Name: establishing in situ closed circuit perfusion of lower abdominal organs and hind limbs in mice
Uploaded: 2020-08-13T12:30:00.000+00:00
Duration: 5 min 27 s
Description: Watch this Scientific Journal Video about Establishing In Situ Closed Circuit Perfusion of Lower Abdominal Organs and Hind Limbs in Mice at JoVE.com

The study was supported by the NIH grant CA194058 to DS, Skaggs School of Pharmacy ADR seed grant program (DS); National Natural Science Foundation of China (Grant No. 31771093), the Project of International Collaboration of Jilin Province (No.201180414085GH), the Fundamental&#8194;Research&#8194;Funds&#8194;for&#8194;the&#8194;Central&#8194;Universities, the Program for JLU Science and Technology Innovative Research Team (2017TD-27, 2019TD-36).

All methods described here have been approved by the University of Colorado&#8217;s Institutional Animal Care and Use Committee (IACUC).
1. Pre-heat the perfusion system
<ol>
	<li>Prepare the perfusion system before surgery by starting a 37 &#176;C circulating water bath for all water-jacketed components (perfusate reservoir, moist chamber, and lid) as shown in a customized configuration in Figure 1A. Make sure the tubing is clean and replace if necessary. To limit perfusate volume, use a bubble trap within a moist chamber as the perfusate reservoir (Figure 1B-6).</li>
</ol>
2. Vascular catheterization
<ol>
	<li>Induce anesthesia in an 8-10 week old BALB/c mouse using an isoflurane veterinary anesthesia machine with 3-5% isoflurane and oxygen flow rate at 0.3 L/min. As an alternative, use ketamine/xylazine or any other type of intraperitoneal anesthesia. Evaluate the depth of anesthesia by 2 methods: toe-pinch and corneal reflex.</li>
	<li>Prewet a 4-0 silk suture with needle in double distilled water.</li>
	<li>Place the anesthetized mouse in a supine position on a Styrofoam board with the head facing the surgeon and immobilize forelimbs and hind limbs with tape. Wipe the abdomen with isopropyl alcohol and cut the abdomen along the midline in a &#34;T&#34; shape with scissors. Stop bleeding around the edge of the incision by electrocoagulation (cauterizing).</li>
	<li>Push the stomach, jejunum and colon to the right side of the abdomen to reveal the abdominal aorta, vena cava, and common iliac and iliolumbar arteries and veins.</li>
	<li>Under a dissection microscope, find and ligate the iliolumbar artery/vein in the male, and ovarian artery/vein and the iliolumbar artery/vein in the female using 4-0 silk sutures (Figure 2 yellow lines).</li>
	<li>Under a dissection microscope, loop two 4-0 silk sutures underneath the abdominal aorta and inferior vena cava (about 1 cm above iliac artery and vein, 1 mm apart, Figure 2), and make a loose knot in the suture closest to the iliac vessels (Figure 2, white dotted line). Alternatively, a 6-0 silk suture can be used for this knot.</li>
	<li>Under a dissection microscope, horizontally align and stretch both the inferior vena cava and the abdominal aorta with porte-aiguille. Use a 24 G winged shielded I.V. catheter to puncture the abdominal aorta, push the button to retract the needle core and insert the catheter about 5 mm into the vessel.
	<ol>
		<li>Repeat the same procedure with the inferior vena cava and tie up knots of both sutures around the catheterized vessels. 
		NOTE: The needle can easily puncture through the blood vessel; therefore, keep the vessels stretched and needle parallel with the vessel. Retract the needle core as soon as the needle penetrates about 1 mm into the vessel. The abdominal aorta is underneath the inferior vena cava and much thinner and more elastic due to being encased in connective tissue. Therefore, the aorta can &#8220;hold on&#8221; to the catheter, and should thus be catheterized before the vena cava to reduce the likelihood of the catheter slipping out.</li>
	</ol>
	</li>
	<li>Apply instant glue to immobilize the catheters to the erector spinae, replace the abdominal organs, and end the surgery while maintaining anesthesia. 
	NOTE: Organs that cannot be completely replaced into the abdominal cavity will need to be periodically moistened with perfusion medium during the perfusion process.</li>
</ol>
3. Set up the perfusion system
<ol>
	<li>Transfer the mouse into the water-jacketed moist chamber prewarmed to 37 &#176;C on a silicon pad and immobilize the catheter wings to the pad with 19 G needles.</li>
	<li>Fill the arterial catheter&#8217;s end (inlet) with prewarmed perfusion buffer (Ringer&#8217;s lactate solution supplemented with 5% BSA), and then connect the catheter end with the inlet perfusion tubing using a screw-on connector (Figure 1B, red arrow). 
	NOTE: Hold the connector with hemostatic forceps and immobilize the tubing with tape to avoid moving the catheters.</li>
	<li>Adjust the peristaltic flow rate to 0.6 mL/min and keep the perfusion outflow (Figure 1B, blue arrow) open-ended for 5-10 min to wash out the blood through the venous catheter. There will be some clots in the outlet catheter; flush out the clots with perfusate buffer before closing the perfusion circuit.</li>
	<li>Connect the venous catheter&#8217;s end with the outlet tubing using a screw-on connector to close the circuit (Figure 1B). At this point, perform CO2&#160;gas euthanasia and verify by chest puncture or any other method.</li>
	<li>Cover the moist chamber with the warmed lid. Check the level of perfusate periodically and add more if needed. Perfusion can be performed for up to 2 hours. 
	NOTE: 5 mL of perfusate will be needed to set up the closed perfusion system. If there is no leaking or edema, the volume of perfusate will decrease by less than 1 mL and additional buffer will not be needed. To avoid edema induced by peripheral circulating thrombus, perfusion buffer containing 0.002% heparin can be used in the first 10 minutes of perfusion, but should be changed to buffer without heparin to avoid the leaking at the edge of the incisions.</li>
	<li>If needed, add a reagent of choice into the perfusion reservoir or to the injection port at any time (Figure 1B-5). For example, 10 &#181;L of 10 mg/mL Hoechst33342 can be added into the perfusate to stain the cell nuclei 2 h before the end of perfusion, or 50 &#181;L of 1 mg/mL DyLight 649-lectin to stain the vascular endothelial cells 30 min before the end of perfusion. 
	NOTE: If attempting to stain bone marrow with Hoechst solution, mice will need to be pre-injected 30 minutes before surgery.</li>
	<li>After perfusion with fluorescent reagents, wash out with perfusion buffer for another 10 minutes to minimize the background fluorescence.</li>
</ol>
4. Analysis of perfused organs
<ol>
	<li>Collect organs including testis, prostate, bladder, femur, muscle, and skin (e.g., feet). Excise a piece of organ about 1 mm3 and flatten between two glass slides.
	<ol>
		<li>Study under inverted fluorescent confocal microscope using DAPI/Cy5 excitation and emission filters (excitation lasers : DAPI, 405 nm; Cy5,640 nm). Use at least 200x magnification objective with a 0.45 numerical aperture.</li>
		<li>Alternatively, fix the organs with 4% formaldehyde solution for 24 h and perform hematoxylin-eosin staining19.</li>
	</ol>
	</li>
	<li>To create a bone window for intact bone marrow observation, immobilize both ends of the femur or tibia and scrape away the cortical bone with the lateral edge of a 19 G needle to expose the periosteum; take care to keep a thin layer of residual bone. Place bone on a cover slip with the window facing the glass and image with inverted fluorescent confocal microscope using DAPI and Cy5 channels as described above. The cells and vascular network in the bone marrow cavity can be readily observed.</li>
</ol>

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The authors have nothing to disclose.

The described circuit can be used to probe various research questions, for example the role of different serum components and tissue barriers in drug delivery, or immune and stem cell trafficking. Different drug delivery systems (e.g., liposomes and nanoparticles) can be added to the perfusate in order to understand the role of physiological and biochemical factors in delivery. The duration of perfusion can vary, depending on the tissue studied, scientific goals, and the composition of perfusate. We present here the resu...

Isolated organ perfusion was originally developed to study organ physiology for transplantation1,2,3, and enabled understanding of functions of the organs without interference from other body systems. For example, isolated kidney and heart perfusion was immensely useful in understanding basic principles of hemodynamics and effects of vasoactive agents, whereas liver perfusion was important to understanding the metabolic function, including drug metabolism in healthy and diseased tissue4,5,6,7. In addition, perfusion studies were critical in understanding viability and function of organs intended for transplantation. In Cancer Researchearch, isolated tumor perfusion has been described by several groups using mouse, rat, and freshly resected human tissues8,9. In some isolated tumor perfusion, the tumor was implanted in the ovary fat pad to force the growth of tumor supplying blood vessels from the mesentery artery10. The Jain group performed pioneering studies using isolated perfusion of colon adenocarcinomas to understand tumor hemodynamics and metastasis8,11,12,13. Other innovative engineered ex vivo setups include a 96-well plate-based perfusion device to culture the primary human multiple myeloma cells14 and a modular flow chamber for engineering bone marrow architecture and function research15.
In addition to physiology and pathology studies, organ perfusion has been used to study the basic principles of drug delivery. Thus, one group described isolated rat limb perfusion and studied accumulation of liposomes in implanted sarcomas16, whereas another group performed dissected human kidney perfusion to study the endothelial targeting of nanoparticles17. Ternullo et al. used an isolated perfused human skin flap as a close-to-in vivo skin drug penetration model18.
Despite these advancements in perfusion of large organs and tissues, there have been no reports on in situ perfusion models in mice that: a) bypass clearance organs such as liver, spleen and kidneys; b) include pelvic organs, skin, muscle, reproductive organs (in male), bladder, prostate and bone marrow. Due to the small size of these organs and the supplying vasculature, ex vivo cannulation and establishment of a perfusion circuit has not been feasible. The mouse is the most important animal model in cancer and immunology research, and drug delivery. The ability to perfuse small mouse organs would allow interesting questions regarding drug delivery to these organs, including to tumors implanted in the pelvis (bladder, prostate, ovary, bone marrow), to be answered, as well as studies of basic physiology and immunology of diseases of these organs. To address this deficiency, we developed an in situ perfusion circuit in mice that can potentially avoid tissue injury and is much better suited for functional research than isolated organ perfusion.

<table><tbody><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>3.5x-90x stereo zoom microscope on boom stand with LED light</td><td>Amscope</td><td>SKU: SM-3BZ-80S</td><td></td></tr><tr><td>Carbon dioxide, USP</td><td>Airgas healthcare</td><td>19087-5283</td><td></td></tr><tr><td>Confocal microscope</td><td>NIKON</td><td>ECLIPSE Ti2</td><td></td></tr><tr><td>Disposable Sterile Cautery Pen with High Temp</td><td>FIAB</td><td>F7244</td><td></td></tr><tr><td>Moist chamber bubble trap (part 6 in Figure 1)</td><td>Harvard Apparatus</td><td>733692</td><td>Customized as the perfisate container; also enabled constant pressure perfusion</td></tr><tr><td>Moist chamber cover with quartz window (part 3 in Figure 1)</td><td>Harvard Apparatus</td><td>733524</td><td>keep the chamber&#039;s temperature</td></tr><tr><td>Moist chamber with metal tube heat exchanger</td><td>Harvard Apparatus</td><td>732901</td><td>Water-jacketed moist chamber with lid to maintain perfusate and mouse temperature</td></tr><tr><td>Olsen-Hegar needle holders with suture cutters</td><td>Fine Science Tools (FST)</td><td>125014</td><td></td></tr><tr><td>Oxygen compressed, USP</td><td>Airgas healthcare</td><td>C2649150AE06</td><td></td></tr><tr><td>Roller pump (part 4 in Figure 1)</td><td>Harvard Apparatus</td><td>730113</td><td>deliver perfusate to cannula in the moist chamber</td></tr><tr><td>SCP plugsys servo control F/Perfusion (part 1 in Figure 1)</td><td>Harvard Apparatus</td><td>732806</td><td>control the purfusion speed</td></tr><tr><td>Silicone pad</td><td>Harvard Apparatus</td><td></td><td></td></tr><tr><td>Silicone tubing set (arrows in Figure 1)</td><td>Harvard Apparatus (TYGON)</td><td>733456</td><td></td></tr><tr><td>Student standard pattern forceps</td><td>Fine Science Tools (FST)</td><td>91100-12</td><td></td></tr><tr><td>Surgical Scissors</td><td>Fine Science Tools (FST)</td><td>14001-14</td><td></td></tr><tr><td>Table for moist chamber</td><td>Harvard Apparatus</td><td>734198</td><td></td></tr><tr><td>Thermocirculator (part 2 in Figure 1)</td><td>Harvard Apparatus</td><td>724927</td><td>circulating water bath for all water-jacketed components</td></tr><tr><td>Three-way stopcock (part 5 in Figure 1)</td><td>Cole-Palmer</td><td>30600-02</td><td></td></tr><tr><td>Veterinary anesthesia machine</td><td>Highland</td><td>HME109</td><td></td></tr><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>19-G BD PrecisionGlide needle</td><td>BD</td><td>305186</td><td>For immobilizing the Insyte Autoguard Winged needle and scratching the cortical bone</td></tr><tr><td>4-0 silk sutures</td><td>Keebomed-Hopemedical</td><td>427411</td><td></td></tr><tr><td>6-0 silk sutures</td><td>Keebomed-Hopemedical</td><td>427401</td><td></td></tr><tr><td>Filter (0.2 &amp;micro;m)</td><td>ThermoFisher</td><td>42225-CA</td><td>Filter for 5% BSA-RINGER&amp;rsquo;S</td></tr><tr><td>Permanent marker</td><td>Staedtler</td><td>342-9</td><td></td></tr><tr><td>Syringe (10 mL)</td><td>Fisher Scientific</td><td>14-823-2E</td><td></td></tr><tr><td>Syringe (60 mL)</td><td>BD</td><td>309653</td><td>Filter for 5% BSA-RINGER&amp;rsquo;S</td></tr><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1% Evans blue ( w/v ) in phosphate-buffered saline (PBS, pH 7.5)</td><td>Sigma</td><td>314-13-6</td><td></td></tr><tr><td>10% buffered formalin</td><td>velleyvet</td><td>36692</td><td></td></tr><tr><td>BALB/c mice ( 8-10 weeks old )</td><td>Charles River</td><td></td><td></td></tr><tr><td>Baxter Viaflex lactate Ringer&#039;s solution</td><td>EMRN Medical Supplies Inc.</td><td>JB2324</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Thermo Fisher</td><td>11021-037</td><td></td></tr><tr><td>Cyanoacrylate glue</td><td>Krazy Glue</td><td></td><td></td></tr><tr><td>DyLight-649-lectin</td><td>Vector Laboratories,Inc.</td><td>ZB1214</td><td></td></tr><tr><td>Ethanol (70% (vol/vol))</td><td>Pharmco</td><td>111000190</td><td></td></tr><tr><td>Hoechst33342</td><td>Life Technologies</td><td>H3570</td><td></td></tr><tr><td>Isoflurane</td><td>Piramal Enterprises Limited</td><td>66794-017-25</td><td></td></tr><tr><td>Phosphate buffered saline</td><td>Gibco</td><td>10010023</td><td></td></tr></tbody></table>

establishing in situ closed circuit perfusion of lower abdominal organs and hind limbs in mice

Ex vivo perfusion is an important physiological tool to study the function of isolated organs (e.g. liver, kidneys). At the same time, due to the small size of mouse organs, ex vivo perfusion of bone, bladder, skin, prostate, and reproductive organs is challenging or not feasible. Here, we report for the first time an in situ lower body perfusion circuit in mice that includes the above tissues, but bypasses the main clearance organs (kidney, liver, and spleen). The circuit is established by cannulating the abdominal aorta and inferior vena cava above the iliac artery and vein and cauterizing peripheral blood vessels. Perfusion is performed via a peristaltic pump with perfusate flow maintained for up to 2 h. In situ staining with fluorescent lectin and Hoechst solution confirmed that the microvasculature was successfully perfused. This mouse model can be a very useful tool for studying pathological processes as well as mechanisms of drug delivery, migration/metastasis of circulating tumor cells into/from the tumor, and interactions of immune system with perfused organs and tissues.

We set up a closed circuit perfusion system through cannulation of the abdominal aorta and the inferior vena cava of 8-10 week old mice while keeping the volume of perfusion buffer less than 10 mL. Figure 3A shows confocal images after perfusing tissues with Ringer&#8217;s solution containing Hoechst 33342 and DyLight 649-lectin. Muscle, bone marrow, testis, bladder, prostate, and foot skin show efficient nuclear and vascular staining. Figure 3B shows hematoxyli...

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Establishing In Situ Closed Circuit Perfusion of Lower Abdominal Organs and Hind Limbs in Mice

A protocol is described for in situ perfusion of the mouse lower body, including the bladder, the prostate, sex organs, bone, muscle and foot skin.

Ex vivo perfusion is an important physiological tool to study the function of isolated organs (e.g. liver, kidneys). At the same time, due to the small ...

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6.0K Views.  The First Hospital of Jilin University. A protocol is described for in situ perfusion of the mouse lower body, including the bladder, the prostate, sex organs, bone, muscle and foot skin.

Video: Establishing In Situ Closed Circuit Perfusion of Lower Abdominal Organs and Hind Limbs in Mice

Programming Stem Cells for Therapeutic Angiogenesis Using Biodegradable Polymeric Nanoparticles

Controlled vascular growth is critical for successful tissue regeneration and wound healing, as well as for treating ischemic diseases such as stroke, heart attack or peripheral arterial diseases. Direct delivery of angiogenic growth factors has the potential to stimulate new blood vessel growth, but is often associated with limitations such as lack of targeting and short half-life in vivo. Gene therapy offers an alternative approach by delivering genes encoding angiogenic factors, but often requires using virus, and is limited by safety concerns. Here we describe a recently developed strategy for stimulating vascular growth by programming stem cells to overexpress angiogenic factors in situ using biodegradable polymeric nanoparticles. Specifically our strategy utilized stem cells as delivery vehicles by taking advantage of their ability to migrate toward ischemic tissues in vivo. Using the optimized polymeric vectors, adipose-derived stem cells were modified to overexpress an angiogenic gene encoding vascular endothelial growth factor (VEGF). We described the processes for polymer synthesis, nanoparticle formation, transfecting stem cells in vitro, as well as methods for validating the efficacy of VEGF-expressing stem cells for promoting angiogenesis in a murine hindlimb ischemia model.

We describe the method of programming stem cells to overexpress therapeutic factors for angiogenesis using biodegradable polymeric nanoparticles. Processes described include polymer synthesis, transfecting adipose-derived stem cells in vitro, and validating the efficacy of programmed stem cells to promote angiogenesis in a murine hindlimb ischemia model.

Controlled vascular growth is critical for successful tissue regeneration and wound healing, as well as for treating ischemic diseases such as stroke, ...

Bioengineering

Murine Model of Metastatic Liver Tumors in the Setting of Ischemia Reperfusion Injury

Liver ischemia and reperfusion (I/R) injury, a common clinical challenge, remains an inevitable pathophysiological process that has been shown to induce multiple tissue and organ damage. Despite recent advances and therapeutic approaches, the overall morbidity has remained unsatisfactory especially in patients with underlying parenchymal abnormalities. In the context of aggressive cancer growth and metastasis, surgical I/R is suspected to be the promoter regulating tumor recurrence. This article aims to describe a clinically relevant murine model of liver I/R and colorectal liver metastasis. In doing so, we aim to assist other investigators in establishing and perfecting this model for their routine research practice to better understand the effects of liver I/R on promoting liver metastases.

We describe in detail a clinically relevant colorectal cancer liver metastases (CRLM) tumor model and the influence of liver ischemia reperfusion (I/R) in tumor growth and metastasis. This model can help to better understand the mechanisms underlying surgery-induced promotion of liver metastatic growth.

Liver ischemia and reperfusion (I/R) injury, a common clinical challenge, remains an inevitable pathophysiological process that has been shown to induce ...

Cancer Research

Murine Model of Hindlimb Ischemia 

In the United States, peripheral arterial disease (PAD) affects about 10 million individuals, and is also prevalent worldwide. Medical therapies for symptomatic relief are limited. Surgical or endovascular interventions are useful for some individuals, but long-term results are often disappointing. As a result, there is a need for developing new therapies to treat PAD. The murine hindlimb ischemia preparation is a model of PAD, and is useful for testing new therapies. When compared to other models of tissue ischemia such as coronary or cerebral artery ligation, femoral artery ligation provides for a simpler model of ischemic tissue. Other advantages of this model are the ease of access to the femoral artery and low mortality rate. 
In this video, we demonstrate the methodology for the murine model of unilateral hindimb ischemia. The specific materials and procedures for creating and evaluating the model will be described, including the assessment of limb perfusion by laser Doppler imaging. This protocol can also be utilized for the transplantation and non-invasive tracking of cells, which is demonstrated by Huang et al.1.

Murine Model of Hindlimb Ischemia

The surgical procedure for induction of unilateral hindlimb ischemia is demonstrated, with confirmation of ischemia by laser Doppler perfusion imaging.

In the United States, peripheral arterial disease (PAD) affects about 10 million individuals, and is also prevalent worldwide.  Medical therapies for ...

Biology

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming increasingly common. Ex vivo liver perfusions allow for a comprehensive analysis of liver metabolism using nuclear magnetic resonance (NMR), in nutritional conditions that can be rigorously controlled. As in silico simulations remain a primarily theoretical means of assessing hormone actions and the effects of pharmaceutical intervention, the perfused liver remains one of the most valuable test beds for understanding hepatic metabolism. As these studies guide basic insights into hepatic physiology, results must be accurate and reproducible. The greatest factor in the reproducibility of ex vivo hepatic perfusion is the quality of surgery. Therefore, we have introduced an organized and streamlined method to perform ex vivo mouse liver perfusions in the context of in situ NMR experiments. We also describe a unique application and discuss common issues encountered in these studies. The overall purpose is to provide an uncomplicated guide to a technique we have refined over several years that we deem the golden standard for obtaining reproducible results in hepatic resections and perfusions in the context of in situ NMR experiments. The distance to the center of the field for the magnet as well as the inaccessibility of the tissue to intervention during the NMR experiment makes our methods novel.

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

The protocol describes a straightforward method of resectioning an intact mouse liver for metabolic studies through portal vein perfusion.

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming ...

Biochemistry

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing modified cannulas and techniques adapted from conventional commercial ex vivo perfusion equipment. The system was utilized to evaluate the preservation outcomes following 12 h of perfusion. C57BL/6J mice served as liver donors, and the livers were explanted by cannulating the portal vein (PV) and bile duct (BD), and subsequently flushing the organ with warm (37 &#176;C) heparinized saline. Then, the explanted livers were transferred to the perfusion chamber and subjected to normothermic oxygenated machine perfusion (NEVLP). Inlet and outlet perfusate samples were collected at 3 h intervals for perfusate analysis. Upon completion of the perfusion, liver samples were obtained for histological analysis, with morphological integrity assessed using modified Suzuki-Score through Hematoxylin-Eosin (HE) staining. The optimization experiments yielded the following findings: (1) mice weighing over 30 g were deemed more suitable for the experiment due to the larger size of their bile duct (BD). (2) a 2 Fr (outer diameter = 0.66 mm) polyurethane cannula was better suited for cannulating the portal vein (PV) when compared to a polypropylene cannula. This was attributed to the polyurethane material&#39;s enhanced grip, resulting in reduced catheter slippage during the transfer from the body to the organ chamber. (3) for cannulation of the bile duct (BD), a 1 Fr (outer diameter = 0.33 mm) polyurethane cannula was found to be more effective compared to the polypropylene UT - 03 (outer diameter = 0.30 mm) cannula. With this optimized protocol, mouse livers were successfully preserved for a duration of 12 h without significant impact on the histological structure. Hematoxylin-Eosin (HE) staining revealed a well-preserved morphological architecture of the liver, characterized by predominantly viable hepatocytes with clearly visible nuclei and mild dilation of hepatic sinusoids.

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

A normothermic ex vivo liver perfusion (NEVLP) system was created for mouse livers. This system requires experience in microsurgery but allows for reproducible perfusion results. The ability to utilize mouse livers facilitates the investigation of molecular pathways to identify novel perfusate additives and enables the execution of experiments focused on organ repair.

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing ...

Engineering

Methods for Acute and Subacute Murine Hindlimb Ischemia

Peripheral artery disease (PAD) is a leading cause of cardiovascular morbidity and mortality in developed countries, and animal models that reliably reproduce the human disease are necessary to develop new therapies for this disease. The mouse hindlimb ischemia model has been widely used for this purpose, but the standard practice of inducing acute limb ischemia by ligation of the femoral artery can result in substantial tissue necrosis, compromising investigators&#39; ability to study the vascular and skeletal muscle tissue responses to ischemia. An alternative approach to femoral artery ligation is the induction of gradual femoral artery occlusion through the use of ameroid constrictors. When placed around the femoral artery in the same or different locations as the sites of femoral artery ligation, these devices occlude the artery over 1 - 3 days, resulting in more gradual, subacute ischemia. This results in less substantial skeletal muscle tissue necrosis, which may more closely mimic the responses seen in human PAD. Because genetic background influences outcomes in both the acute and subacute ischemia models, consideration of the mouse strain being studied is important in choosing the best model. This paper describes the proper procedure and anatomical placement of ligatures or ameroid constrictors on the mouse femoral artery to induce subacute or acute hindlimb ischemia in the mouse.

Surgical induction of hindlimb ischemia in the mouse is useful to examine angiogenesis, however this is compromised in certain inbred mouse strains that display marked ischemia-induced tissue necrosis. Methods are described to induce subacute limb ischemia using ameroid constrictors to circumvent this problem through the induction of gradual arterial occlusion.

Peripheral artery disease (PAD) is a leading cause of cardiovascular morbidity and mortality in developed countries, and animal models that reliably ...

Laser Doppler Perfusion Imaging in the Mouse Hindlimb

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is a common, noninvasive, repeatable method for assessing blood flow recovery. The technique calculates overall blood flow in the sampled tissue from the Doppler shift in frequency caused when a laser hits moving red blood cells. Measurements are expressed in arbitrary perfusion units, so the contralateral non-intervened upon leg is usually used to help control measurements. Measurement depth is in the range of 0.3-1 mm; for hindlimb ischemia, this means that dermal perfusion is assessed. Dermal perfusion is dependent on several factors&#8212;most importantly skin temperature and anesthetic agent, which must be carefully controlled to result in reliable readings. Furthermore, hair and skin pigmentation can alter the ability of the laser to either reach or penetrate to the dermis. This article demonstrates the technique of LDPI in the mouse hindlimb.

Here, we present a protocol that demonstrates the technique and necessary controls for Laser Doppler perfusion imaging to measure blood flow in the mouse hindlimb.

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is ...

The Mouse Isolated Perfused Kidney Technique

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a prerequisite for studying the physiology of the isolated organ and for many innovative applications that may be possible in the future, including perfusion decellularization for kidney bioengineering or the administration of anti-rejection or genome-editing drugs in high doses to prime the kidney for transplantation. During the time of the perfusion, the kidney can be manipulated, renal function can be assessed, and various pharmaceuticals administered. After the procedure, the kidney can be transplanted or processed for molecular biology, biochemical analysis, or microscopy.
This paper describes the perfusate and the surgical technique needed for the ex vivo perfusion of mouse kidneys. Details of the perfusion apparatus are given and data are presented showing the viability of the kidney&#39;s preparation: renal blood flow, vascular resistance, and urine data as functional, transmission electron micrographs of different nephron segments as morphological readouts, and western blots of transport proteins of different nephron segments as molecular readout.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. The buffers and surgical technique are described in detail.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a ...

In Situ Mouse Liver Perfusion Model to Study Metabolic Responses to Substrates

Hepatic Glucose Production, Ureagenesis, and Lipolysis Quantified using the Perfused Mouse Liver Model

The liver has numerous functions, including nutrient metabolism. In contrast to other in vitro and in vivo models of liver research, the isolated perfused liver allows the study of liver biology and metabolism in the whole liver with an intact hepatic architecture, separated from the influence of extra-hepatic factors. Liver perfusions were originally developed for rats, but the method has been adapted to mice as well. Here we describe a protocol for in situ perfusion of the mouse liver. The liver is perfused antegradely through the portal vein with oxygenated Krebs-Henseleit bicarbonate buffer, and the output is collected from the suprahepatic inferior vena cava with clamping of the infrahepatic inferior vena cava to close the circuit. Using this method, the direct hepatic effects of a test compound can be evaluated with a detailed time resolution. Liver function and viability are stable for at least 3 h, allowing the inclusion of internal controls in the same experiment. The experimental possibilities using this model are numerous and may infer insight into liver physiology and liver diseases.

Author Spotlight: Investigating Hepatic Adaptations and Prediabetic Progression in Liver Diseases 

Here, we present a robust method for in situ perfusion of the mouse liver to study the acute and direct regulation of liver metabolism without disturbing the hepatic architecture but in the absence of extra-hepatic factors.

The liver has numerous functions, including nutrient metabolism. In contrast to other in vitro and in vivo models of liver research, the isolated perfused ...

A Single Cell Dissociation Approach for Molecular Analysis of Urinary Bladder in the Mouse Following Spinal Cord Injury

We describe the implementation of spinal cord injury in mice to elicit detrusor-sphincter dyssynergia, a functional bladder outlet obstruction, and subsequent bladder wall remodeling. To facilitate assessment of the cellular composition of the bladder wall in non-injured control and spinal cord injured mice, we developed an optimized dissociation protocol that supports high cell viability and enables the detection of discrete subpopulations by flow cytometry.
Spinal cord injury is created by complete transection of the thoracic spinal cord. At the time of tissue harvest, the animal is perfused with phosphate-buffered saline under deep anesthesia and bladders are harvested into Tyrode&#8217;s buffer. Tissues are minced prior to incubation in digestion buffer that has been optimized based on the collagen content of mouse bladder as determined by interrogation of publicly available gene expression databases. Following generation of a single cell suspension, material is analyzed by flow cytometry for assessment of cell viability, cell number and specific subpopulations. We demonstrate that the method yields cell populations with greater than 90% viability, and robust representation of cells of mesenchymal and epithelial origin. This method will enable accurate downstream analysis of discrete cell types in mouse bladder and potentially other organs.

The goal of this protocol is to apply an optimized tissue dissociation protocol to a mouse model of spinal cord injury and validate the approach for single cell analysis by flow cytometry.

We describe the implementation of spinal cord injury in mice to elicit detrusor-sphincter dyssynergia, a functional bladder outlet obstruction, and ...

Establishing In Situ Closed Circuit Perfusion of Lower Abdominal Organs and Hind Limbs in Mice

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Programming Stem Cells for Therapeutic Angiogenesis Using Biodegradable Polymeric Nanoparticles

Murine Model of Metastatic Liver Tumors in the Setting of Ischemia Reperfusion Injury

Murine Model of Hindlimb Ischemia

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

Methods for Acute and Subacute Murine Hindlimb Ischemia

Laser Doppler Perfusion Imaging in the Mouse Hindlimb

The Mouse Isolated Perfused Kidney Technique

Hepatic Glucose Production, Ureagenesis, and Lipolysis Quantified using the Perfused Mouse Liver Model

A Single Cell Dissociation Approach for Molecular Analysis of Urinary Bladder in the Mouse Following Spinal Cord Injury