Name: use of a central venous line for fluids, drugs and nutrient administration in a mouse model of critical illness
Uploaded: 2017-05-02T14:45:00.000+00:00
Duration: 9 min 17 s
Description: Watch this Scientific Journal Video about Use of a Central Venous Line for Fluids, Drugs and Nutrient Administration in a Mouse Model of Critical Illness at JoVE.com

GVdB, via the University of Leuven (KU Leuven), receives long-term structural research support from the Methusalem Program funded by the Flemish Government (METH08/07) and holds a European Research Council Advanced Grant AdvG-2012-321670 from the Ideas Program of the European Union seventh framework program. SET received a Research Foundation-Flanders (FWO) Research Assistant Fellowship.

The protocol was approved by the University of Leuven Ethical Review Board for Animal Research.
1. Preparation of the Venous Line
<ol>
	<li>Prepare the tip of the venous catheter by quickly immersing the middle part of 60 cm microrenathane (MRE) tubing in hot (&#62; 220 &#176;C) sesame oil. Subsequently stretch the middle part of the tube to produce a narrow diameter (outer diameter (OD) &#60; 0.5 mm) by gently moving the ends of the tubing away from each other.</li>
	<li>Use a scalpel blade to cut the tubing into two parts of 30 cm each (see Figure 1, Table 1).</li>
	<li>Connect the MRE tubing to polyethylene tubing PE10 using polyethylene PE50 connector and connect this PE10 tubing to the bottom part of the swivel. Connect the upper part of the swivel to PE10 tubing, and connect this to a Luer stub needle with PE50 connector tubing as per Figure 1.</li>
	<li>Apply strong fast-acting adhesive glue to all connections and test the catheter for leakage by flushing with air. Gas sterilize the catheter prior to use.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/55553/55553fig1.jpg" /> 
Figure 1: Construction of Venous Line. Venous lines are prepared by stretching the MRE tubing to a small diameter and connecting it via polyethylene tubing to a rodent swivel device. See Table 1 for instructions on lengths of the different parts. <a href="http://cloudfront.jove.com/files/ftp_upload/55553/55553fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Legend</td>
			<td>Length</td>
			<td>Volume</td>
		</tr>
		<tr>
			<td>1) Luer stub needle 22 G</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>2) PE50 &#8211; polyethylene .023&#34; x .038&#34;</td>
			<td>5 cm</td>
			<td>13 &#181;L</td>
		</tr>
		<tr>
			<td>3) PE10 &#8211; polyethylene .011&#34; x .024&#8221;&#160;</td>
			<td>50 cm</td>
			<td>30 &#181;L</td>
		</tr>
		<tr>
			<td>4) Rodent Swivel 20 G</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5) PE-10&#160;</td>
			<td>15 cm</td>
			<td>9 &#181;L</td>
		</tr>
		<tr>
			<td>6) PE-50 (connector)</td>
			<td>5 cm</td>
			<td>13 &#181;L</td>
		</tr>
		<tr>
			<td>7) MRE025 Tip stretched microrenathane .025&#34; x .012&#34;</td>
			<td>30 cm</td>
			<td>27 &#181;L</td>
		</tr>
		<tr>
			<td>Total sum</td>
			<td>105 cm</td>
			<td>92 &#181;L</td>
		</tr>
	</tbody>
</table>
Table 1: Construction of Venous Line. This table provides a legend for Figure 1.
2. Anesthesia and Pre-surgery Handling
<ol>
	<li>Use 24-week old male C57BL/6J mice (27 - 32 g). 
	NOTE: We use 24-weeks-old mice (mature adult) as this age corresponds better with the mean age of intensive care patients. We use only male mice to avoid the cyclic influence of estrogens. The technique can also be used on younger animals (tested in 16-week old mice) and females, if preferred.</li>
	<li>Sterilize all surgical instruments before use. Flush the sterile home-made venous catheter with saline to remove any air bubbles and to test for patency and potential leaks.</li>
	<li>Cut a metal wire (0.8 mm diameter) of 80 cm length, double it by folding and make a loop. Lead 3 pearls over the doubled wire to the loop. Attach the catheter to the metal wire by guiding the tip of the catheter through the 3 pearls. 
	NOTE: The metal attachment wire is necessary for rotation of the swivel to which the catheter is connected. As such, the mouse can move freely, without blocking the venous line.</li>
	<li>Anesthetize the mouse by an intraperitoneal (IP) injection of a mixture of 0.03 mL ketamine (100 mg/kg) and 0.02 mL xylazine (13&#160;mg/kg). Ensure that anesthesia is adequate by checking the absence of reflexes. Gently pull the mouse tongue outside with forceps to avoid suffocation by swallowing of the tongue. 
	NOTE: If after 10 min, the mouse still attempts to withdraw its limbs give additional ketamine as a 0.01 mL bolus.</li>
	<li>Shave the surgical area including abdomen, ventral side of the neck (triangle between chin, sternum and clavicle), and between shoulder blades at the base of the head).&#160;</li>
	<li>Place the mouse in prone position on a pre-warmed heating pad, disinfect the skin with 70% ethanol and apply a small amount of ophthalmic lubricant to protect eyes from drying out. Infiltrate the surgical sites (abdomen, neck-front and back) with 0.2 mL ropivacaine (0.67 mg/kg).</li>
	<li>While the mouse is still in prone position, make a small incision at the base of the dorsal side of the head with scalpel or scissors. Expose the posterior cervical muscles, and tie the muscle to the loop of the attachment wire with 3.0 nylon by leading a 3.0 nylon thread under the muscle.</li>
	<li>Place the mouse on its right side. Make a small vertical incision in the skin of the ventral neck with a scalpel or scissors. Under visual guidance, tunnel an 18-gauge needle subcutaneously through this ventral incision towards the incision on the back made previously and thread the catheter through the needle to exteriorize it at the ventral side.</li>
	<li>Place the mouse on its back. Pass a 3.0 nylon thread behind the top incisor teeth of the mouse and tape it down to the heating pad. Secure the head in a face mask for delivering oxygen (2 L/min). Fix the mouse in a stretched position by taping down the tail and the two forelimbs. 
	NOTE: If the ketamine-xylazine anesthesia alone is not sufficient, inhaled isoflurane (0.5 - 1.5%) can be delivered during surgery.</li>
</ol>
3. Placement of a Chronic Indwelling Catheter
<ol>
	<li>Place the mouse under the dissecting microscope. In the incision made previously in the ventral neck, gently tease fat tissue and glands away with forceps until the jugular vein can be visualized.</li>
	<li>Bluntly dissect to free the jugular vein from the connective and subcutaneous tissue above and around the vessel by placing the tip of the forceps between the vein and connective tissue and opening the forceps repeatedly in parallel with the vein. As such, damage to the vein will be avoided.</li>
	<li>Isolate the jugular vein by placing blunt forceps under the vein. Feed three pieces of 3.0 silk thread under the vein, position one piece proximal to the bifurcation of the jugular vein (cranial ligature) and one piece close to the sternocleidomastoid muscle (caudal ligature). Tighten the cranial ligature and the caudal ligature to stretch the vein and prevent excessive bleeding while placing the catheter. Tie a loose ligature with the middle thread to ensure easy securing of the catheter later on.</li>
	<li>Using micro scissors, make an incision along the vein between the cranial and caudal ligatures and large enough to pass the catheter. Grab the catheter with forceps, and insert the catheter 11 mm into the vein. Loosely secure the catheter by tying off the middle ligature and confirm correct placement by gently flushing with sterile saline.</li>
	<li>Secure the catheter by firmly tying off the middle and caudal ligature around the vessel and catheter. Tie ends of caudal and middle ligature together to firmly secure the catheter. Ensure that the knots do not occlude the vein, by flushing the catheter after every knot made. Finally, tie the caudal and cranial ligatures. Close the incision with 5.0 silk sutures.</li>
</ol>
4. Cecal Ligation and Puncture
<ol>
	<li>Make a 1-cm midline incision through skin of the lower half of the abdomen with scalpel or scissors; be careful not to penetrate into the peritoneal cavity.</li>
	<li>Identify the linea alba of the abdominal musculature and make an intermuscular incision to gain entry into the peritoneal cavity. Locate the cecum, and use blunt anatomical forceps to isolate the cecum and exteriorize it.</li>
	<li>Ligate the cecum at 50% of its length with 3.0 silk sutures. Make sure not to ligate the ileocecal valve so that intestinal continuity is maintained.</li>
	<li>Perforate the cecum with an 18-gauge needle by a single through-and-through puncture midway between the ligation and the tip of the cecum. After removing the needle, extrude a small amount of feces from the hole to ensure patency.</li>
	<li>Reposition the bowel in the abdominal cavity, and close the peritoneum and skin with 5.0 silk sutures. 
	NOTE: On average, a new trainee requires 10 to 15 animals to be able to smoothly place the venous catheter and perform CLP within 45 min. After the training period, an anesthesia/surgery related mortality of 10% can be expected.</li>
</ol>
5. Post-surgical Treatment and Fluid Resuscitation
<ol>
	<li>Place the mouse in supine position and secure the catheter to the attachment wire with tape. Move the mouse to an individual cage. Use a stand with adjustable clamp to hold the swivel device 25 cm above the mouse and firmly tape the free part of the metal attachment wire to the rotating point of the swivel.</li>
	<li>Attach a syringe containing the mixture of balanced colloids and crystalloids (1:4) to the venous line to start fluid resuscitation.</li>
	<li>Place the cage in a temperature controlled (27 &#176;C) animal cabinet with 12 h light and dark cycles and start intravenous fluid resuscitation (10&#160;mL/kg/h) via an accurate syringe-driven infusion pump. Provide cage enrichment such as nesting material and a wooden block.</li>
</ol>
6. Intensive Care
<ol>
	<li>At 6 h post-operation, subcutaneously inject pain medication and antibiotics (0.3 mL buprenorphine (0.15 mg/kg) and 0.2 mL imipenem (16.67mg/kg)). NOTE: Following standard laboratory animal medicine practice, the first dose of pain meds is given prior to the initial surgical incision and then as directed by the institution&#8217;s veterinary policies. Subcutaneous injections should be carefully injected into the subcutaneous space.</li>
	<li>After 20 - 24 h of fluid resuscitation, replace the crystalloids/colloids by total parenteral nutrition (6.67&#160;mL/kg/h). 
	NOTE: Total parenteral nutrition administered (5.8 kcal/24 h) covers around 40% of the daily caloric needs of mice, similar to the early caloric intake of patients in the intensive care unit.</li>
	<li>Subcutaneously administer pain medication and antibiotics (0.6 mL buprenorphine (0.3 mg/kg) and 0.2 mL imipenem (16.67 mg/kg) every 12 h during the whole period of critical illness.</li>
	<li>Check the animals at least every 3 h during the day. Assess suffering by giving a pain score based on the validated mouse grimace scale11. Intensify monitoring for animals with a high pain score.</li>
</ol>
7.&#160;End of the Experiment
NOTE: Approval of and recommendations on the level of severity of the model and guidelines and policies for human endpoints should be sought from the local Institutional Ethical Review Board for Animal Research.
NOTE: In case of a nonfunctional venous line such as blocked catheter, delocalization of the catheter, problems with the syringe pump, the animal is excluded from the study and euthanized.
<ol>
	<li>At the end of the experimental period, deeply anesthetize the mouse using a mixture of 0.03 mL ketamine (100 mg/kg) and 0.02 mL (13&#160;mg/kg) xylazine. Euthanize the mouse by withdrawing blood by cardiac puncture. Store snap frozen tissue samples of interest at -80 &#176;C.</li>
</ol>

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S., Flierl, M. A., Ward, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunodesign+of+experimental+sepsis+by+cecal+ligation+and+puncture.">Immunodesign of experimental sepsis by cecal ligation and puncture.</a> Nat Protoc. 4 (1), 31-36 (2009).</li><li>Zolfaghari, P. S., Pinto, B. B., Dyson, A., Singer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+metabolic+phenotype+of+rodent+sepsis:+cause+for+concern.">The metabolic phenotype of rodent sepsis: cause for concern.</a> Intensive Care Med Exp. 1 (1), 25 (2013).</li><li>Radermacher, P., Haouzi, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+mouse+is+not+a+rat+is+not+a+man:+species-specific+metabolic+responses+to+sepsis+-+a+nail+in+the+coffin+of+murine+models+for+critical+care+research.">A mouse is not a rat is not a man: species-specific metabolic responses to sepsis - a nail in the coffin of murine models for critical care research.</a> Intensive Care Med Exp. 1 (1), 26 (2013).</li><li>Dyson, A., Singer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+sepsis:+why+does+preclinical+efficacy+fail+to+translate+to+the+clinical+setting.">Animal models of sepsis: why does preclinical efficacy fail to translate to the clinical setting.</a> Crit Care Med. 37 (1 Suppl), S30-S37 (2009).</li><li>Osuchowski, M. 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The authors have nothing to disclose.

We developed a more clinically relevant mouse model of critical illness, by combining the cecal ligation and puncture method to induce sepsis with the use of a central venous line for fluids, drugs and nutrient administration. This experimental setup is reproducible, allows to study the prolonged phase of critical illness and results in a stable mortality rate, hereby mimicking the human clinical situation1.
Cecal ligation and puncture has been extensively recognized as...

Critical illness is a disease state in which the function of one or more organ systems is hampered to the extent that the patient will die, unless intensive medical support is administered. Whereas the initial cause for admission to the intensive care unit (ICU) can vary, ranging from trauma, complicated surgery, burns, disease exacerbations to sepsis, all critically ill patients suffer from cellular damage, caused by hypoperfusion, hypoxia and excessive inflammation among others, which leads to organ failure. Most patients survive their acute insult, but an important fraction of the patients do not immediately recover and need prolonged intensive care. They are at high risk of death due to non-resolving multiple organ failure. Furthermore, the prolonged phase of critical illness is hallmarked by profound muscle weakness and endocrine and metabolic change,&#160; of which the pathogenesis is currently incompletely understood.
Several rodent models are being used in the critical care research setting. The two mostly used models are the exogenous administration of lipopolysaccharide (LPS) and cecal ligation and puncture (CLP). Both models are developed to mimic the acute phase of sepsis, defined as a life-threatening organ dysfunction caused by a dysregulated host response to infection, and one of the primary reasons for admission to the ICU worldwide1,2. The LPS model has several disadvantages as it only transiently affects the release of cytokines and the hemodynamic status of the animal3. Unlike humans, rodents are also particularly resistant to endotoxin and the use of &#39;high doses&#39; of endotoxin is necessary to produce hypotension and mortality, hereby further raising concerns of the validity of this method4,5. The other model, cecal ligation and puncture model (CLP) features a ligation of a portion of the cecum followed by a needle puncture through-and-through. This procedure causes a polymicrobial abdominal infection with tissue damage, followed by a translocation of the bacteria into the blood compartment. This will trigger a systemic inflammatory response and the development of sepsis. The CLP model has been extensively recognized as an animal model of acute critical illness that reproduces the main features of sepsis: hyperinflammation, vasodilation, hypotension and increased cardiac output6,7. However, this model does not allow study of non-resolving multiple organ failure, muscle wasting, and endocrine and metabolic changes, which are typical for the prolonged phase of critical illness. Furthermore, recently the validity of mouse models for critical illness have been questioned, since findings from mouse models cannot always be translated to the human setting8,9,10. A possible explanation might be that the supportive care that is provided to critically ill human patients differs substantially from the care that is provided to critically ill mice.
Therefore, to resemble the human setting more closely and to allow investigation of the prolonged phase of critical illness, we developed a mouse model that mimics the acute intensive care as given to humans, such as extensive intravenous fluid administration and antibiotic treatment, and which allows to administer supportive care in order to survive the prolonged phase of critical illness, such as nutritional support. For this purpose, we adapted the mouse model of CLP-induced sepsis, being the golden standard for sepsis, and placed a central venous line which enables the administration of fluids, nutrition and drugs.

<table><tbody><tr><td>Buprenorphine</td><td>Ecuphar</td><td></td><td>Vetergesic 0.3 mg/mL</td></tr><tr><td>C57BL/6&amp;nbsp;</td><td>Janvier labs</td><td>C57BL/6JRj</td><td></td></tr><tr><td>colloids</td><td>Fresenius Kabi</td><td></td><td>Volulyte 6%</td></tr><tr><td>crystalloids</td><td>Baxter</td><td></td><td>Plasmalyte a viaflo</td></tr><tr><td>Ethilon 3.0</td><td>Ethicon</td><td>F3211</td><td></td></tr><tr><td>Imipenem</td><td>MSD</td><td></td><td>Tienam 500&amp;nbsp;mg powder for injection fluid</td></tr><tr><td>Isoflurane</td><td>Eurovet</td><td></td><td>Iso-vet</td></tr><tr><td>Ketamine</td><td>Eurovet</td><td></td><td>Nimatek 100 mg/mL</td></tr><tr><td>LocTite Super glue3 all plastics&amp;nbsp;</td><td>Rectavit</td><td>119818</td><td></td></tr><tr><td>Mersilk 3.0</td><td>Ethicon</td><td>L192</td><td></td></tr><tr><td>Mersilk 5.0</td><td>Ethicon</td><td>F682</td><td></td></tr><tr><td>Microrenathane .025&quot; O.D. x .012&quot; I.D.</td><td>Bioseb</td><td>MRE-025</td><td></td></tr><tr><td>olimel N7E</td><td>Baxter</td><td></td><td></td></tr><tr><td>PE10 - Polyethylene .011&quot; x .024&quot;&amp;nbsp;</td><td>Instech Solomon</td><td>BTPE-10</td><td></td></tr><tr><td>PE-50 tubing .023&quot; x.038&quot;</td><td>Instech Solomon</td><td>BTPE-50</td><td></td></tr><tr><td>Rodent Swivel 20 G</td><td>Bioseb</td><td>RS-20G&amp;nbsp;</td><td></td></tr><tr><td>Ropivaca&amp;iuml;ne</td><td>Astrazenica</td><td></td><td>Naropin 2&amp;nbsp;mg/mL</td></tr><tr><td>Xylazine</td><td>VMD</td><td></td><td>Xylazine hydrochloride 2%</td></tr></tbody></table>

use of a central venous line for fluids, drugs and nutrient administration in a mouse model of critical illness

This protocol describes a centrally catheterized mouse model of prolonged critical illness. We combine the cecal ligation and puncture method to induce sepsis with the use of a central venous line for fluids, drugs and nutrient administration to mimic the human clinical setting. Critically ill patients require intensive medical support in order to survive. While the majority of patients will recover within a few days, about a quarter of the patients need prolonged intensive care and are at high risk of dying from non-resolving multiple organ failure. Furthermore, the prolonged phase of critical illness is hallmarked by profound muscle weakness, and endocrine and metabolic changes, of which the pathogenesis is currently incompletely understood. The most widely used animal model in critical care research is the cecal ligation and puncture model to induce sepsis. This is a very reproducible model, with acute inflammatory and hemodynamic changes similar to human sepsis, which is designed to study the acute phase of critical illness. However, this model is hallmarked by a high lethality, which is different from the clinical human situation, and is not developed to study the prolonged phase of critical illness. Therefore, we adapted the technique by placing a central venous catheter in the jugular vein allowing us to administer clinically relevant supportive care, to better mimic the human clinical situation of critical illness. This mouse model requires an extensive surgical procedure and daily intensive care of the animals, but it results in a relevant model of the acute and prolonged phase of critical illness.

C57BL/6 mice were made critically ill as described above. We performed two experiments to assess post-CLP survival until two time points: survival until day 5 (n = 15) and survival until day 7 (n = 22) post-CLP. Survival curves of the two experiments were not significantly different (compared until day 5), indicating the reproducibility of the experimental setup. Non&#8211;surviving animals were found death or euthanized due to reaching human endpoints.&#160;&#160;Ligation of 50% of the c...

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Use of a Central Venous Line for Fluids, Drugs and Nutrient Administration in a Mouse Model of Critical Illness

This protocol describes a centrally catheterized mouse model of prolonged critical illness. We combine the cecal ligation and puncture method to induce sepsis with the use of a central venous line for fluids, drugs and nutrient administration to mimic the human clinical setting.

This protocol describes a centrally catheterized mouse model of prolonged critical illness. We combine the cecal ligation and puncture method to induce ...

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8.8K Views.  KU Leuven. This protocol describes a centrally catheterized mouse model of prolonged critical illness. We combine the cecal ligation and puncture method to induce sepsis with the use of a central venous line for fluids, drugs and nutrient administration to mimic the human clinical setting. 

Video: Use of a Central Venous Line for Fluids, Drugs and Nutrient Administration in a Mouse Model of Critical Illness

Hyperinsulinemic-euglycemic Clamps in Conscious, Unrestrained Mice

Type 2 diabetes is characterized by a defect in insulin action. The hyperinsulinemic-euglycemic clamp, or insulin clamp, is widely considered the "gold standard" method for assessing insulin action in vivo. During an insulin clamp, hyperinsulinemia is achieved by a constant insulin infusion. Euglycemia is maintained via a concomitant glucose infusion at a variable rate. This variable glucose infusion rate (GIR) is determined by measuring blood glucose at brief intervals throughout the experiment and adjusting the GIR accordingly. The GIR is indicative of whole-body insulin action, as mice with enhanced insulin action require a greater GIR. The insulin clamp can incorporate administration of isotopic 2[14C]deoxyglucose to assess tissue-specific glucose uptake and [3-3H]glucose to assess the ability of insulin to suppress the rate of endogenous glucose appearance (endoRa), a marker of hepatic glucose production, and to stimulate the rate of whole-body glucose disappearance (Rd).
The miniaturization of the insulin clamp for use in genetic mouse models of metabolic disease has led to significant advances in diabetes research. Methods for performing insulin clamps vary between laboratories. It is important to note that the manner in which an insulin clamp is performed can significantly affect the results obtained. We have published a comprehensive assessment of different approaches to performing insulin clamps in conscious mice1 as well as an evaluation of the metabolic response of four commonly used inbred mouse strains using various clamp techniques2. Here we present a protocol for performing insulin clamps on conscious, unrestrained mice developed by the Vanderbilt Mouse Metabolic Phenotyping Center (MMPC; URL: www.mc.vanderbilt.edu/mmpc). This includes a description of the method for implanting catheters used during the insulin clamp. The protocol employed by the Vanderbilt MMPC utilizes a unique two-catheter system3. One catheter is inserted into the jugular vein for infusions. A second catheter is inserted into the carotid artery, which allows for blood sampling without the need to restrain or handle the mouse. This technique provides a significant advantage to the most common method for obtaining blood samples during insulin clamps which is to sample from the severed tip of the tail. Unlike this latter method, sampling from an arterial catheter is not stressful to the mouse1. We also describe methods for using isotopic tracer infusions to assess tissue-specific insulin action. We also provide guidelines for the appropriate presentation of results obtained from insulin clamps.

The hyperinsulinemic-euglycemic clamp, or insulin clamp, is the gold standard for assessing insulin action in vivo. A method for performing insulin clamps in mice is described. This includes a method for arterial catheterization that permits experiments to be performed in conscious, unrestrained mice with minimal stress.

Type 2 diabetes is characterized by a defect in insulin action. The hyperinsulinemic-euglycemic clamp, or insulin clamp, is widely considered the "gold ...

Methods for Intravenous Self Administration in a Mouse Model

Animal models have been developed to study the reinforcing effects of drugs, including the intravenous self-administration (IVSA) paradigm. The advantages of using an IVSA paradigm to study the reinforcing properties of drugs of abuse such as cocaine include the fact that the drug is self-administered instead of experimenter-administered, the schedule of reinforcement can be altered, and accurate measurement of the quantities of drug consumed as well as the timing and pattern of IV injections can be obtained. Furthermore, the intravenous route of administration avoids potential confounds related to first pass metabolism or taste, and produces rapid increases in blood and brain drug levels. As outlined in this video, intravenous self-administration can be obtained without prior food restriction or prior drug training following careful catheter placement during surgery and meticulous daily catheter flushing and maintenance. Experimental procedures outlined in this paper include a description of animal housing and acclimation methods, operant training using sweetened milk solutions, and catheter implantation surgery.

The intravenous self-administration (IVSA) paradigm is considered to be the gold standard in examining the reinforcing properties of drugs of abuse in rodents. This manuscript outlines the experimental procedures and surgical techniques necessary to obtain reliable IVSA data. In particular, meticulous catheter implantation and maintenance are highlighted.

Animal models have been developed to study the reinforcing effects of drugs, including the intravenous self-administration (IVSA) paradigm. The advantages ...

Neuroscience

Catheterization of the Carotid Artery and Jugular Vein to Perform Hemodynamic Measures, Infusions and Blood Sampling in a Conscious Rat Model

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. Intra-tracheal inoculation of a known amount of bacteria has been successfully used to reproduce the pathogenesis of pneumonia which then develops into sepsis. Monitoring hemodynamic parameters and providing standard clinical treatment including infusion of antibiotics, fluids and drugs to maintain blood pressure is critical to simulate routine supportive care in this model but to do so requires both arterial and venous vascular access. The video details the surgical technique for implanting carotid artery and common jugular vein catheters in an anesthetized rat. Following a 72 hr recovery period, the animals will be re-anesthetized and connected to a tether and swivel setup attached to the rodent housing which connects the implanted catheters to the hemodynamic monitoring system. This setup allows free movement of the rat during the study while continuously monitoring pressures, infusing fluids and drugs (antibiotics, vasopressors) and performing blood sampling.

Vascular accesses to measure hemodynamics, provide fluids and perform blood sampling are important to any small animal model study. We present a technique for implanting catheters into the carotid artery and the common jugular vein in an anesthetized rat for connecting to a system to perform monitoring, infusions and sampling.

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. ...

Monitoring of Systemic and Hepatic Hemodynamic Parameters in Mice

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

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

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

Stenosis of the Inferior Vena Cava: A Murine Model of Deep Vein Thrombosis

Deep vein thrombosis (DVT) and its devastating complication, pulmonary embolism, are a severe health problem with high mortality. Mechanisms of thrombus formation in veins remain obscure. Lack of mobility (e.g., after surgery or long-haul flights) is one of the main factors leading to DVT. The pathophysiological consequence of the lack of mobility is blood flow stagnation in venous valves. Here, a model is described that mimics such flow disturbance as a thrombosis-driving factor. In this model, partial flow restriction (stenosis) in the inferior vena cava (IVC) is created. Closure of about 90% of the IVC lumen for 48 h results in development of thrombi structurally similar to those in humans. The similarities are: i) most of the thrombus volume is red, i.e., consists of red blood cells and fibrin, ii) presence of a white part (lines of Zahn), iii) non-denuded endothelial monolayer, iv) elevated plasma D-Dimer levels, and v) possibility to prevent thrombosis by low molecular weight heparin. Limitations include variable size of thrombi and the fact that a certain percentage of wild-type mice (0 - 35%) may not produce a thrombus. In addition to visual observation and measurement, thrombi may be visualized by non-invasive technologies, such as ultrasonography, which allows for monitoring the dynamics of thrombus development. At shorter time points (1 - 6 h), intravital microscopy may be applied to directly observe events (e.g., recruitment of cells to the vessel wall) preceding thrombus formation. Use of this method by several teams around the world has made it possible to uncover basic mechanisms of DVT initiation and identify potential targets that might be beneficial for its prevention.

We describe here stenosis in the inferior vena cava as a murine model of deep vein thrombosis. This model recapitulates blood flow restriction, one of the major triggers of venous thrombosis in humans.

Deep vein thrombosis (DVT) and its devastating complication, pulmonary embolism, are a severe health problem with high mortality. Mechanisms of thrombus ...

Immunology and Infection

Continuous IV Infusion is the Choice Treatment Route for Arginine-vasopressin Receptor Blocker Conivaptan in Mice to Study Stroke-evoked Brain Edema

Stroke is one of the major causes of morbidity and mortality in the world. Stroke is complicated by brain edema and other pathophysiological events. Among the most important players in the development and evolution of stroke-evoked brain edema is the hormone arginine-vasopressin and its receptors, V1a and V2. Recently, the V1a and V2 receptor blocker conivaptan has been attracting attention as a potential drug to reduce brain edema after stroke. However, animal models which involve conivaptan applications in stroke research need to be modified based on feasible routes of administration. Here the outcomes of 48 hr continuous intravenous (IV) are compared with intraperitoneal (IP) conivaptan treatments after experimental stroke in mice. We developed a protocol in which middle cerebral artery occlusion was combined with catheter installation into the jugular vein for IV treatment of conivaptan (0.2 mg) or vehicle. Different cohorts of animals were treated with 0.2 mg bolus of conivaptan or vehicle IP daily. Experimental stroke-evoked brain edema was evaluated in mice after continuous IV and IP treatments. Comparison of the results revealed that the continuous IV administration of conivaptan alleviates post-ischemic brain edema in mice, unlike the IP administration of conivaptan. We conclude that our model can be used for future studies of conivaptan applications in the context of stroke and brain edema.

Our studies have revealed that the beneficial effects of conivaptan are dependent on the method of delivery after experimental stroke in mice. We have developed a research protocol for delivery of the receptor blocker via IV catheter on stroke-evoked brain edema formation in mice.

Stroke is one of the major causes of morbidity and mortality in the world. Stroke is complicated by brain edema and other pathophysiological events. Among ...

Veno-Venous Extracorporeal Membrane Oxygenation in a Mouse

The use of extracorporeal membrane oxygenation (ECMO) has increased substantially in recent years. ECMO has become a reliable and effective therapy for acute as well as end-stage lung diseases. With the increase in clinical demand and prolonged use of ECMO, procedural optimization and prevention of multi-organ damage are of critical importance. The aim of this protocol is to present a detailed technique of veno-venous ECMO in a non-intubated, spontaneously breathing mouse. This protocol demonstrates the technical design of the ECMO and surgical steps. This murine ECMO model will facilitate the study of pathophysiology related to ECMO (e.g., inflammation,bleeding and thromboembolic events). Due to the abundance of genetically modified mice, the molecular mechanisms involved in ECMO-related complications can also be dissected.

Here we present a protocol describing the technique of veno-venous extracorporeal membrane oxygenation (ECMO) in a non-intubated, spontaneously breathing mouse. This murine model of ECMO can be effectively implemented in experimental studies of acute and end-stage lung diseases.

The use of extracorporeal membrane oxygenation (ECMO) has increased substantially in recent years. ECMO has become a reliable and effective therapy for ...

Murine Model of Central Venous Stenosis using Aortocaval Fistula with an Outflow Stenosis

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ligation of the inferior vena cava (IVC) in the outflow of the fistula, mimicking central venous stenosis. Technical aspects of this model are introduced. The aorta and IVC are exposed, following an abdominal incision. The infra-renal aorta and IVC are dissected for proximal clamping, and the distal aorta is exposed for puncture. The IVC at the midpoint between the left renal vein and the aortic bifurcation is carefully dissected to place an 8-0 suture beneath the IVC. After clamping the aorta and IVC, an AVF is created by puncturing the infra-renal aorta through both walls into the IVC with a 25 G needle, followed by ligating a 22 G intra-venous (IV) catheter and IVC together. The catheter is then removed, creating a reproducible venous stenosis without occlusion. The aorta and IVC are unclamped after confirming primary hemostasis. This novel model of central vein stenosis is easy to perform, reproducible, and will facilitate studies on AVF failure.

An aortocaval fistula was created by puncturing the murine infra-renal aorta through both walls into the inferior vena cava and was followed by creation of a stenosis in its outflow via partial ligation of the inferior vena cava. This reproducible model can be used to study central venous stenosis.

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ...

Drug Treatment by Central Venous Catheter in a Mouse Model of Angiotensin II Induced Abdominal Aortic Aneurysm and Monitoring by 3D Ultrasound

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse models, are applied to advance the understanding of the disease pathogenesis and to identify potential therapeutic targets. Testing novel drug candidates to block AAA growth in these models generally requires repeated drug administration during the time course of the experiment. Here, we describe a compiled protocol for AAA induction, insertion of an intravenous catheter to facilitate prolonged therapy, and serial AAA monitoring by 3D ultrasound. Aneurysms are induced in apolipoprotein E (ApoE) deficient mice by angiotensin II release over 28 days from osmotic mini-pumps implanted subcutaneously into the mouse back. Subsequently, the surgical procedure for external jugular vein catheterization is conducted to allow for daily intravenous drug treatment or repeated blood sampling via a subcutaneous vascular access button. Despite the two dorsal implants, the monitoring of AAA development is readily facilitated by sequential semi-automated 3D ultrasound analysis, which yields comprehensive information on the expansion of aortic diameter and volume and on aneurysm morphology, as illustrated by experimental examples.

This protocol describes the consecutive implantation of an osmotic pump to induce abdominal aortic aneurysm by angiotensin II release in apolipoprotein E (ApoE) deficient mice and of a vascular access port with a jugular vein catheter for repeated drug treatment. Monitoring of aneurysm development by 3D ultrasound is effectively conducted despite dorsal implants.

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse ...

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, ...