This work was supported by funding from the National Institutes of Health (R01-DK105346, P41-GM122698, 5U2C-DK119889). A portion of this work was performed in the McKnight Brain Institute at the National High Magnetic Field Laboratory&#39;s Advanced Magnetic Resonance Imaging and Spectroscopy (AMRIS) Facility, which is supported by the National Science Foundation Cooperative Agreement No. DMR-1644779 and the State of Florida.

Experiments involving mice were handled in compliance with the University of Florida Institutional Animal Care and Use Committee (protocol number #201909320). The mouse strain used was C57BL/6J; all mice were male. This method is generally applicable for studies using other standard mouse strains as well. This surgery is optimally performed by two individuals working together.
1. Initial set-up
<ol>
	<li>Perfuse livers with perfusate containing Krebs-Henseleit electrolytes6(25 mM NaHCO3, 112 mM NaCl, 4.7 mM KCl, 1.2 mM each of MgSO4, KH2PO4, and 0.5 mM sodium-EDTA, 1.25 mM CaCl2), 6 mM sodium lactate, 0.6 mM sodium pyruvate, 0.2 mM [U-13C] sodium propionate, 10% (v/v) D2O, and 0.63 mM mixed fatty acids (containing palmitic acid (22.1% of total), palmitoleic acid (5.2%), stearic acid (2.7%), oleic acid (27%), linoleic acid (37.7%), &#947;-linolenic acid (2.4%), and decosahexanoic acid (2.8%)) along with 2% (w/v) bovine serum albumin. Set final pH of perfusate to 7.3 using HCl (and NaOH, if necessary).</li>
</ol>
2. Pre-surgery set-up
<ol>
	<li>Assemble two 1 mL syringes with a 23G needle which are 19.05 mm long. Fill one syringe with 0.01 mL of 1000 units/mL heparin and 0.19 mL of saline (0.9% (w/v) NaCl in water; Table 1).</li>
	<li>Fill the second syringe with 0.2 mL of 2% lidocaine and 0.6 mL of 0.9% saline (Table 1). In another 1 mL syringe with a 27 G 38.1 mm needle, fill with the perfusate and maintain at 37 &#176;C.</li>
</ol>
3. Perfusate column set-up
<ol>
	<li>Set the glass bottle containing 500 mL of perfusate into the water bath (Figure 1B). Turn the water bath on and set the temperature to ~42 &#176;C. The higher temperature in the water bath enables maintaining 37 &#176;C in the perfusion column.</li>
	<li>Once the water heats up to 42 &#176;C, turn on the two pumps to circulate the perfusate from the bottle throughout the thin film oxygenator and the water-jacketed 100 cm glass column (Figure 1A-E).</li>
	<li>Turn on the oxygenating gas (95% oxygen and 5% carbon dioxide) to pressurize the oxygenator7 (Figure 1C). Adjust perfusate column height to achieve a flow rate of 8 mL/min with the catheter attached (see step 9 for flow rate measurement)5,8,9. 
	&#8203;NOTE: Flow rate refers to the rate at which perfusate is being expelled by the liver.</li>
</ol>
4. Anesthetization of the mouse
<ol>
	<li>Don PPE as required by IACUC protocol and other appropriate safety guidelines. 
	NOTE: The following steps have been optimized for mice that are 9 - 13 weeks old.</li>
	<li>Place the mouse in the isoflurane chamber. Turn the delivery gas to 100% oxygen, a flow rate of 1 L/min, and the isoflurane to 2%10. Wait till breathing slows down and is steady. 
	NOTE: For the mouse to reach a stable surgical plane, as evidenced by a slow and steady respiratory rate and lack of toe pinch reflex, the oxygen flow rate can be adjusted to ~1.5 L/min to ~3 L/min and the isoflurane concentration from&#160;1 - 3%. The delivery rate of carrier gas and isoflurane concentration depends on the animal&#39;s age and weight and factors such as noise and light.</li>
	<li>Disinfect the abdomen with 70% alcohol. Administer heparin through a deep subcutaneous injection in the abdominal fat layer (Figure 2). Place the mouse back into the anesthesia chamber for 10 min. 
	&#8203;NOTE: Shaving is not required as this procedure is terminal.</li>
</ol>
5. Celiotomy
<ol>
	<li>Transfer the mouse from the anesthesia chamber to the surgery platform and place it in the supine position (Figure 3).</li>
	<li>Place the nose of the mouse into a nose cone and tape the paws down. Take care to not apply any strain on the neck that may lead to suffocation.</li>
	<li>Administer lidocaine through a subcutaneous injection bilaterally in the anterior iliac crest region11 (Figure 3). Perform a toe pinch test to confirm the absence of all pain reflexes.</li>
	<li>Perform celiotomy to expose the internal organs (Figure 4). Make a 3 cm wide incision (the width of the entire abdomen of the mouse). 
	&#8203;NOTE: Width will change with the age and diet of the mouse.</li>
	<li>Expand the incision by using a hemostat that pulls traction by clamping on the xiphoid process (Figure 4).</li>
</ol>
6. Cannulation of portal vein
<ol>
	<li>Use a cotton-tipped applicator to clear the small and large intestines covering the portal vein. Position a silk suture under the arch of the portal vein proximal to the liver (Figure 4A).</li>
	<li>Depending on the anatomical structure, place the second silk suture proximal or distal of the inferior mesenteric vein distal from the liver (Figure 4A)12,13. Use a 2-0 suture for both sutures.</li>
	<li>Once the sutures are in place, cannulate the portal vein with a 22G catheter14 (Figure 4B). When inserting the catheter, keep the bevel pointed up. Enter the portal vein at no more than a 15&#176; angle.</li>
	<li>Tie the first suture past the catheter tapper. After the portal vein is cannulated, anchor the catheter 2-3 mm distal from the branch of the portal vein with the silk suture (Figure 4B). 
	NOTE: The assistant should roll shoulders and wrists to prevent dislodging the catheter or tearing of the portal vein. Each suture requires two knots.</li>
	<li>Next, secure the lower portion of the catheter with the second suture. With the help of the surgical assistant, tie a knot with the suture to secure the catheter to the distal portion of the portal vein and the surrounding tissue.</li>
</ol>
7. Resection of liver post portal vein cannulation
<ol>
	<li>After the catheter is secured, insert a 1 mL syringe with a 27G needle which is 38.1 mm long into the catheter to flush blood and air bubbles. 
	NOTE: There is usually a backflow of blood out of the catheter from the pressure.</li>
	<li>Use a 1 mm I.D. x 5 mm O.D. silicone tube with a fixed stopcock to couple the perfusion column (Figure 1A) to the catheter allowing the flow of the buffer into the liver marking the start of the perfusion. Start a timer at this point to mark the start of the perfusion.</li>
	<li>Relieve increased vascular pressure by making an incision, using scissors, into the inferior vena cava.</li>
	<li>Confirm flow of perfusate through the liver by observing the homogenous change in liver color from pink/red to a pale yellow. Once the flow is confirmed, excise the stomach, small intestine, large intestine, and the right kidney from surrounding tissue.</li>
	<li>With the help of the surgical assistant, maneuver the liver around the abdominal and thoracic cavity as the surgeon cuts through the parietal peritoneum and thoracic tissue to resect the liver</li>
	<li>Lastly, lift the liver upwards and cut the remaining connective tissues holding the liver in place with scissors. Slowly manipulate the liver for ease of view. Remove any fur that sticks to the liver by rinsing off with perfusate before encapsulating it within the NMR tube. 
	NOTE: In this procedure, only the liver is removed. All other organs are left in the body of the animal. The bile duct can be removed based on the protocol of the experiment. Although for this experiment, it was left in place.</li>
</ol>
8. Hanging the liver from the column
<ol>
	<li>Once the surgeon hands the liver and tubing to the assistant, the assistant disconnects the tubing from the catheter and column.</li>
	<li>Fill the catheter with perfusate until a meniscus is formed on the top of the catheter. Attach the catheter to the column for the liver to hang and perfuse. 
	NOTE: The bead of perfusate on the catheter provides sufficient volume for the liver to function until its connected.The catheter attached to the liver is press fitted to the bottom of the column.</li>
	<li>Screw a 20 mm NMR tube onto the 100 cm glass column to encapsulate the liver (Figure 5). To avoid torsion of the liver and the portal vein, slowly screw on the NMR tube. If torsion does occur and flow is stopped, unscrew and rescrew the NMR tube. This will remedy the occlusion and flow will return.</li>
	<li>Perfuse the liver for 30-60 min based on the details of the study. 
	&#8203;NOTE: Time is based on the perfusion experiment. For this experiment, metabolic turnover was measured within 30 min. The liver perfusion can take up to 10 min to reach a steady state. The time to steady-state starts once the inferior vena cava has been cut and hepatic flow is established.</li>
</ol>
9. Flow measurement
<ol>
	<li>Place a weigh boat on a top loading balance and zero the balance. Place the tubing from the roller pump pulling efferent perfusate from the NMR into the weigh boat and start the timer.</li>
	<li>Weigh the mass of liquid accumulated over 1 min which yields the flow rate of the liver. Place the tube back into the waste/collection container.</li>
</ol>
10. Oxygen Measurement
NOTE: Oxygen meter measurements were set up according to the manufacturer&#39;s instructions15.
<ol>
	<li>Place the electrode containing 20 &#181;L of 50% KCl saturated solution on the platinum dome and place five 10 &#181;L drops around the lower platinum ring of the electrode.</li>
	<li>Remove the adhesive off the cigarette paper. Lay a piece of polytetrafluoroethylene membrane over the cigarette paper.</li>
	<li>Place the two pieces on top of the electrode. Fit a small O-ring around the top of the electrode. Trim the paper to lay flat onto the lower platinum ring on the electrode. 
	NOTE: Some overhang is acceptable. It&#39;s essential to cover the silver of the electrode.</li>
	<li>Place the larger O-ring on the electrode. Couple the electrode to the water chamber and tighten the base to hold the electrode in place. Turn on the water bath and allow to heat up to 37 &#176;C.</li>
	<li>Open oxygen meter software. Click on Calibrate &#62; Air-Saturated Water. Set stirrer speed to 75 and the temperature to 37 &#176;C.</li>
	<li>Fill a 50 mL vial halfway with water and shake vigorously for 2 min. This is air-saturated water, use it as 100% standard. Fill the oxygen meter chamber with ~2 mL of water and place the two-piece stopper on.</li>
	<li>Click Okay on the screen and allow the signal to plateau. Once the signal has reached a plateau, click Okay. Dispose of liquid in the chamber and dry with tissue paper.</li>
	<li>Repeat steps 10.6-10.7 but with 200 mM sodium sulfite (0% standard). Click on Save calibration. 
	NOTE: No vigorous shaking is needed for the 0% standard.</li>
	<li>During perfusion, use two 5 mL syringes. One syringe for circulating perfusate (oxygen in) and the second for efferent perfusate from the NMR tube (oxygen out).</li>
	<li>When drawing up perfusate for both in and out measurements, draw 3-4 mL each time. 
	NOTE: This column has glass tubing to allow for access into the NMR tube to withdraw the perfusate that has flowed through the liver.</li>
	<li>Measure the circulating perfusate, first just as water in step 10.6 and dispose of in step 10.7. Repeat the same steps for the efferent perfusate.</li>
	<li>Perform oxygen measurements every 10 min.</li>
</ol>

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X., 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=Real-time+detection+of+hepatic+gluconeogenic+and+glycogenolytic+states+using+hyperpolarized+[2-13C]+dihydroxyacetone.">Real-time detection of hepatic gluconeogenic and glycogenolytic states using hyperpolarized [2-13C] dihydroxyacetone.</a> The Journal of Biological Chemistry. 289 (52), 35859-35867 (2014).</li><li>Moreno, K. X., 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=Hyperpolarized+&#948;-[1-13C]+gluconolactone+as+a+probe+of+the+pentose+phosphate+pathway.">Hyperpolarized &#948;-[1-13C] gluconolactone as a probe of the pentose phosphate pathway.</a> NMR in Biomedicine. 30 (6), (2017).</li><li>Merritt, M. E., Harrison, C., Sherry, A. D., Malloy, C. R., Burgess, S. 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The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

This surgical procedure is challenging and requires extensive practice to achieve reproducible results. Isoflurane and carrier gas should be adjusted as needed to maintain the viability of the animal through as much of the surgical procedure as possible. Environment, time of day, age, weight, and several other factors will affect anesthesia. Weight, diet, strain of mice and age could affect surgery as fat buildup can interfere with visualizing the portal vein. When taping the paws down, care must be taken to not apply an...

Ex vivo perfusions are crucial in the study of hepatic metabolism, and perfusion through the portal vein is the standard for these studies. In order to study hepatic metabolism in isolation, the liver must be resected from the body to avoid complications arising from metabolism in other organs (i.e., whole-body metabolism) and to exert control over hormone availability (insulin, glucagon, etc.). This approach can be essential for understanding the effects of diseases such as diabetes, NAFLD, and NASH on hepatic metabolism as well as mechanisms of drug action. This article serves as a guide to hepatic resection and perfusion. We have developed a streamlined procedure to perform these metabolic liver studies with sufficient rigor and reproducibility. If the surgery is not performed correctly, there is pronounced variability in the metabolic data obtained. We describe an organized method to perform portal vein catheterization and liver resection in the context of metabolic studies in situ in a nuclear magnetic resonance (NMR) spectrometer, as described in the literature1,2,3,4,5.
Currently, there is no literature describing an ex vivo hepatic perfusion using a glass column within an NMR. Nor is there a video or text publication providing a clear example of how to perform the procedure with the mouse liver, specifically, demonstrating how to catheterize the portal vein, resect a liver, transfer, and hang the liver onto a glass column. As the genetically modified mouse is ubiquitously used for studying liver metabolism, this is an essential procedure that deserves a complete description. Liver perfusion surgeries are not new, but this article is a gold standard method accompanied by a video demonstrating the technical excellence described in this paper to aid everyone interested in this procedure. The method presented here would be best applied to real-time metabolism to detect the function and turnover of metabolites in disease models.
This method uses a 100 cm water-jacketed glass column, which allows the liver to hang at the bottom of the cannula encapsulated by perfusate inside an NMR tube. Heated water in the glass jacket is used to control perfusate temperature. A thin layer oxygenator is pressurized with 95%/5% O2/CO2 for pH control. By using three separate pumps, the perfusate column height is set, which provides constant pressure to the liver. Flow rates are not controlled beyond the application of constant pressure (Figure 1). To confirm the liver is appropriately functioning, oxygen measurements are taken along with flow rates. In our hands, this set of preconditions leads to highly repeatable NMR experiments for the assessment of liver metabolic function.

<table><tbody><tr><td>1 mL Luer-Lock Single Use Sterile Disposable Syringe</td><td>N/A</td><td>N/A</td><td>Non-specific Brand</td></tr><tr><td>100 cm Water Jacketed Glass Column</td><td>N/A</td><td>N/A</td><td>Custom Made</td></tr><tr><td>2-0 Silk Suture</td><td>Braintree Scientific</td><td>N/A</td><td></td></tr><tr><td>22 Gauge Catherter 1 in. Without Safety</td><td>Terumo</td><td>SRFF2225</td><td></td></tr><tr><td>23 G 0.75 in. Hypodemeric Needles</td><td>Exel International</td><td>26407</td><td></td></tr><tr><td>27 G 1.5 in. Hypodemeric Needles</td><td>Exel International</td><td>26426</td><td></td></tr><tr><td>4x4 in. Surgical Platform</td><td>N/A</td><td>N/A</td><td>Custom Made</td></tr><tr><td>70% Alcohol Wipe</td><td>N/A</td><td>N/A</td><td>Non-specific Brand</td></tr><tr><td>Circulating Water Bath</td><td>MS Lauda</td><td>N/A</td><td>Model no longer manufactured</td></tr><tr><td>Cotton Tip Applicator</td><td>N/A</td><td>N/A</td><td>Non-specific Brand</td></tr><tr><td>Delicate Operating Scissors; Straight; Sharp-Sharp; 30mm Blade Length; 4 3/4 &quot;</td><td>Roboz</td><td>RS-6702</td><td></td></tr><tr><td>Dumont #5/45 Forceps</td><td>Fine Scientific Tools</td><td>11251-35</td><td></td></tr><tr><td>Dumont #7 - Fine Forceps</td><td>Fine Scientific Tools</td><td>11274-20</td><td></td></tr><tr><td>Hemostats</td><td>Fine Scientific Tools</td><td>13015-14</td><td></td></tr><tr><td>Heparin Sodium Injectable 1000 units/mL</td><td>RX Generics</td><td>71288-0402-02</td><td></td></tr><tr><td>Isoflurane</td><td>Patterson Veterinary</td><td>14043-0704-06</td><td></td></tr><tr><td>Lidocaine HCl 2%</td><td>VEDCO Inc.</td><td>50989-0417-12</td><td></td></tr><tr><td>Membrane-Thin-Layer Oxygenator</td><td>Radnoti</td><td>N/A</td><td></td></tr><tr><td>Metzenbaum Scissors; Curved; Blunt; 27 mm Blade Length; 5 &quot;</td><td>Roboz</td><td>RS-6013</td><td></td></tr><tr><td>Oxygen Meter System</td><td>Hanstech Instruments Ltd.</td><td>N/A</td><td></td></tr><tr><td>Saline 0.9% Solution</td><td>N/A</td><td>N/A</td><td>Saline is made in lab</td></tr><tr><td>Scale</td><td>N/A</td><td>N/A</td><td>Non-specific Brand</td></tr><tr><td>&amp;nbsp;Variable Speed Analog Console Pump Systems</td><td>Cole Palmer</td><td>N/A</td><td>Models are custom per application</td></tr><tr><td>Weigh boats</td><td>N/A</td><td>N/A</td><td>Non-specific Brand</td></tr></tbody></table>

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.

Liver function is primarily assessed by oxygen consumption and flow rate. A flow rate of 4-8 mL/min and oxygen consumption of 1 &#181;mol/min.g is typical. These measures will vary depending on specific experimental conditions and biological differences.
The exact amount of isoflurane used will depend on the type of anesthesia system being used as well as the environment and age/weight of the mouse. During the surgery, the isoflurane and delivery gas do not change, although some changes may be...

Watch this Scientific Journal Video about Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse at JoVE.com

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.

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 ...

ex-vivo-hepatic-perfusion-through-the-portal-vein-in-mouse

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

8.6K Views.  University of Florida. The protocol describes a straightforward method of resectioning an intact mouse liver for metabolic studies through portal vein perfusion.

Video: Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Purification of Hepatocytes and Sinusoidal Endothelial Cells from Mouse Liver Perfusion

This protocol demonstrates a method for obtaining high yield and viability for mouse hepatocytes and sinusoidal endothelial cells (SECs) suitable for culturing or for obtaining cell lysates. In this protocol, the portal vein is used as the site for catheterization, rather than the vena cava, as this limits contamination of other possible cell types in the final liver preparation. No special instrumentation is required throughout the procedure. A water bath is used as a source of heat to maintain the temperature of all the buffers and solutions. A standard peristaltic pump is used to drive the fluid, and a refrigerated table-top centrifuge is required for the centrifugation procedures. The only limitation of this technique is the placement of the catheter within the portal vein, which is challenging on some of the mice in the 18 - 25 g size range. An advantage of this technique is that only one vein is utilized for the perfusion and the access to the vein is quick, which minimizes ischemia and reperfusion of the liver that reduces hepatic cell viability. Another advantage to this protocol is that it is easy to distinguish live from dead hepatocytes by eyesight due to the difference in cellular density during the centrifugation steps. Cells from this protocol may be used in cell culture for any downstream application as well as processed for any biochemical assessment.

The goal of this protocol is to obtain high viability and high yield of hepatocytes and sinusoidal endothelial cells from liver. This is accomplished by perfusing the liver with a type IV collagenase solution via the portal vein, followed by differential centrifugation to obtain hepatocytes and sinusoidal endothelial cells.

This protocol demonstrates a method for obtaining high yield and viability for mouse hepatocytes and sinusoidal endothelial cells (SECs) suitable for ...

Measurement of Fatty Acid &#946;-Oxidation in a Suspension of Freshly Isolated Mouse Hepatocytes

Fatty acid &#946;-oxidation is a key metabolic pathway to meet the energy demands of the liver and provide substrates and cofactors for additional processes, such as ketogenesis and gluconeogenesis, which are essential to maintain whole-body glucose homeostasis and support extra-hepatic organ function in the fasted state. Fatty acid &#946;-oxidation occurs within the mitochondria and peroxisomes and is regulated through multiple mechanisms, including the uptake and activation of fatty acids, enzyme expression levels, and availability of cofactors such as coenzyme A and NAD+. In assays that measure fatty acid &#946;-oxidation in liver homogenates, cell lysis and the common addition of supraphysiological levels of cofactors mask the effects of these regulatory mechanisms. Furthermore, the integrity of the organelles in the homogenates is hard to control and can vary significantly between preparations. The measurement of fatty acid &#946;-oxidation in intact primary hepatocytes overcomes the above pitfalls. This protocol describes a method for the measurement of fatty acid &#946;-oxidation in a suspension of freshly isolated primary mouse hepatocytes incubated with 14C-labeled palmitic acid. By avoiding hours to days of culture, this method has the advantage of better preserving the protein expression levels and metabolic pathway activity of the original liver, including the activation of fatty acid &#946;-oxidation observed in hepatocytes isolated from fasted mice compared to fed mice.

Fatty acid &#946;-oxidation is an essential metabolic pathway responsible for generating energy in many different cell types, including hepatocytes. Here, we describe a method to measure fatty acid &#946;-oxidation in freshly isolated primary hepatocytes using 14C-labeled palmitic acid.

Fatty acid &#946;-oxidation is a key metabolic pathway to meet the energy demands of the liver and provide substrates and cofactors for additional ...

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

Visualization of Vascular and Parenchymal Regeneration after 70% Partial Hepatectomy in Normal Mice

A modified silicone injection procedure was used for visualization of the hepatic vascular tree. This procedure consisted of in-vivo injection of the silicone compound, via a 26 G catheter, into the portal or hepatic vein. After silicone injection, organs were explanted and prepared for ex-vivo micro-CT (&#181;CT) scanning. The silicone injection procedure is technically challenging. Achieving a successful outcome requires extensive microsurgical experience from the surgeon. One of the challenges of this procedure involves determining the adequate perfusion rate for the silicone compound. The perfusion rate for the silicone compound needs to be defined based on the hemodynamic of the vascular system of interest. Inappropriate perfusion rate can lead to an incomplete perfusion, artificial dilation and rupturing of vascular trees.
The 3D reconstruction of the vascular system was based on CT scans and was achieved using preclinical software such as HepaVision. The quality of the reconstructed vascular tree was directly related to the quality of silicone perfusion. Subsequently computed vascular parameters indicative of vascular growth, such as total vascular volume, were calculated based on the vascular reconstructions. Contrasting the vascular tree with silicone allowed for subsequent histological work-up of the specimen after &#181;CT scanning. The specimen can be subjected to serial sectioning, histological analysis and whole slide scanning, and thereafter to 3D reconstruction of the vascular trees based on histological images. This is the prerequisite for the detection of molecular events and their distribution with respect to the vascular tree. This modified silicone injection procedure can also be used to visualize and reconstruct the vascular systems of other organs. This technique has the potential to be extensively applied to studies concerning vascular anatomy and growth in various animal and disease models.

Tools used for visualizing vascular regeneration require methods for contrasting the vascular trees. This film demonstrated a delicate injection technique used to achieve optimal contrasting of the vascular trees and illustrate the potential benefits resulting from a detailed analysis of the resulting specimen using &#181;CT and histological serial sections.

A modified silicone injection procedure was used for visualization of the hepatic vascular tree. This procedure consisted of in-vivo injection of the ...

Medicine

Electroporation-Mediated Delivery of Cas9 Ribonucleoproteins and mRNA into Freshly Isolated Primary Mouse Hepatocytes

This protocol describes a fast and effective method for isolating primary mouse hepatocytes followed by electroporation-mediated delivery of CRISPR-Cas9 as ribonucleoproteins (RNPs) and mRNA. Primary mouse hepatocytes were isolated using a three-step retrograde perfusion method resulting in high yields of up to 50 &#215; 106 cells per liver and cell viability of &#62;85%. This protocol provides detailed instructions for plating, staining, and culturing hepatocytes. The results indicate that electroporation provides a high transfection efficiency of 89%, as measured by the percentage of green fluorescent protein (GFP)-positive cells and modest cell viability of &#62;35% in mouse hepatocytes.
To demonstrate the utility of this approach, CRISPR-Cas9 targeting the hydroxyphenylpyruvate dioxygenase gene was electroporated into primary mouse hepatocytes as proof-of-principle gene editing to disrupt a therapeutic gene related to an inherited metabolic disease (IMD) of the liver. A higher on-target edit of 78% was observed for RNPs compared to 47% editing efficiency with mRNA. The functionality of hepatocytes was evaluated in vitro using an albumin assay that indicated that delivering CRISPR-Cas9 as RNPs and mRNA results in comparable cell viability in primary mouse hepatocytes. A promising application for this protocol is the generation of mouse models for human genetic diseases affecting the liver.

This protocol describes techniques for isolating primary mouse hepatocytes from the liver and electroporating CRISPR-Cas9 as ribonucleoproteins and mRNA to disrupt a therapeutic target gene associated with an inherited metabolic disease of the liver. The methods described result in high viability and high levels of gene modification after electroporation.

This protocol describes a fast and effective method for isolating primary mouse hepatocytes followed by electroporation-mediated delivery of CRISPR-Cas9 ...

Bioengineering

3D Imaging of the Liver Extracellular Matrix in a Mouse Model of Non-Alcoholic Steatohepatitis

Non-alcoholic steatohepatitis (NASH) is the most common chronic liver disease in the United States, affecting more than 70 million Americans. NASH can progress to fibrosis and eventually to cirrhosis, a significant risk factor for hepatocellular carcinoma. The extracellular matrix (ECM) provides structural support and maintains liver homeostasis via matricellular signals. Liver fibrosis results from an imbalance in the dynamic ECM remodeling process and is characterized by excessive accumulation of structural elements and associated changes in glycosaminoglycans. The typical fibrosis pattern of NASH is called &#34;chicken wire,&#34; which usually consists of zone 3 perisinusoidal/pericellular fibrosis, based on features observed by Masson&#39;s trichrome stain and Picrosirius Red stains. However, these traditional thin two-dimensional (2D) tissue slide-based imaging techniques cannot demonstrate the detailed three-dimensional (3D) ECM structural changes, limiting the understanding of the dynamic ECM remodeling in liver fibrosis.
The current work optimized a fast and efficient protocol to image the native ECM structure in the liver via decellularization to address the above challenges. Mice were fed either with chow or fast-food diet for 14 weeks. Decellularization was performed after in situ portal vein perfusion, and the two-photon microscopy techniques were applied to image and analyze changes in the native ECM. The 3D images of the normal and NASH livers were reconstituted and analyzed. Performing in situ perfusion decellularization and analyzing the scaffold by two-photon microscopy provided a practical and reliable platform to visualize the dynamic ECM remodeling in the liver.

The present protocol optimizes the liver in situ perfusion/decellularization and two-photon microscopy methods to establish a reliable platform to visualize the dynamics of extracellular matrix (ECM) remodeling during non-alcoholic steatohepatitis (NASH).

Non-alcoholic steatohepatitis (NASH) is the most common chronic liver disease in the United States, affecting more than 70 million Americans. NASH can ...

Isolation of Regenerating Hepatocytes after Partial Hepatectomy in Mice

Partial hepatectomy has been widely used to investigate liver regeneration in mice, but the isolation of high yields of viable hepatocytes for downstream single-cell applications is challenging. A marked accumulation of lipids within regenerating hepatocytes is observed during the first 2 days of normal liver regeneration in mice. This so-called transient regeneration-associated steatosis (TRAS) is temporary but partially overlaps the major proliferative phase. Density-gradient purification is the backbone of most existing protocols for the isolation of primary hepatocytes. As gradient purification relies on the density and size of cells, it separates non-steatotic from steatotic hepatocyte populations. Therefore, fatty hepatocytes often are lost, yielding non-representative hepatocyte fractions. 
The presented protocol describes an easy and reliable method for the in vivo isolation of regenerating hepatocytes regardless of their lipid content. Hepatocytes from male C57BL/6 mice are isolated 24-48 h after hepatectomy by a classic two-step collagenase perfusion approach. A standard peristaltic pump drives the warmed solutions via the catheterized inferior vena cava into the remnant, using a retrograde perfusion technique with outflow through the portal vein. Hepatocytes are dissociated by collagenase for their release from the Glisson's capsule. After washing and careful centrifugation, the hepatocytes can be used for any downstream analyses. In conclusion, this paper describes a straightforward and reproducible technique for the isolation of a representative population of regenerating hepatocytes after partial hepatectomy in mice. The method may also aid the study of fatty liver disease.

Lipid-laden hepatocytes are inherent to liver regeneration but are usually lost upon density-gradient centrifugation. Here, we present an optimized cell isolation protocol that retains steatotic hepatocytes, yielding representative populations of regenerating hepatocytes after partial hepatectomy in mice.

Partial hepatectomy has been widely used to investigate liver regeneration in mice, but the isolation of high yields of viable hepatocytes for downstream ...

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 ...

Noninvasive Intraocular In Vivo Imaging of Engrafted Liver Spheroids in Mice

In Vivo Imaging of Liver Spheroids Engrafted in the Anterior Chamber of the Mouse Eye

Biomedical studies of the liver in mammals are hindered by the lack of methods for in vivo noninvasive longitudinal imaging at cellular resolution. Until now, optical imaging of the liver in situ is possible by intravital imaging, which offers high-resolution imaging at the cellular level but cannot be performed multiple times and, therefore, longitudinally in the same animal. Noninvasive imaging methods, such as bioluminescence, allow repeated imaging sessions on the same animal but do not achieve cell resolution. To address this methodology gap, we have developed a platform for noninvasive in vivo imaging of liver spheroids engrafted in the anterior chamber of the mouse eye. In the workflow described in this study, primary mouse liver spheroids are generated in vitro and transplanted into the anterior chamber of the eye of recipient mice, where they engraft on the iris. The cornea acts as a natural body window through which we can image the engrafted spheroids by conventional confocal microscopy. The spheroids survive for months in the eye, during which the cells can be studied in contexts of health and disease, as well as being monitored in response to different stimuli over repeated imaging sessions using appropriate fluorescent probes. In this protocol, we provide a breakdown of the necessary steps to implement this imaging system and explain how to best harness its potential.

Author Spotlight: Harnessing Mouse Eye Chambers for Noninvasive Liver Spheroid Studies

Here, we describe a platform that allows noninvasive in vivo imaging of liver spheroids engrafted in the anterior chamber of the mouse eye. The workflow spans from generating spheroids from primary liver cells to transplantation into the mouse eye and in vivo imaging at cellular resolution by confocal microscopy.

Biomedical studies of the liver in mammals are hindered by the lack of methods for in vivo noninvasive longitudinal imaging at cellular resolution. Until ...

Biology

Mice Liver Perfusion for Accurate Evaluation of Hepatic Insulin Clearance

Assessing Insulin Clearance in Mice via In Situ Liver Perfusion

Hepatic insulin clearance is essential for maintaining glucose homeostasis and is closely linked to metabolic disorders such as obesity, insulin resistance, and diabetes. Accurate measurement of insulin clearance is vital for understanding the underlying mechanisms of these conditions. This protocol presents a straightforward and user-friendly hepatic perfusion procedure in mice, specifically designed to directly evaluate the hepatic insulin clearance rate. The method involves precise cannulation of the portal vein and suprahepatic inferior vena cava to create an in situ perfusion system that mimics physiological conditions. The protocol guides researchers through every stage of the procedure, from surgical preparation and setting up the perfusion system to sample collection and analysis. Detailed instructions are provided, along with representative results and important tips for optimizing the procedure. A video tutorial accompanies the written protocol, offering visually in-depth instructions and illustrations, making it an accessible and comprehensive reference for scientists exploring the molecular mechanisms behind hepatic insulin metabolism and clearance.

Hepatic insulin clearance is critical for regulating glucose homeostasis. This article describes a user-friendly hepatic perfusion procedure for directly evaluating the hepatic insulin clearance rate in situ in mice.

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Summary

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Chapters in this video

Purification of Hepatocytes and Sinusoidal Endothelial Cells from Mouse Liver Perfusion

Measurement of Fatty Acid β-Oxidation in a Suspension of Freshly Isolated Mouse Hepatocytes

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

Visualization of Vascular and Parenchymal Regeneration after 70% Partial Hepatectomy in Normal Mice

Electroporation-Mediated Delivery of Cas9 Ribonucleoproteins and mRNA into Freshly Isolated Primary Mouse Hepatocytes

3D Imaging of the Liver Extracellular Matrix in a Mouse Model of Non-Alcoholic Steatohepatitis

Isolation of Regenerating Hepatocytes after Partial Hepatectomy in Mice

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

In Vivo Imaging of Liver Spheroids Engrafted in the Anterior Chamber of the Mouse Eye

Assessing Insulin Clearance in Mice via In Situ Liver Perfusion