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>

<ol>
	<li>Angus, D. C., van der Poll, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Severe+sepsis+and+septic+shock.">Severe sepsis and septic shock.</a> N Engl J Med. 369 (9), 840-851 (2013).</li><li>Singer, 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=The+Third+International+Consensus+Definitions+for+Sepsis+and+Septic+Shock+(Sepsis-3).">The Third International Consensus Definitions for Sepsis and Septic Shock (Sepsis-3).</a> JAMA. 315 (8), 801-810 (2016).</li><li>Remick, D. G., Newcomb, D. E., Bolgos, G. L., Call, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+the+mortality+and+inflammatory+response+of+two+models+of+sepsis:+lipopolysaccharide+vs.+cecal+ligation+and+puncture.">Comparison of the mortality and inflammatory response of two models of sepsis: lipopolysaccharide vs. cecal ligation and puncture.</a> Shock. 13 (2), 110-116 (2000).</li><li>Copeland, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+inflammatory+response+to+endotoxin+in+mice+and+humans.">Acute inflammatory response to endotoxin in mice and humans.</a> Clin Diagn Lab Immunol. 12 (1), 60-67 (2005).</li><li>Seok, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genomic+responses+in+mouse+models+poorly+mimic+human+inflammatory+diseases.">Genomic responses in mouse models poorly mimic human inflammatory diseases.</a> Proc Natl Acad Sci U S A. 110 (9), 3507-3512 (2013).</li><li>Rittirsch, D., Huber-Lang, M. 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. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Abandon+the+mouse+research+ship?+Not+just+yet.">Abandon the mouse research ship? Not just yet.</a> Shock. 41 (6), 463-475 (2014).</li><li>Langford DJ1, B. A., Chanda, M. L., Clarke, S. E., Drummond, T. E., Echols, S., Glick, S., Ingrao, J., Klassen-Ross, T., Lacroix-Fralish, M. L., Matsumiya, L., Sorge, R. E., Sotocinal, S. G., Tabaka, J. M., Wong, D., van den Maagdenberg, A. M., Ferrari, M. D., Craig, K. D., Mogil, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coding+of+facial+expressions+of+pain+in+the+laboratory+mouse.">Coding of facial expressions of pain in the laboratory mouse.</a> Nat Methods. 7, (2010).</li><li>Hollenberg, S. 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=Characterization+of+a+hyperdynamic+murine+model+of+resuscitated+sepsis+using+echocardiography.">Characterization of a hyperdynamic murine model of resuscitated sepsis using echocardiography.</a> Am J Respir Crit Care Med. 164 (5), 891-895 (2001).</li><li>Heuer, J. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cecal+ligation+and+puncture+with+total+parenteral+nutrition:+a+clinically+relevant+model+of+the+metabolic,+hormonal,+and+inflammatory+dysfunction+associated+with+critical+illness.">Cecal ligation and puncture with total parenteral nutrition: a clinically relevant model of the metabolic, hormonal, and inflammatory dysfunction associated with critical illness.</a> J Surg Res. 121 (2), 178-186 (2004).</li><li>Conti, M., Merlani, P., Ricou, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prognosis+and+quality+of+life+of+elderly+patients+after+intensive+care.">Prognosis and quality of life of elderly patients after intensive care.</a> Swiss Med Wkly. 142, (2012).</li></ol>

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Buprenorphine</td>
			<td>Ecuphar</td>
			<td></td>
			<td>Vetergesic 0,3mg/ml</td>
		</tr>
		<tr>
			<td>C57BL/6&#160;</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&#160;</td>
			<td></td>
			<td>Tienam 500mg 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 100mg/ml</td>
		</tr>
		<tr>
			<td>LocTite Super glue3 all plastics&#160;</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 O.D.&#34; x .012 I.D.&#34;</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&#34; x .024&#34;&#160;</td>
			<td>Instech Solomon</td>
			<td>BTPE-10</td>
			<td></td>
		</tr>
		<tr>
			<td>PE-50 tubing .023&#34;x.038&#34;</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&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Ropivaca&#239;ne</td>
			<td>Astrazenica</td>
			<td></td>
			<td>Naropin 2mg/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

Manual Muscle Testing:  A Method of Measuring Extremity Muscle Strength Applied to Critically Ill Patients

Survivors of acute respiratory distress syndrome (ARDS) and other causes of critical illness often have generalized weakness, reduced exercise tolerance, and persistent nerve and muscle impairments after hospital discharge.1-6 Using an explicit protocol with a structured approach to training and quality assurance of research staff, manual muscle testing (MMT) is a highly reliable method for assessing strength, using a standardized clinical examination, for patients following ARDS, and can be completed with mechanically ventilated patients who can tolerate sitting upright in bed and are able to follow two-step commands. 7, 8 
This video demonstrates a protocol for MMT, which has been taught to &ge;43 research staff who have performed &gt;800 assessments on &gt;280 ARDS survivors. Modifications for the bedridden patient are included. Each muscle is tested with specific techniques for positioning, stabilization, resistance, and palpation for each score of the 6-point ordinal Medical Research Council scale.7,9-11 Three upper and three lower extremity muscles are graded in this protocol: shoulder abduction, elbow flexion, wrist extension, hip flexion, knee extension, and ankle dorsiflexion. These muscles were chosen based on the standard approach for evaluating patients for ICU-acquired weakness used in prior publications. 1,2.

Survivors of acute respiratory distress syndrome (ARDS) and critical illness frequently develop long-lasting muscle weakness. Manual muscle testing (MMT) is a standardized clinical examination commonly used to measure strength of peripheral skeletal muscle groups. This video demonstrates MMT using the 6-point Medical Research Council scale.

Survivors of acute respiratory distress syndrome (ARDS) and other causes of critical illness often have generalized weakness, reduced exercise tolerance, ...

Femoral Arterial and Venous Catheterization for Blood Sampling, Drug Administration and Conscious Blood Pressure and Heart Rate Measurements

In multiple fields of study, access to the circulatory system in laboratory studies is necessary. Pharmacological studies in rats using chronically implanted catheters permit a researcher to effectively and humanely administer substances, perform repeated blood sampling and assists in conscious direct measurements of blood pressure and heart rate. Once the catheter is implanted long-term sampling is possible. Patency and catheter life depends on multiple factors including the lock solution used, flushing regimen and catheter material. This video will demonstrate the methodology of femoral artery and venous catheterization of the rat. In addition the video will demonstrate the use of the femoral venous and arterial catheters for blood sampling, drug administration and use of the arterial catheter in taking measurements of blood pressure and heart rate in a conscious freely-moving rat. A tether and harness attached to a swivel system will allow the animal to be housed and have samples taken by the researcher with minimal disruption to the animal. To maintain patency of the catheter, careful daily maintenance of the catheter is required using lock solution (100 U/ml heparinized saline), machine-ground blunt tip syringe needles and the use of syringe filters to minimize potential contamination. With careful aseptic surgical techniques, proper catheter materials and careful catheter maintenance techniques, it is possible to sustain patent catheters and healthy animals for long periods of time (several weeks).

Chronic catheterization of blood vessels in the rat is often required for administration of substances, obtain blood sample over a period of time or for direct conscious blood pressure measurements. Femoral arterial catheterization of the rat and corresponding measurements of blood pressure in the conscious animal will be demonstrated.

In multiple fields of study, access to the circulatory system in laboratory studies is necessary. Pharmacological studies in rats using chronically ...

A Novel Surgical Approach for Intratracheal Administration of Bioactive Agents in a Fetal Mouse Model

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and acquired diseases. Apart from congenital or inherited abnormalities with the requirement for long-term expression of the delivered gene, several non-inherited perinatal conditions, where short-term gene expression or pharmacological intervention is sufficient to achieve therapeutic effects, are considered as potential future indications for this kind of approach. Candidate diseases for the application of short-term prenatal therapy could be the transient neonatal deficiency of surfactant protein B causing neonatal respiratory distress syndrome1,2 or hyperoxic injuries of the neonatal lung3. Candidate diseases for permanent therapeutic correction are Cystic Fibrosis (CF)4, genetic variants of surfactant deficiencies5 and &alpha;1-antitrypsin deficiency6.
Generally, an important advantage of prenatal gene therapy is the ability to start therapeutic intervention early in development, at or even prior to clinical manifestations in the patient, thus preventing irreparable damage to the individual. In addition, fetal organs have an increased cell proliferation rate as compared to adult organs, which could allow a more efficient gene or stem cell transfer into the fetus. Furthermore, in utero gene delivery is performed when the individual's immune system is not completely mature. Therefore, transplantation of heterologous cells or supplementation of a non-functional or absent protein with a correct version should not cause immune sensitization to the cell, vector or transgene product, which has recently been proven to be the case with both cellular and genetic therapies7.
In the present study, we investigated the potential to directly target the fetal trachea in a mouse model. This procedure is in use in larger animal models such as rabbits and sheep8, and even in a clinical setting9, but has to date not been performed before in a mouse model. When studying the potential of fetal gene therapy for genetic diseases such as CF, the mouse model is very useful as a first proof-of-concept because of the wide availability of different transgenic mouse strains, the well documented embryogenesis and fetal development, less stringent ethical regulations, short gestation and the large litter size.
Different access routes have been described to target the fetal rodent lung, including intra-amniotic injection10-12, (ultrasound-guided) intrapulmonary injection13,14 and intravenous administration into the yolk sac vessels15,16 or umbilical vein17. Our novel surgical procedure enables researchers to inject the agent of choice directly into the fetal mouse trachea which allows for a more efficient delivery to the airways than existing techniques18.

We developed a novel surgical approach for intratracheal administration of bioactive agents into the mouse fetus. The delivery route is more efficient in targeting the fetal mouse lungs than the commonly used intra-amniotic injection. This procedure has to date not been described in a mouse model.

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and ...

A Model of Chronic Nutrient Infusion in the Rat

Chronic exposure to excessive levels of nutrients is postulated to affect the function of several organs and tissues and to contribute to the development of the many complications associated with obesity and the metabolic syndrome, including type 2 diabetes. To study the mechanisms by which excessive levels of glucose and fatty acids affect the pancreatic beta-cell and the secretion of insulin, we have established a chronic nutrient infusion model in the rat. The procedure consists of catheterizing the right jugular vein and left carotid artery under general anesthesia; allowing a 7-day recuperation period; connecting the catheters to the pumps using a swivel and counterweight system that enables the animal to move freely in the cage; and infusing glucose and/or Intralipid (a soybean oil emulsion which generates a mixture of approximately 80% unsaturated/20% saturated fatty acids when infused with heparin) for 72 hr. This model offers several advantages, including the possibility to finely modulate the target levels of circulating glucose and fatty acids; the option to co-infuse pharmacological compounds; and the relatively short time frame as opposed to dietary models. It can be used to examine the mechanisms of nutrient-induced dysfunction in a variety of organs and to test the effectiveness of drugs in this context.

A protocol for chronic infusions of glucose and Intralipid in rats is described. This model can be used to study the impact of nutrient excess on organ function and physiological parameters.

Chronic exposure to excessive levels of nutrients is postulated to  affect the function of several organs and tissues and to contribute to the  ...

A Mouse Fetal Skin Model of Scarless Wound Repair

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs across species but is age dependent. The transition from a scarless to scarring phenotype occurs in the third trimester of pregnancy in humans and around embryonic day 18 (E18) in mice. However, this varies with the size of the wound with larger defects generating a scar at an earlier gestational age. The emergence of lineage tracing and other genetic tools in the mouse has opened promising new avenues for investigation of fetal scarless wound healing. However, given the inherently high rates of morbidity and premature uterine contraction associated with fetal surgery, investigations of fetal scarless wound healing in vivo require a precise and reproducible surgical model. Here we detail a reliable model of fetal scarless wound healing in the dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

During mammalian development, early gestational skin wounds heal without a scar. Here we detail a reliable and reproducible model of fetal scarless wound healing in the cutaneous dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs ...

A Simple Critical-sized Femoral Defect Model in Mice

While bone has a remarkable capacity for regeneration, serious bone trauma often results in damage that does not properly heal. In fact, one tenth of all limb bone fractures fail to heal completely due to the extent of the trauma, disease, or age of the patient. Our ability to improve bone regenerative strategies is critically dependent on the ability to mimic serious bone trauma in test animals, but the generation and stabilization of large bone lesions is technically challenging. In most cases, serious long bone trauma is mimicked experimentally by establishing a defect that will not naturally heal. This is achieved by complete removal of a bone segment that is larger than 1.5 times the diameter of the bone cross-section. The bone is then stabilized with a metal implant to maintain proper orientation of the fracture edges and allow for mobility. 
Due to their small size and the fragility of their long bones, establishment of such lesions in mice are beyond the capabilities of most research groups. As such, long bone defect models are confined to rats and larger animals. Nevertheless, mice afford significant research advantages in that they can be genetically modified and bred as immune-compromised strains that do not reject human cells and tissue.
Herein, we demonstrate a technique that facilitates the generation of a segmental defect in mouse femora using standard laboratory and veterinary equipment. With practice, fabrication of the fixation device and surgical implantation is feasible for the majority of trained veterinarians and animal research personnel. Using example data, we also provide methodologies for the quantitative analysis of bone healing for the model.

Animal models are frequently employed to mimic serious bone injury in biomedical research. Due to their small size, establishment of stabilized bone lesions in mice are beyond the capabilities of most research groups. Herein, we describe a simple method for establishing and analyzing experimental femoral defects in mice.

While bone has a remarkable capacity for regeneration, serious bone trauma often results in damage that does not properly heal. In fact, one tenth of all ...

A Novel Approach for the Administration of Medications and Fluids in Emergency Scenarios and Settings

The available routes of administration commonly used for medications and fluids in the acute care setting are generally limited to oral, intravenous, or intraosseous routes, but in many patients, particularly in the emergency or critical care settings, these routes are often unavailable or time-consuming to access. A novel device is now available that offers an easy route for administration of medications or fluids via rectal mucosal absorption (also referred to as proctoclysis in the case of fluid administration and subsequent absorption). Although originally intended for the palliative care market, the utility of this device in the emergency setting has recently been described. Specifically, reports of patients being treated for dehydration, alcohol withdrawal, vomiting, fever, myocardial infarction, hyperthyroidism, and cardiac arrest have shown success with administration of a wide variety of medications or fluids (including water, aspirin, lorazepam, ondansetron, acetaminophen, methimazole, and buspirone). Device placement is straightforward, and based on the observation of expected effects from the medication administrations, absorption is rapid. The rapidity of absorption kinetics are further demonstrated in a recent report of the measurement of phenobarbital pharmacokinetics. We describe here the placement and use of this device, and demonstrate methods of pharmacokinetic measurements of medications administered by this method.

This study presents a novel device that offers an easy route to quickly provide medications and fluids in patients with limited or difficult IV access. This small-diameter device is placed in the distal third of the rectum, allowing for ongoing medication and fluids administration.

The available routes of administration commonly used for medications and fluids in the acute care setting are generally limited to oral, intravenous, or ...

Use of a Piglet Model for the Study of Anesthetic-induced Developmental Neurotoxicity (AIDN): A Translational Neuroscience Approach

Anesthesia cannot be avoided in many cases when surgery is required, particularly in children. Recent investigations in animals have raised concerns that anesthesia exposure may lead to neuronal apoptosis, known as anesthesia-induced developmental neurotoxicity (AIDN). Furthermore, some clinical studies in children have suggested that anesthesia exposure may lead to neurodevelopmental deficits later in life. Nonetheless, an ideal animal model for preclinical study has yet to be developed. The neonatal piglet represents a valuable model for preclinical study, as they share a striking number of developmental similarities with humans.
The anatomy and physiology of piglets allow for implementation of rigorous human perioperative conditions in both survival and non-survival procedures. Femoral artery catheterization allows for close monitoring, thus enabling prompt correction of any deviation of the piglet's vital signs and chemistries. In addition, there are multiple developmental similarities between piglets and human neonates. The techniques required to use piglets for experimentation will require experience to master. A pediatric anesthesiologist is a critical member of the investigative team. We describe, in a general sense, the appropriate use of a piglet model for neurodevelopmental study.

Use of a Piglet Model for the Study of Anesthetic-induced Developmental Neurotoxicity AIDN: A Translational Neuroscience Approach

Anesthesia-induced developmental neurotoxicity (AIDN) research has focused on rodents, which are not broadly applicable to humans. Non-human primate models are more relevant, but are cost-prohibitive and difficult to use for experimentation. The piglet, in contrast, is a clinically relevant, practical animal model ideal for the study of anesthetic neurotoxicity.

Anesthesia cannot be avoided in many cases when surgery is required, particularly in children.  Recent investigations in animals have raised concerns that ...

Dissection of the Mouse Pancreas for Histological Analysis and Metabolic Profiling

We have been investigating the pancreas specific transcription factor, 1a cre-recombinase; lox-stop-lox- Kristen rat sarcoma, glycine to aspartic acid at the 12 codon (Ptf1acre/+;LSL-KrasG12D/+) mouse strain as a model of human pancreatic cancer. The goal of our current studies is to identify novel metabolic biomarkers of pancreatic cancer progression. We have performed metabolic profiling of urine, feces, blood, and pancreas tissue extracts, as well as histological analyses of the pancreas to stage the cancer progression. The mouse pancreas is not a well-defined solid organ like in humans, but rather is a diffusely distributed soft tissue that is not easily identified by individuals unfamiliar with mouse internal anatomy or by individuals that have little or no experience performing mouse organ dissections. The purpose of this article is to provide a detailed step-wise visual demonstration to guide novices in the removal of the mouse pancreas by dissection. This article should be especially valuable to students and investigators new to research that requires harvesting of the mouse pancreas by dissection for metabolic profiling or histological analyses.

This video article provides a detailed demonstration of the procedures required to successfully remove the pancreas from a mouse by dissection for histological analysis and metabolic profiling.

We have been investigating the pancreas specific transcription factor, 1a cre-recombinase; lox-stop-lox- Kristen rat sarcoma, glycine to aspartic acid at ...

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 ...