The studies and Nicolai J. Wewer Albrechtsen were supported by Novo Nordisk Foundation Excellence Emerging Investigator Grant - Endocrinology and Metabolism (Application No. NNF19OC0055001), European Foundation for the Study of Diabetes Future Leader Award (NNF21SA0072746) and Independent Research Fund Denmark, Sapere Aude (1052-00003B). Novo Nordisk Foundation Center for Protein Research is supported financially by the Novo Nordisk Foundation (Grant agreement NNF14CC0001). Figure 1B was created with biorender.com.&#160;We thank Dr. Rune E. Kuhre (Novo Nordisk A/S) for fruitful discussions on the perfused mouse liver.&#160;

All animal experiments were conducted with permission from the Danish Animal Experiments Inspectorate, Ministry of Environment and Food of Denmark (permit 2018-15-0201-01397), and the local ethics committee in accordance with the EU directive 2010/63/EU, the National Institutes of Health (publication No. 85-3) and following the guidelines of Danish legislation governing animal experimentation (1987). This is a terminal procedure, and the cause of death is exsanguination and perforation of the diaphragm under deep anesthesia.
1. Experimental animals
<ol>
	<li>Obtain mice of desired strain, age, and gender. This study used male C57BL/6JRj mice aged 11-16 weeks. House up to 5 male or 8 female mice per cage with ad libitum access to chow and water and maintain a 12 h/12 h light-dark cycle with lights on from 6 AM to 6 PM.</li>
</ol>
2. Preoperative preparations
<ol>
	<li>Make liver perfusion buffer.
	<ol>
		<li>Krebs-Henseleit Buffer. Mix and dissolve 118 mmol/L NaCl, 4.7 mmol/L KCl, 1.2 mmol/L MgSO4, and 1.2 mmol/L KH2PO4 in dH2O. Dissolve 1.25 mmol/L CaCl2 in a separate beaker and add the buffer.</li>
		<li>Dissolve 25 mmol/L NaHCO3 and add slowly to the buffer while stirring. Store the buffer at 4 deg. The buffer is stable for at least 1 month. 
		NOTE: Precipitation may occur if CaCl2 and NaHCO3 are not dissolved individually before being added.</li>
	</ol>
	</li>
	<li>Filter the perfusion buffer through a 2 &#181;m filter and adjust pH to 7.5 using HCl. 
	NOTE: This step should be performed on the experimental day, as pH increases over time, even when stored in closed flasks.</li>
	<li>Prepare test compounds in a higher concentration than the desired final concentration (e.g., 20x concentration when infused at a rate of 0.175 mL/min via a sidearm pump) in filtered and pH-adjusted Krebs-Henseleit perfusion buffer. If relevant, dilute test compounds in Krebs-Henseleit perfusion buffer with 1% BSA added as a carrier (to avoid adhesion to tubing and glassware, essential for all peptides), filtered through an appropriate size filter.</li>
</ol>
3. Operation and perfusion
NOTE: An illustration of the perfusion setup used in this study is provided in Figure 1.
<ol>
	<li>Gas the perfusion buffer (95% O2, 5% CO2) for at least 30 min to supply sufficient oxygen to the liver and to maintain the correct pH from the start of the operation (the bicarbonate buffer system will reach a pH of 7.4 under continuous gassing, see Supplementary Figure 1).</li>
	<li>Anesthetize a mouse by administering ketamine (90 mg/kg) and xylazine (10 mg/kg) by intraperitoneal injection.</li>
	<li>Place the mouse supine on a heated operation table and confirm the lack of reflexes in response to the toe pinch. Spray with 70% ethanol to keep hair from sticking to the scissors. Fixing the mouse to the warm surface helps to increase stability for the following procedures.</li>
	<li>Make an incision with scissors at the base of the abdomen and cut upwards to the ribcage on both sides to expose the abdominal cavity. Move the intestines to the right using a cotton swab, exposing the portal vein.</li>
	<li>Place two ligatures under the portal vein using curved forceps and prepare a loose knot for each ligature.</li>
	<li>Insert a 0.7 mm catheter into the portal vein. When perforated, remove the needle from the catheter and guide the catheter through the vein until the tip of the catheter is close to the liver, as illustrated in Figure 2. Blood flows into the catheter.</li>
	<li>Tighten the ligatures. If the catheter is not filled with blood, fill it up with perfusion buffer to avoid the introduction of an air bubble.</li>
	<li>Attach the perfusion tube and initiate perfusion of the liver with Krebs-Henseleit bicarbonate buffer at 37 &#176;C by starting the roller pump at a perfusion flow rate of 0.8 mL/min. The liver turns pale within seconds.</li>
	<li>Cut the ribcage and diaphragm using scissors. At this point, the animal is euthanized. Place a ligature under the suprahepatic inferior vena cava using fine point forceps. Place a pen, rolled gauze, or another disposable item under the back of the mouse to make the vein more accessible.</li>
	<li>Insert a catheter into the suprahepatic inferior vena cava via the right atrium of the heart. When perforated, remove the needle from the catheter and guide the catheter through the vein until the tip of the catheter is close to the liver. Blood and perfusion buffer runs out immediately.</li>
	<li>Tighten the ligature and attach a tube for the collection of perfusion effluent. Secure all tubes with waterproof tape (sleek tape or similar).</li>
	<li>Use a vessel clamp adaptor to place a vessel clamp across the infrahepatic vena cava immediately above the right renal vein to prevent admixture (Figure 2).</li>
	<li>Increase the perfusion flow rate to 3.5 mL/min and start a timer. Start the pressure recording. Successful perfusion usually results in a pressure of ~10 mm/Hg.</li>
	<li>Cover the liver with a sterile drape moistened with saline and add saline during the experiment to prevent it from drying.</li>
	<li>Collect the perfusion effluent for 1 min and measure the volume. The volume should be approximately 3.5 mL/min. Admixture of blood is not expected at this stage, and the heart does not pump any longer.</li>
	<li>Wait for an equilibration period of 30 min before starting the experiment.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65596/65596fig01.jpg" /> 
Figure 1: An illustration of the perfusion setup. (A)The operating table is elevated on a tripod stand and heated to 37 &#176;C. The perfusion buffer is gassed (95% O2, 5% CO2), pumped via a peristaltic roller pump, and heated in the heat exchanger with a built-in bubble trap. The system furthermore consists of a pressure gauge and spindle pump for adjustment of the perfusion pressure. The perfusion pressure is continuously recorded and visualized via a transducer on a PC, a pressure recording program. (B) The red box captures the connections of three-way stopcocks. The first three-way stopcock is open for the infusion of a test compound via a syringe pump, and the second is closed. The third is open for continuous pressure measurements. The fourth stopcock may be used to collect input samples, for e.g., gas analysis across the perfused liver. The connectors can be modified as needed for specific experiments requiring more or fewer infusion lines. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65596/65596fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65596/65596fig02.jpg" /> 
Figure 2: Photos of the mouse abdominal cavity before and during liver perfusion. (A)The green dot indicates the location of the tip of the portal vein catheter. It is important that the tip of the catheter is positioned just below the branching point of the portal vein into the left and right hepatic portal veins but above the pancreato-duodenal branch to avoid leakage. The yellow dot indicates the correct location of the vessel clamp on the infrahepatic inferior vena cava between the right renal vein and the liver to avoid the backflow of blood into the perfused liver. (B, C) A perfused mouse liver with the two catheters inserted in the portal vein (B) and suprahepatic inferior vena cava (C) and the vessel clamp on the infrahepatic inferior vena cava (B). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65596/65596fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Experiment
<ol>
	<li>Start the experiment by collecting the first baseline samples using a fraction collector at the end of the equilibration period. Collect samples at the desired time interval and place them on ice immediately.&#160;</li>
	<li>Check the bubble trap regularly and refill with perfusion buffer when close to empty.</li>
	<li>Collect a sample of the buffer through a three-way stopcock immediately before it enters the organ and from the collecting catheter inserted in the inferior vena cava (after it has been perfused through the liver).</li>
	<li>Measure the samples with a blood-gas analyzer to confirm that the organ is metabolically active (indicated by an increase in the partial pressure of CO2 and a decrease in pH).</li>
	<li>Repeat steps 4.3-4.4 at the end of the experiment to assess viability throughout the experiment (Supplementary Figure 2). 
	NOTE: The decrease in oxygen partial pressure does not provide a reliable measure of respiration because of losses of oxygen from tubing, the organ etc.</li>
	<li>After 15 min of baseline perfusion, start the first stimulation by infusing a test substance through a three-way stopcock at desired flow rate using a syringe pump (e.g., 20x concentration of the test substance when infused at a rate of 0.175 mL/min via a sidearm pump). Alternatively, switch the baseline buffer to a new (oxygenated and heated) buffer containing the test compound in the final concentration.</li>
	<li>Stop the stimulation and collect baseline samples for 20-30 min before starting the second stimulation.</li>
	<li>At the end of the experiment, infuse an appropriate positive control for 5-10 min.</li>
	<li>After the experiment, excise the perfused liver and weigh it to normalize the output to liver weight. Snap-freeze the liver in liquid nitrogen for potential measurement of protein content to normalize the output to protein content.</li>
</ol>
5. Biochemical measurements
<ol>
	<li>Quantify the concentration of the molecule of interest using assays appropriate for measurements in perfusion buffer (in-house or commercially available colorimetric or ELISA assays). The buffer is compatible with most omics-based techniques such as metabolomics and proteomics. 
	NOTE: In this study, urea was measured based on a colorimetric assay previously described14. Glucose and non-esterified fatty acids were quantified using commercially available kits.</li>
</ol>
6. Data analysis
<ol>
	<li>Present the data in XY graphs showing the secretory output over time. 
	NOTE: It is one of the virtues of the in vitro perfusion system that it is possible to express data as the actual output (concentration &#215; flow rate, e.g., &#181;mol/min) instead of the measured concentration in the output (mmol/L) as in in vivo studies. Consider normalizing the output to the liver weight when comparing different mouse models or to the total protein content (measured by BCA) when comparing control mice to mouse models with obesity or liver diseases where an increase in liver weight may be due to increased fat content.</li>
	<li>Present summary data as individual dots per animal representing the mean or total output during baseline and over a stimulation period (typically 15-30 min) or incremental output using the preceding baseline for each stimulation period dependent on the study design.</li>
	<li>Analyze the data using paired t-test (two groups) or one-way ANOVA with repeated stimulations (more than two groups) with an appropriate post-hoc test for multiple testing.</li>
</ol>

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

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	<li>Bale, S. S., Geerts, S., Jindal, R., Yarmush, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+co-culture+of+rat+parenchymal+and+non-parenchymal+liver+cells+to+evaluate+cellular+interactions+and+response.">Isolation and co-culture of rat parenchymal and non-parenchymal liver cells to evaluate cellular interactions and response.</a> Scientific Reports. 6, 25329 (2016).</li><li>Lauschke, V. 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C., 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+primary+human+hepatocyte+spheroids+as+a+model+system+for+drug-induced+liver+injury,+liver+function,+and+disease.">Characterization of primary human hepatocyte spheroids as a model system for drug-induced liver injury, liver function, and disease.</a> Scientific Reports. 6, 25187 (2016).</li><li>Dewyse, L., Reynaert, H., van Grunsven, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Best+practices+and+progress+in+precision-cut+liver+slice+cultures.">Best practices and progress in precision-cut liver slice cultures.</a> International Journal of Molecular Sciences. 22 (13), 7137 (2021).</li><li>Li, X., George, S. M., Vernetti, L., Gough, A. H., Taylor, D. 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The authors declare no conflicts of interest relevant to this article.

The isolated perfused mouse liver is a strong research tool for studies of the dynamics and molecular mechanisms of hepatic metabolism. The possibility of minute-to-minute sample collection provides a detailed evaluation of the direct effect of a test compound on the liver. Compared to in vivo studies, the perfused liver allows us to study liver metabolism in an isolated manner avoiding extra-hepatic factors carried by the blood and with complete control over the experimental conditions. The advantages of liver perfusion compared to in vitro studies using isolated hepatocytes are the maintenance of the hepatic architecture, polarity, zonal division, and vascular integrity. Another advantage of liver perfusion is the large sample size volume (3.5 mL/min), allowing measurement of several outcomes in the same sample. The perfused liver is, however, not ideal when studying effects dependent on transcriptional regulation since such mechanisms may take several hours to occur. In addition to metabolic research, the protocol may be applied in other research fields to study the endocrine functions of the liver (secretion of hormones and hepatokines) and liver metabolism of drugs and xenobiotics.
The most critical step for this method is the placement of the catheter in the portal vein. It is important that the tip of the catheter is positioned immediately before the branch point of the portal vein into the left and right hepatic portal veins (Figure 2). If the tip of the catheter is pushed too far, only part of the liver will be properly perfused. If it is placed too far down in the portal vein, leakage through the pancreato-duodenal branch may occur. Placement of the second catheter for collection is less urgent since the liver at this step is already perfused with the oxygenated buffer. When the operation has been successfully performed, the liver will be respiring and responsive for at least 3 h with a constant pressure and perfusion output (Supplementary Figure 2). At this stage, the avoidance of air bubbles in the perfusion system is the most important factor for a successful experiment. Air bubbles are difficult to avoid completely since the buffer is continuously gassed with oxygen, but the bubbles should be trapped in the bubble trap of the perfusion system. It is therefore important to keep an eye on the bubble trap and refill it with perfusion buffer when it is close to empty.
Krebs-Henseleit buffer (KHB) is the most widely used solution for liver perfusions13. The classical formulation contains 2.5 mmol/L CaCl2 but based on a previous study showing that calcium contributes to mitochondrial damage, we use a buffer with only 1.25 mmol/L CaCl212. The addition of erythrocytes, albumin, or glucose to the buffer is not needed11. The proposed buffer also allows molecular profiling using advanced biochemical techniques such as mass-spectrometry-based metabolomics. Different concentrations of a test compound may be tested to determine the optimal dose and to evaluate physiological vs. pharmacological effects. In the experiments conducted in this study, 10 mM AA is in the high physiological range corresponding to postprandial levels, whereas 10 nM glucagon corresponds to a pharmacological dose. As demonstrated in Figure 4, it is essential to include 20-30 min baseline period between two consecutive stimulations to be able to compare the dynamics and molecular mechanisms of the respective responses.
It is important to avoid blood in the perfusate samples as it may interfere with the subsequent analyses, e.g., colorimetric determination of urea and glucose concentrations. If blood appears in the collected samples after ~15 min into the equilibration period, when a 3-way valve is opened for infusions, or when the bubble trap is filled, the vessel clamp on the infrahepatic vena cava is likely not placed correctly.
In our experience, a successful operation results in a stable portal pressure of ~10 mmHg and an output flow rate of 3.5 mL/min within a few minutes after inserting the collecting catheter. If the pressure is considerably higher or continuously increasing, or if the collected output flow rate is not 3.5 mL/min or continuously dropping, several events should be considered: 1) Are all liver lobes equally perfused and showing a uniform pale color? If not, an occlusion may have occurred (often an air bubble; carefully &#34;massage&#34; the liver lobe with a cotton stick; this sometimes helps to release the air bubble whereby the pressure will drop immediately), or the portal vein catheter has been pushed too far up, thus supplying only one of the two branches of the hepatic portal vein (cautiously try to pull back the catheter a few millimeters). 2) Is the suprahepatic inferior vena cava twisted? This can occur when attaching the male part of the collecting tube to the female part of the catheter inserted in the vein; cautiously remove the collecting tube and observe if the pressure drops. 3) Is the collecting catheter pushed too far down the suprahepatic inferior vena cava so the tip of the catheter has wedged into the liver? Cautiously retract a few millimeters if needed.
In summary, the perfused mouse liver is a physiologically relevant in vitro experimental model that can be used to study the dynamic effects of a given compound on the liver. The quality of the operation is critical for the quality of data obtained, and in our experience, it takes ~2 months of training before data of satisfactory quality can be expected. Once the technique has been learned, the possibilities with this method are numerous, including the use of transgenic or knockout donor animals regarding the genes of interest as well as mouse models of liver diseases. Experiments may, however, be limited by potential toxic side effects (which are likely to be much more severe in vivo) or poor solubility of test compounds. Such compounds may be given orally to the donor animals at an appropriate time period before perfusion.

The liver is an essential organ in metabolism. It plays a key role in the control of whole-body energy balance by regulating glucose, lipid, and amino acid metabolism. The increase in liver diseases worldwide is emerging as a major global health burden, and more knowledge is needed about the pathophysiology and its consequences for liver functions.
Various in vitro models have been developed for research on the liver to complement in vivo studies. Isolated and cultured primary hepatocytes from rodents and humans are widely used. Non-parenchymal cells can be separated from hepatocytes using differential and gradient centrifugation, and the co-culture of different cell types is useful for studying intercellular crosstalk1. Although primary human hepatocytes are considered the golden standard for testing drug toxicity, several studies have shown that the hepatocytes rapidly dedifferentiate in tissue culture resulting in loss of hepatic functions2,3,4. Hepatocyte culture in a 3D spheroid system ameliorates the dedifferentiation, is more stable, and appears to mimic the liver in vivo to a higher degree than the traditional 2D culture systems5. Precision-cut liver slices are another well-established in vitro model that keeps the tissue architecture intact and contains the non-parenchymal cells present in the liver6. More advanced in vitro models include liver-on-a-chip7 and liver organoids8. However, with all these approaches, there is a loss of structural integrity and flow dynamics, including vectorial portal-hepatic vein flow, which likely impacts the generalizability.
The isolated perfused rat liver was first described by Claude Bernard in 18559, and is still used in various scientific fields for studies of liver biology, toxicology, and pathophysiology. Advantages of the perfused liver compared to the above-mentioned in vitro models include the maintenance of the hepatic architecture, the vascular flow, the hepatocyte polarity and zonation, and the interactions between hepatocytes and non-parenchymal cells. Compared to in vivo studies, the perfused liver allows the study of liver metabolism in an isolated manner avoiding extra-hepatic factors carried by the blood and with complete control over the experimental conditions. Several modifications have been made to improve the rat liver perfusion model over the years10,11,12,13. Although mice have been used for isolated perfused liver studies, less literature is available. Here, we present a method for in situ perfusion of the mouse liver by cannulation of the portal vein and the suprahepatic vena cava inferior to study the acute and direct metabolic responses to metabolic substrates and hormones as measured in the hepatic venous effluent from the mouse liver in real-time.

<table><tbody><tr><td>3-way stopcock</td><td>BD</td><td>394601</td><td></td></tr><tr><td>Altromin breeding diet</td><td>Altromin Spezialfutter</td><td>1319</td><td></td></tr><tr><td>Calcium chloride dihydrate (CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;#183;2H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma</td><td>C8106</td><td></td></tr><tr><td>Catheters (0.7 mm)</td><td>BD</td><td>381812</td><td></td></tr><tr><td>Filter paper (pore size 2.0 &amp;#181;m)</td><td>Millipore</td><td>AP2029325</td><td></td></tr><tr><td>Glucose kit</td><td>QuantiChromTM</td><td>DIGL-100</td><td>Low concentration protocol</td></tr><tr><td>Ketamine</td><td>MSD Animal Health</td><td>511485</td><td>90 mg/kg</td></tr><tr><td>Ligature (black sterile silk)</td><td>Agnthos</td><td>14739</td><td></td></tr><tr><td>Magnesium sulfate (MgSO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma</td><td>230391</td><td></td></tr><tr><td>Non-esterified fatty acids kit</td><td>Fujifilm Wako Chemicals</td><td>NEFA-HR(2)</td><td></td></tr><tr><td>Operating table, heated on tripod stand, type 873</td><td>Harvard Bioscience, Inc.</td><td>733776</td><td></td></tr><tr><td>Potassium chloride (KCl)</td><td>Sigma</td><td>P9541</td><td></td></tr><tr><td>Potassium dihydrogen phosphate(KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Merck</td><td>1.04877</td><td></td></tr><tr><td>Roller Pump, with four channels</td><td>Harvard Bioscience, Inc.</td><td>730100</td><td></td></tr><tr><td>Sleek tape</td><td>Mediq danmark</td><td>4001910</td><td></td></tr><tr><td>Sodium bicarbonate (NaHCO&lt;sub&gt;3&lt;/sub&gt;)</td><td>Sigma</td><td>S5761</td><td></td></tr><tr><td>Sodium chloride (NaCl)</td><td>Sigma</td><td>S1679</td><td></td></tr><tr><td>Thermostatic Circulator</td><td>Harvard Bioscience, Inc.</td><td>730125</td><td>Bath Volume 3 L, 230 V/50 Hz</td></tr><tr><td>Tubing</td><td>Tygon</td><td>E3603</td><td>Inner diameter 1.59 mm, outer diameter 3.18 mm</td></tr><tr><td>Universal perfusion system</td><td>Harvard Bioscience, Inc.</td><td>732316</td><td>Basic unit uniper UP-100, type 834</td></tr><tr><td>Vamin</td><td>Fresenius Kabi</td><td>B05ABA01</td><td>Mixed amino acids</td></tr><tr><td>Vessel clamp adaptor</td><td>Deutsche Biomedical</td><td>DBC1002</td><td></td></tr><tr><td>Vessel clamps</td><td>Deutsche Biomedical</td><td>DBC1005</td><td></td></tr><tr><td>Windkessel</td><td>Harvard Bioscience, Inc.</td><td>732068</td><td></td></tr><tr><td>Xylazine</td><td>Rompun Vet</td><td>530701</td><td>10 mg/kg</td></tr></tbody></table>

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.

A steady baseline is required to determine whether a stimulus or substrate leads to the release of the molecule of interest. Figure 3A shows an example of a successful experiment. Production of urea in the perfused liver is measured in 2 min intervals and shown as mean &#177; SEM. The baseline periods preceding each of the two stimulation periods are steady. The mean urea production during the two stimulation periods and the respective preceding baselines are shown in Figure 3B. Statistical significance between the periods was tested using one-way ANOVA with repeated measurements. Based on these results, it can be concluded that the urea response to two consecutive stimulations with mixed amino acids is similar, which is an important control experiment in evaluating repeated stimulations with different doses or test compounds.
In addition to ureagenesis, it is possible to study hepatic glycogenolysis and lipolysis using the above protocol. Figure 3C-F show data from a separate experiment during an infusion with glucagon. The release of glucose (Figure 3C) and the non-esterified fatty acids (NEFA) (Figure 3E) was measured in 4 min periods. A steady basal release of glucose (Figure 3C) and NEFA (Figure 3E) is observed prior to a robust increase in glucose and NEFA during the glucagon infusion, respectively. Based on the mean values at the basal state and during the glucagon infusion, it can be concluded that glucagon rapidly stimulates hepatic glycogenolysis (Figure 3D) and lipolysis (Figure 3F).
Figure 4&#160;shows an example of apparently unsuccessful experiments. The basal release of urea is unsteady, and the stimulation periods are too close to each other for the urea production to return to basal levels before the next stimulation begins. Without steady basal periods, it is not possible to distinguish one urea response from the next. From these data, it is impossible to conclude whether different glucose concentrations influence amino acid-induced ureagenesis in the perfused mouse liver. Experimental protocols should instead be designed with 20-30 min of baseline periods in-between two consecutive stimulations to reach a steady baseline before each stimulation period.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65596/65596fig03.jpg" /> 
Figure 3: Good-quality data from the perfused mouse liver. (A,B) Urea total is shown during basal conditions and in response to periods of administration of mixed amino acids (10 mM) and glucose (6 mM). Data are presented as (A) mean &#177; SEM and (B) mean values during each stimulation and preceding basal period (each dot represents a mouse), n = 6. (C, D) Glucose and (E,F) non-esterified free fatty acids (NEFA) total outputs are shown in basal conditions and response to the administration of glucagon (10 nM). Data are presented as (C,E) mean &#177; SEM and (D,F) mean values during the basal and stimulation period (each dot represents a mouse), n = 6. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65596/65596fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65596/65596fig04.jpg" /> 
Figure 4: Poor-quality data from the perfused mouse liver. Urea total output is shown in basal conditions in response to mixed amino acids (10 mM) and decreasing concentrations of glucose (18 mM, 12 mM, 6 mM, 3 mM, and 0 mM). Data are presented as mean &#177; SEM, n = 3. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65596/65596fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Time-dependent partial pressure of oxygen and carbon dioxide and pH in Krebs-Henseleit bicarbonate buffer during gassing with 95% O2 + 5% CO2. (A) Partial pressure of oxygen, (B) carbon dioxide, and (C) pH in samples of Krebs-Henseleit perfusion buffer (pH not adjusted prior to the experiment) collected after having run through the perfusion system at indicated time points after the start of the gassing with 95% O2 + 5% CO2. Perfusion buffer samples were measured with a blood-gas analyzer. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65596/Supplementary_Figure_1.pdf" target="_blank">Please click here to download this figure.</a>
Supplementary Figure 2: Changes in partial pressure of oxygen and carbon dioxide and pH across the perfused liver. (A) Partial pressure of oxygen, (B) carbon dioxide, and (C) pH in samples of Krebs-Henseleit perfusion buffer collected immediately before reaching the liver and after being passed the perfused liver at 1 min and 180 min into an experiment. Perfusion samples were measured with a blood-gas analyzer. Data is shown as mean &#177; SD. **P &#60; 0.01, ****P &#60; 0.0001 by multiple paired t-tests corrected for multiple testing using the Holm-Sidak method. n =14. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65596/Supplementary_Figure_2.pdf" target="_blank">Please click here to download this figure.</a>

Watch this Scientific Journal Video about Investigating Hepatic Adaptations and Prediabetic Progression in Liver Diseases at JoVE.com

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.

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 ...

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1.7K Views.  University of Copenhagen. 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.

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

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 ...

Biochemistry

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 ...

Extracellular Glucose Depletion as an Indirect Measure of Glucose Uptake in Cells and Tissues Ex Vivo

The ongoing worldwide epidemic of diabetes increases the demand for the identification of environmental, nutritional, endocrine, genetic, and epigenetic factors affecting glucose uptake. The measurement of intracellular fluorescence is a widely used method to test the uptake of fluorescently-labeled glucose (FD-glucose) in cells in vitro, or for imaging glucose-consuming tissues in vivo. This assay assesses glucose uptake at a chosen time point. The intracellular analysis assumes that the metabolism of FD-glucose is slower than that of endogenous glucose, which participates in catabolic and anabolic reactions and signaling. However, dynamic glucose metabolism also alters uptake mechanisms, which would require kinetic measurements of glucose uptake in response to different factors. This article describes a method for measuring extracellular FD-glucose depletion and validates its correlation with intracellular FD-glucose uptake in cells and tissues ex vivo. Extracellular glucose depletion may be potentially applicable for high-throughput kinetic and dose-dependent studies, as well as identifying compounds with glycemic activity and their tissue-specific effects.

Extracellular Glucose Depletion as an Indirect Measure of Glucose Uptake in Cells and Tissues Ex Vivo

Extracellular depletion of fluorescently labeled glucose correlates with glucose uptake and could be used for high-throughput screening of glucose uptake in excised organs and cell cultures.

The ongoing worldwide epidemic of diabetes increases the demand for the identification of environmental, nutritional, endocrine, genetic, and epigenetic ...

Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Establishing a system of procedures to qualitatively and quantitatively characterize in vivo and in vitro hepatic steatosis is important for metabolic study in the liver. Here, numerous assays are described to comprehensively measure the phenotype and parameters of hepatic steatosis in mouse and hepatocyte models. 
Combining the physiological, histological, and biochemical methods, this system can be used to assess the progress of hepatic steatosis. In vivo, the measurements of body weight and nuclear magnetic resonance (NMR) provide a general understanding of mice in a non-invasive manner. Hematoxylin and Eosin (H&amp;E) and Oil Red O staining determine the histological morphology and lipid deposition of liver tissue under nutrient overload conditions, such as high-fat diet feeding. Next, the total lipid contents are isolated by chloroform/methanol extraction, which are followed by a biochemical analysis for triglyceride and cholesterol. Moreover, mouse primary hepatocytes are treated with high glucose plus insulin to stimulate lipid accumulation, an efficient in vitro model to mimic diet-induced hyperglycemia and hyperinsulinemia in vivo. Then, the lipid deposition is measured by Oil Red O staining and chloroform/methanol extraction. Oil Red O staining determines intuitive hepatic steatotic phenotypes, while the lipid extraction analysis determines the parameters that can be analyzed statistically. The present protocols are of interest to scientists in the fields of fatty liver diseases, insulin resistance, and type 2 diabetes.

Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Here, optimized methods to generate in vivo and in vitro models of hepatic steatosis and to analyze the steatotic phenotypes and related physiological parameters are described.

Establishing a system of procedures to qualitatively and quantitatively characterize in vivo and in vitro hepatic steatosis is important for metabolic ...

Determination of Fatty Acid Oxidation and Lipogenesis in Mouse Primary Hepatocytes

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid oxidation, or alternatively, synthesize new lipid via de novo lipogenesis. The net sum of these pathways is dictated by a number of factors, which in certain disease states leads to fatty liver disease. Excess hepatic lipid accumulation is associated with whole body insulin resistance and coronary heart disease. Tools to study lipid metabolism in hepatocytes are useful to understand the role of hepatic lipid metabolism in certain metabolic disorders.
In the liver, hepatocytes regulate the breakdown and synthesis of fatty acids via &#946;-fatty oxidation and de novo lipogenesis, respectively. Quantifying metabolism in these pathways provides insight into hepatic lipid handling. Unlike in vitro quantification, using primary hepatocytes, making measurements in vivo is technically challenging and resource intensive. Hence, quantifying &#946;-fatty acid oxidation and de novo lipogenesis in cultured mouse hepatocytes provides a straight forward method to assess hepatocyte lipid handling. 
Here we describe a method for the isolation of primary mouse hepatocytes, and we demonstrate quantification of &#946;-fatty acid oxidation and de novo lipogenesis, using radiolabeled substrates.

De novo lipogenesis and &#946;-fatty acid oxidation constitute key metabolic pathways in hepatocyte, pathways that are perturbed in several metabolic disorders, including fatty liver disease. Here we demonstrate isolation of mouse primary hepatocytes and describe quantification of &#946;-fatty acid oxidation and lipogenesis.

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid ...

Biology

Evaluation of Hepatic Glucose Production in a Polycystic Ovary Syndrome Mouse Model

Polycystic ovary syndrome (PCOS) is a common disease that results in disorders of glucose metabolism, such as insulin resistance and glucose intolerance. Dysregulated glucose metabolism is an important manifestation of the disease and is the key to its pathogenesis. Therefore, studies involving evaluation of glucose metabolism in PCOS are of utmost importance. Very few studies have quantified hepatic glucose production directly in PCOS models using non-radioactive glucose tracers. In this study, we discuss step-by-step instructions for the quantification of the rate of hepatic glucose production in a PCOS mouse model by measuring M+2 enrichment of [6,6-2H2]glucose, a stable isotopic glucose tracer, via gas chromatography - mass spectrometry (GCMS). This procedure involves creation of stable isotopic glucose tracer solution, use of tail vein catheter placement and infusion of the glucose tracer in both fasting and glucose-rich states in the same mouse in tandem. The enrichment of [6,6-2H2]glucose is measured using pentaacetate derivative in GCMS. This technique can be applied to a wide variety of studies involving direct measurement of the rate of hepatic glucose production.

This study describes the direct measurement of hepatic glucose production in a polycystic ovary syndrome mouse model by using a stable isotopic glucose tracer via tail vein in both fasting and glucose-rich states in tandem.

Polycystic ovary syndrome (PCOS) is a common disease that results in disorders of glucose metabolism, such as insulin resistance and glucose intolerance. ...

An Improved Time- and Labor- Efficient Protocol for Mouse Primary Hepatocyte Isolation

Primary hepatocytes are used extensively in liver in vitro research, especially in glucose metabolism studies. A base technique has been adapted based on different needs, like time, labor, cost, and primary hepatocyte usage, resulting in various primary hepatocyte isolation protocols. However, the numerous steps and time-consuming reagent preparations in primary hepatocyte isolation are major drawbacks for efficiency. After comparing different protocols for their pros and cons, the advantages of each were combined, and a rapid and efficient primary hepatocyte isolation protocol was formulated. Within only ~35 min, this protocol could yield as much, if not more, healthy primary hepatocytes as other protocols. Further, glucose metabolism experiments performed using the isolated primary hepatocytes validated the usefulness of this protocol in in vitro liver metabolism studies. We also extensively reviewed and analyzed the significance and purpose of each step in this study so that future researchers can further optimize this protocol based on needs.

Primary hepatocytes are a valuable tool to study liver response and metabolism in vitro. Utilizing commercially available reagents, an improved time- and labor-efficient protocol for mouse primary hepatocyte isolation was developed.

Primary hepatocytes are used extensively in liver in vitro research, especially in glucose metabolism studies. A base technique has been adapted based on ...

Ultrafast Doppler Ultrasound for Hepatic Microvasculature Analysis in Mice

Imaging and Quantification of the Hepatic Vasculature of Mice Using Ultrafast Doppler Ultrasound

Non-invasive in vivo imaging of the vasculature is a powerful tool for studying disease mechanisms in rodents. To achieve high sensitivity imaging of the microvasculature using Doppler ultrasound methods, imaging modalities employing the concept of ultrafast imaging are preferred. By increasing the frame rate of the ultrasound scanner to thousands of frames per second, it becomes possible to improve the sensitivity of the blood flow down to 2 mm/s and to obtain functional information about the microcirculation in comparison to a sensitivity of around 1 cm/s in conventional Doppler modes. While Ultrafast Doppler ultrasound (UFUS) imaging has become adopted in neuroscience, where it can capture brain activity through neurovascular coupling, it presents greater challenges when imaging the vasculature of abdominal organs due to larger motions linked to breathing. The liver, positioned anatomically under the diaphragm, is particularly susceptible to out-of-plane movement and oscillating respiratory motion. These artifacts not only adversely affect Doppler imaging but also complicate the anatomical analysis of vascular structures and the computation of vascular parameters. Here, we present a qualitative and quantitative imaging analysis of the hepatic vasculature in mice by UFUS. We identify major anatomical vascular structures and provide graphical illustrations of the hepatic macroscopical anatomy, comparing it to an in-depth anatomical assessment of the hepatic vasculature based on Doppler readouts. Additionally, we have developed a quantification protocol for robust measurements of hepatic blood volume of the microvasculature over time. To contemplate further research, qualitative vascular analysis provides a comprehensive overview and suggests a standardized terminology for researchers working with mouse models of liver disease. Furthermore, it offers the opportunity to apply ultrasound as a non-invasive complementary method to inspect hepatic vascular defects in vivo and measure functional microvascular alterations deep within the organ before unraveling blood vessel anomalies at the micron scale levels using ex vivo staining on tissue sections.

Ultrafast Doppler ultrasound (UFUS) with high spatial resolution (100 &#956;m) and sensitivity represents a suitable noninvasive imaging modality for obtaining a comprehensive qualitative overview of the hepatic vasculature. UFUS also facilitates quantitative measurements of the microvasculature, aiming to enhance our understanding of vascular disease mechanisms.

Non-invasive in vivo imaging of the vasculature is a powerful tool for studying disease mechanisms in rodents. To achieve high sensitivity imaging of the ...

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.

Hepatic insulin clearance is essential for maintaining glucose homeostasis and is closely linked to metabolic disorders such as obesity, insulin ...

Measurement of Extracellular Fluorescent Glucose Analog Depletion: A Surrogate Measurement Technique to Assess Glucose Uptake in Mouse Organs Ex Vivo

Source: Kumar, S. B. et al., <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/63681">Extracellular Glucose Depletion as an Indirect Measure of Glucose Uptake in Cells and Tissues Ex Vivo.</a>&#160;J. Vis. Exp.&#160;(2022)
In this video, we demonstrate an ex vivo technique to measure extracellular glucose analog depletion in harvested mouse organs. It involves incubating the mouse tissue in a solution containing fluorescently-labeled deoxyglucose, which is taken up by the cells. The glucose analog uptake can be indirectly determined by measuring the fluorescence of the incubated medium.

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

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Chapters in this video

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

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Extracellular Glucose Depletion as an Indirect Measure of Glucose Uptake in Cells and Tissues Ex Vivo

Optimized Analysis of In Vivo and In Vitro Hepatic Steatosis

Determination of Fatty Acid Oxidation and Lipogenesis in Mouse Primary Hepatocytes

Evaluation of Hepatic Glucose Production in a Polycystic Ovary Syndrome Mouse Model

An Improved Time- and Labor- Efficient Protocol for Mouse Primary Hepatocyte Isolation

Imaging and Quantification of the Hepatic Vasculature of Mice Using Ultrafast Doppler Ultrasound

Assessing Insulin Clearance in Mice via In Situ Liver Perfusion

Measurement of Extracellular Fluorescent Glucose Analog Depletion: A Surrogate Measurement Technique to Assess Glucose Uptake in Mouse Organs Ex Vivo