>TERUMO</td>
			<td>MDSS20ESE</td>
			<td></td>
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		<tr>
			<td>Anesthesia induction chamber</td>
			<td>TEMSEGA</td>
			<td>HUBBIV4</td>
			<td></td>
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			<td>BD Microlance 3 23 G needle</td>
			<td>Becton Dickinson</td>
			<td>300800</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Microlance 3 26 G needle</td>
			<td>Becton Dickinson</td>
			<td>304300</td>
			<td></td>
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			<td>Blood pressure transducer</td>
			<td>emka TECHNOLOGIES</td>
			<td>BP_T</td>
			<td></td>
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		<tr>
			<td>Buprecare</td>
			<td>Axience</td>
			<td>N/A</td>
			<td>1 mL vial, buprenorphine 0.3 mg/mL</td>
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		<tr>
			<td>DE BAKEY, Atraumatic Vascular Forceps</td>
			<td>ALLGAIER instrumente medical</td>
			<td>09-543-150</td>
			<td></td>
		</tr>
		<tr>
			<td>Dermal Betadine 10%</td>
			<td>Mylan</td>
			<td>N/A</td>
			<td>125 mL bottle</td>
		</tr>
		<tr>
			<td>Fine Forceps - Curved / Serrated</td>
			<td>Fine Science Tools</td>
			<td>11065-07</td>
			<td></td>
		</tr>
		<tr>
			<td>GraphPad Prism 8</td>
			<td>GraphPad by Dotmatics</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating mats</td>
			<td>TEMSEGA</td>
			<td>OPT/THERM_MATELASSTEREORATS</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin sodium</td>
			<td>PANPHARMA&#160;</td>
			<td>N/A</td>
			<td>5 mL bottle, 5,000 UI/mL</td>
		</tr>
		<tr>
			<td>IOX2 software</td>
			<td>emka TECHNOLOGIES</td>
			<td>IOX_BASE_4c + IOX_FULLCARDIO_4a</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris Scissors - ToughCut</td>
			<td>Fine Science Tools</td>
			<td>14058-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine</td>
			<td>Fresenius</td>
			<td>N/A</td>
			<td>10 mL bootle, 8.11 mg, lidocaine hydrochloride</td>
		</tr>
		<tr>
			<td>MiniHub-V3.2</td>
			<td>TEMSEGA</td>
			<td>PF006</td>
			<td></td>
		</tr>
		<tr>
			<td>Moria 201/A Vessel Clamp - Straight</td>
			<td>Fine Science Tools</td>
			<td>18320-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Non sterile compresses</td>
			<td>Raffin</td>
			<td>70189</td>
			<td></td>
		</tr>
		<tr>
			<td>Non sterile drape</td>
			<td>Dutscher&#160;</td>
			<td>30786</td>
			<td></td>
		</tr>
		<tr>
			<td>Olsen-Hegar Needle Holder with Scissors&#160;</td>
			<td>Fine Science Tools</td>
			<td>12002-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene tubing PE10</td>
			<td>PHYMEP</td>
			<td>BTPE-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene tubing PE50</td>
			<td>PHYMEP</td>
			<td>BTPE-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Rats</td>
			<td>Charles Rivers</td>
			<td>-</td>
			<td>Male WISTAR HAN (10 weeks)</td>
		</tr>
		<tr>
			<td>Rectal probe</td>
			<td>TEMSEGA</td>
			<td>SONDE_TEMP_RATS</td>
			<td></td>
		</tr>
		<tr>
			<td>Ringer Lactates</td>
			<td>Fresenius Kabi</td>
			<td>964175</td>
			<td></td>
		</tr>
		<tr>
			<td>Scrub Betadine 4%</td>
			<td>Mylan</td>
			<td>N/A</td>
			<td>125 mL bottle</td>
		</tr>
		<tr>
			<td>Sevoflurane</td>
			<td>Abbott</td>
			<td>N/A</td>
			<td>250 mL bottle, gas 100%</td>
		</tr>
		<tr>
			<td>Sevoflurane Vaporizer</td>
			<td>TEMSEGA</td>
			<td>SEVOTEC3NSELEC</td>
			<td></td>
		</tr>
		<tr>
			<td>StatStrip lactate test strips</td>
			<td>Nova Biomedical</td>
			<td>47486</td>
			<td></td>
		</tr>
		<tr>
			<td>StatStrip Xpress lactate Meter</td>
			<td>Nova Biomedical</td>
			<td>47486</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile compresses</td>
			<td>Laboratoire SYLAMED</td>
			<td>211S05-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile drape</td>
			<td>M&#246;lnlycke</td>
			<td>800330</td>
			<td></td>
		</tr>
		<tr>
			<td>Steriles gloves</td>
			<td>MEDLINE</td>
			<td>MSG7275</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture</td>
			<td>Optilene</td>
			<td>3097141</td>
			<td></td>
		</tr>
		<tr>
			<td>Suture for vessels</td>
			<td>SMI</td>
			<td>8150046</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td>Vial m&#233;dical</td>
			<td>16010</td>
			<td></td>
		</tr>
		<tr>
			<td>usbAMP</td>
			<td>emka TECHNOLOGIES</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Vaseline</td>
			<td>Cooper</td>
			<td>N/A</td>
			<td>10 mL vial</td>
		</tr>
		<tr>
			<td>Vitamin A Dulcis (ALLERGAN)</td>
			<td>Allergan</td>
			<td>N/A</td>
			<td>10 g tube, Retinol</td>
		</tr>
	</tbody>
</table>

developing a clinically relevant hemorrhagic shock model in rats

Over the recent decades, the development of animal models allowed us to better understand various pathologies and identify new treatments. Hemorrhagic shock, i.e., organ failure due to rapid loss of a large volume of blood, is associated with a highly complex pathophysiology involving several pathways. Numerous existing animal models of hemorrhagic shock strive to replicate what happens in humans, but these models have limits in terms of clinical relevance, reproducibility, or standardization. The aim of this study was to refine these models to develop a new model of hemorrhagic shock. Briefly, hemorrhagic shock was induced in male Wistar Han rats (11-13 weeks old) by a controlled exsanguination responsible for a drop in the mean arterial pressure. The next phase of 75 min was to maintain a low mean arterial blood pressure, between 32 mmHg and 38 mmHg, to trigger the pathophysiological pathways of hemorrhagic shock. The final phase of the protocol mimicked patient care with an administration of intravenous fluids, Ringer Lactate solution, to elevate the blood pressure. Lactate and behavioral scores were assessed 16 h after the protocol started, while hemodynamics parameters and plasmatic markers were evaluated 24 h after injury. Twenty-four hours post-hemorrhagic shock induction, the mean arterial and diastolic blood pressure were decreased in the hemorrhagic shock group (p&#160;&#60; 0.05). Heart rate and systolic blood pressure remained unchanged. All organ damage markers were increased with the hemorrhagic shock (p &#60; 0.05). The lactatemia and behavioral scores were increased compared to the sham group (p &#60; 0.05). In conclusion, we demonstrated that the protocol described here is a relevant model of hemorrhagic shock that can be used in subsequent studies, particularly to evaluate the therapeutic potential of new molecules.

Following the protocol described above, we evaluated several hemodynamics parameters 24 hours after the induction of the hemorrhagic shock. Basal mean arterial pressure (before the start of the hemorrhagic shock protocol) is similar between the sham and hemorrhagic shock groups (Figure 2A). As expected, the mean arterial pressure is significantly decreased with the hemorrhagic shock protocol, which can be explained by the drop in the diastolic blood pressure (Mean arterial pressure: Sham: 92 mmHg &#177; 3 mmHg; HS: 82 mmHg &#177; 2 mmHg; Diastolic blood pressure: 73 mmHg &#177; 3 mmHg; HS: 61 mmHg &#177; 2 mmHg) (Figure 2B, C). Hemorrhagic shock does not impact systolic blood pressure, pulse pressure, and heart rate (Figure 2D-F). The shock index (heart rate/systolic blood pressure ratio) and the modified shock index (MSI) (heart rate/mean blood pressure ratio) are two predictors of mortality in severe patients14,15. The higher the values are, the greater the risk of mortality. In this model, the shock index is unmodified between the two groups, while the modified shock index tends to increase in the hemorrhagic shock (MSI: Sham: 4.24 &#177; 0.11; HS: 4.70 &#177; 0.15) (Figure 2G,H).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66523/66523fig02.jpg" /> 
Figure 2: Impact of hemorrhagic shock on hemodynamics parameters. (A) Basal mean arterial pressure, (B) mean arterial pressure, (C) diastolic blood pressure, (D) systolic blood pressure, (E) pulse pressure, (F) heart rate,&#160;(G) shock index, and (H) modified shock index between Sham and hemorrhagic shock animals. Results are represented as mean &#177; SEM. Statistical significance was assessed by unpaired t-test. *: p &#60; 0.05; ***: p &#60; 0.001. n = 6-12. <a href="https://www.jove.com/files/ftp_upload/66523/66523fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Global metabolic impairment during hemorrhagic shock can be assessed by lactatemia. As expected, the lactatemia increased after the hemorrhagic shock protocol and 16 h after (End of protocol: Sham: 1.13 mmol/L &#177; 0.14 mmol/L; HS: 5.98 mmol/L &#177; 0.39 mmol/L; H+16: Sham: 1.95 mmol/L &#177; 0.23 mmol/L; HS: 2.95 mmol/L &#177; 0.19 mmol/L) (Figure 3A,B). Temperature and respiration rate are two components of the Systemic Inflammatory Response Syndrome (SIRS), a pro-inflammatory response characteristic of the state of shock. Neither temperature nor respiration rate are modified between the two groups 16 h after the hemorrhagic shock induction (Figure 3C,D). We evaluated the impact of the hemorrhagic shock on a few behavior parameters like posture, activity, etc. (Supplementary File 1). The behavioral score is increased in the hemorrhagic shock group 16 h after the protocol (Sham: 0.33 &#177; 0.21; HS: 2.27 &#177; 0.69) (Figure 3E).
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66523/66523fig03.jpg" /> 
Figure 3: Impact of hemorrhagic shock on lactatemia, temperature, respiration rate, and behavioral score. (A) Lactatemia at the end of the hemorrhagic shock protocol, (B) lactatemia, (C) temperature, (D) respiration rate, and (E) behavioral score 16 h after hemorrhagic shock induction between Sham and hemorrhagic shock animals. Results are represented as mean &#177; SEM. Statistical significance was assessed by unpaired t-test. *: p &#60; 0.05; **: p &#60; 0.01; ***: p &#60; 0.001. n = 6-12. <a href="https://www.jove.com/files/ftp_upload/66523/66523fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The hemorrhagic shock is associated with an organ dysfunction. In order to evaluate if the model could be clinically relevant, we assessed plasmatic markers of organ injury 24 h after the protocol. The creatininemia (Sham: 19.13 &#181;mol/L &#177; 0.33 &#181;mol/L; HS: 28.88 &#181;mol/L &#177; 2.69 &#181;mol/L), the cardiac troponin T (Sham: 9.38 ng/L &#177; 1.87 ng/L; HS: 35.62 ng/L &#177; 2.28 ng/L), and the aspartate and alanine amino transferase (ASAT: Sham: 221 UI/L &#177; 48 UI/L; HS: 963 UI/L &#177; 144 UI/L; ALAT: Sham: 36 UI/L &#177; 4 UI/L; HS: 323 UI/L &#177; 13 UI/L) which reflect damages to the kidney, heart and liver respectively are all significantly increased with the hemorrhagic shock (Figure 4).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66523/66523fig04.jpg" /> 
Figure 4: The hemorrhagic shock model is associated with organ dysfunction. (A) Creatininemia, (B) cardiac troponin T, (C) aspartate aminotransferase, and (D) alanine aminotransferase levels 24 h after hemorrhagic shock induction between Sham and hemorrhagic shock animals. Results are represented as mean &#177; SEM. Statistical significance was assessed by unpaired t-test. *: p &#60; 0.05; **: p &#60; 0.01; ***: p &#60; 0.001. n = 4 <a href="https://www.jove.com/files/ftp_upload/66523/66523fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary file 1: Behavioral score details <a href="https://www.jove.com/files/ftp_upload/66523/Supplementary File 1.zip" target="_blank">Please click here to download this File.</a>

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Author Spotlight: Developing Innovative Therapeutic Strategies for Hemorrhagic Shock Research

Hemorrhagic shock kills 1.9 million people worldwide every year. Small animals are frequently used as hemorrhagic shock models but are associated with issues of standardization, reproducibility, and clinical significance, thus limiting their relevance. This article describes developing a new clinically relevant hemorrhagic shock model in rats.

Developing a Clinically Relevant Hemorrhagic Shock Model in Rats

Over the recent decades, the development of animal models allowed us to better understand various pathologies and identify new treatments. Hemorrhagic ...

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1.1K Views.  Nantes Université. Hemorrhagic shock kills 1.9 million people worldwide every year. Small animals are frequently used as hemorrhagic shock models but are associated with issues of standardization, reproducibility, and clinical significance, thus limiting their relevance. This article describes developing a new clinically relevant hemorrhagic shock model in rats.

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

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

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

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

A Contusion Model of Severe Spinal Cord Injury in Rats

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is described to obtain a consistent model of severe SCI. Use of a stereotactic frame and computer controlled impactor allows for creation of reproducible injury. Hypothermia and urinary tract infection pose significant challenges in the post-operative period. Careful monitoring of animals with daily weight recording and bladder expression allows for early detection of post-operative complications. The functional results of this contusion model are equivalent to transection models. The contusion model can be utilized to evaluate the efficacy of both neuroprotective and neuroregenerative approaches.

A contusion model of severe spinal cord injury is described. Detailed pre-operative, operative and post-operative steps are described to obtain a consistent model.

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is ...

Optimization of High Grade Glioma Cell Culture from Surgical Specimens for Use in  Clinically Relevant Animal Models and 3D Immunochemistry

Glioblastomas, the most common and aggressive form of astrocytoma, are refractory to therapy, and molecularly heterogeneous. The ability to establish cell cultures that preserve the genomic profile of the parental tumors, for use in patient specific in vitro and in vivo models, has the potential to revolutionize the preclinical development of new treatments for glioblastoma tailored to the molecular characteristics of each tumor.
Starting with fresh high grade astrocytoma tumors dissociated into single cells, we use the neurosphere assay as an enrichment method for cells presenting cancer stem cell phenotype, including expression of neural stem cell markers, long term self-renewal in vitro, and the ability to form orthotopic xenograft tumors. This method has been previously proposed, and is now in use by several investigators. Based on our experience of dissociating and culturing 125 glioblastoma specimens, we arrived at the detailed protocol we present here, suitable for routine neurosphere culturing of high grade astrocytomas and large scale expansion of tumorigenic cells for preclinical studies. We report on the efficiency of successful long term cultures using this protocol&#160;and suggest affordable alternatives for culturing dissociated glioblastoma cells that fail to grow as neurospheres. We also describe in detail a protocol for preserving the neurospheres 3D architecture for immunohistochemistry. Cell cultures enriched in CSCs, capable of generating orthotopic xenograft models that preserve the molecular signatures and heterogeneity of GBMs, are becoming increasingly popular for the study of the biology of GBMs and for the improved design of preclinical testing of potential therapies.

Protocol for propagation of dissociated high grade glioma surgical specimens in serum-free neurosphere medium to select for cells with cancer stem cell phenotype. For specimens that fail to grow as neurospheres, an alternative protocol is suggested. A paraffin embedding technique for maintaining the 3D neurosphere architecture for immunocytochemistry is described.

Glioblastomas, the most common and aggressive form of astrocytoma, are refractory to therapy, and molecularly heterogeneous. The ability to establish cell ...

Implementation of In Vitro Drug Resistance Assays: Maximizing the Potential for Uncovering Clinically Relevant Resistance Mechanisms

Although targeted therapies are initially effective, resistance inevitably emerges. Several methods, such as genetic analysis of resistant clinical specimens, have been applied to uncover these resistance mechanisms to facilitate follow-up care. Although these approaches have led to clinically relevant discoveries, difficulties in attaining the relevant patient material or in deconvoluting the genomic data collected from these specimens have severely hampered the path towards a cure. To this end, we here describe a tool for expeditious discovery that may guide improvement in first-line therapies and alternative clinical management strategies. By coupling preclinical in vitro or in vivo drug selection with next-generation sequencing, it is possible to identify genomic structural variations and/or gene expression alterations that may serve as functional drivers of resistance. This approach facilitates the spontaneous emergence of alterations, enhancing the probability that these mechanisms may be observed in the patients. In this protocol we provide guidelines to maximize the potential for uncovering single nucleotide variants that drive resistance using adherent lines.

Implementation of In Vitro Drug Resistance Assays: Maximizing the Potential for Uncovering Clinically Relevant Resistance Mechanisms

Drug resistance to targeted therapeutics is widespread and the need to identify mechanisms of resistance--prior to or following clinical onset--is critical for guiding alternative clinical management strategies. Here, we present a protocol to couple derivation of drug-resistant lines in vitro with sequencing to expedite discovery of these mechanisms.

Although targeted therapies are initially effective, resistance inevitably emerges. Several methods, such as genetic analysis of resistant clinical ...

A Model to Simulate Clinically Relevant Hypoxia in Humans

In case of apnea, arterial partial pressure of oxygen (pO2) decreases, while partial pressure of carbon dioxide (pCO2) increases. To avoid damage to hypoxia sensitive organs such as the brain, compensatory circulatory mechanisms help to maintain an adequate oxygen supply. This is mainly achieved by increased cerebral blood flow. Intermittent hypoxia is a commonly seen phenomenon in patients with obstructive sleep apnea. Acute airway obstruction can also result in hypoxia and hypercapnia. Until now, no adequate model has been established to simulate these dynamics in humans. Previous investigations focusing on human hypoxia used inhaled hypoxic gas mixtures. However, the resulting hypoxia was combined with hyperventilation and is therefore more representative of high altitude environments than of apnea. Furthermore, the transferability of previously performed animal experiments to humans is limited and the pathophysiological background of apnea induced physiological changes is poorly understood. In this study, healthy human apneic divers were utilized to mimic clinically relevant hypoxia and hypercapnia during apnea. Additionally, pulse-oximetry and Near Infrared Spectroscopy (NIRS) were used to evaluate changes in cerebral and peripheral oxygen saturation before, during, and after apnea.

Hypoxia simulation in humans has usually been performed by inhaling hypoxic gas mixtures. For this study, apneic divers were used to simulate dynamic hypoxia in humans. Additionally, physiological changes in desaturation and re-saturation kinetics were evaluated with non-invasive tools such as Near-Infrared-Spectroscopy (NIRS) and peripheral oxygenation saturation (SpO2).

In case of apnea, arterial partial pressure of oxygen (pO2) decreases, while partial pressure of carbon dioxide (pCO2) increases. To avoid damage to ...

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 ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

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 ...

A Model of Glaucoma Induced by Circumlimbal Suture in Rats and Mice

The circumlimbal suture is a technique for inducing experimental glaucoma in rodents by chronically elevating intraocular pressure (IOP), a well-known risk factor for glaucoma. This protocol demonstrates a step-by-step guide on this technique in Long Evans rats and C57BL/6 mice. Under general anesthesia, a "purse-string" suture is applied on the conjunctiva, around the equator and behind the limbus of the eye. The fellow eye serves as an untreated control. Over the duration of our study, which was a period of 8 weeks for rats and 12 weeks for mice, IOP remained elevated, as measured regularly by rebound tonometry in conscious animals without topical anesthesia. In both species, the sutured eyes showed electroretinogram features consistent with preferential inner retinal dysfunction. Optical coherence tomography showed selective thinning of the retinal nerve fiber layer. Histology of the rat retina in cross-section found reduced cell density in the ganglion cell layer, but no change in other cellular layers. Staining of flat-mounted mouse retinae with a ganglion cell specific marker (RBPMS) confirmed ganglion cell loss. The circumlimbal suture is a simple, minimally invasive and cost-effective way to induce ocular hypertension that leads to ganglion cell injury in both rats and mice.

Chronic ocular hypertension is induced by applying a circumlimbal suture in rats and mice, leading to functional and structural deterioration of the retinal ganglion cells consistent with glaucoma.

The circumlimbal suture is a technique for inducing experimental glaucoma in rodents by chronically elevating intraocular pressure (IOP), a well-known ...

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.