Intravital imaging of hepatic blood biliary barrier in live mice

Abstract

Understanding the kinetics and spatiotemporal interactions of living cells within the tissue environment requires real time imaging. The introduction of two-photon microscopy has substantially boosted the power of live intravital imaging, making it possible to obtain information of individual cells in near physiological conditions within intact tissues non-destructively. Intravital imaging of the liver has proved useful in understanding its 3D structure, function, and dynamic cellular interactions. Recently we have shown that integrity of blood bile barrier in different physiological and pathophysiological conditions can be imaged in real time using intravital microscopy. We here discuss the real time intravital imaging method to visualize blood bile barrier integrity in the murine liver.

INTRODUCTION

End-point assays, which provide a snapshot image of biological processes, make up the majority of analytical methods used to analyze experimental systems. While these assays can provide a wealth of information, it is frequently necessary to visualize the dynamics and spatiotemporal interactions of cells and their environment in real time. Intravital live imaging has emerged as one of the powerful tools to image physiological processes in real time(Bennewitz et al., 2014; Weigert et al., 2013). It has been routinely applied to many tissues including the brain(De Niz et al., 2019; Ricard & Debarbieux, 2014), cremaster(Donndorf et al., 2013), lung(Bennewitz et al., 2014; Brzoska et al., 2020; Yipp et al., 2017), liver(Pradhan-Sundd et al., 2018b), kidney(Hato et al., 2018; Peti-Peterdi et al., 2016) and lymph nodes(Meijer et al., 2017; Miller et al., 2002). Liver intravital imaging is an emerging field which has been instrumental in understanding three dimensional dynamics of liver tissue including immune environment(Liew & Kubes, 2016; Matsumoto et al., 2018), dynamics of the tumor microenvironment(Babes & Kubes, 2016), barrier function and pathological manifestation of liver disease progression(Pradhan-Sundd et al., 2018; Pradhan-Sundd et al., 2018b; Vats et al., 2020). We have recently shown that intravital imaging can be especially useful in understanding the blood bile barrier (BBlB)(Pradhan-Sundd et al., 2018; Pradhan-Sundd et al., 2018b; Pradhan-Sundd & Monga, 2019b) integrity in different physiological and pathophysiological conditions in real time, which we discuss in detail in this methods article here.

Basic Protocol 1 detail the methods for surgery and microscope set up for intravital imaging in the mouse liver. This protocol relies on image stabilization with an abdominal/liver suction window. Basic Protocol 2 provides the methods needed to image blood bile barrier integrity in the liver of a live mouse. Both the protocols require familiarity with mouse surgery and advanced microscopy.

NOTE: All protocols involving live animals must first be evaluated and approved by an Institutional Animal Care and Use Committee (IACUC), and must adhere to officially authorized regulations for laboratory animal care and use. (IACUC)

BASIC PROTOCOL I

The quantitative liver intravital imaging (qLIM) method described here is adopted from (Bennewitz et al., 2014; Brzoska et al., 2020). The detailed methods of surgery, anesthesia and image capturing as well as materials required for these procedures are described in this section.

LIVE IMAGING IN THE MOUSE LIVER

Materials

Mice (procedure is best performed in adult mice >20 g and >30 g in body weight)

Anesthetics: ketamine, xylazine, isoflurane

Compressible gases (21%, 100% oxygen)

Phosphate-buffered saline (PBS)/sodium chloride 0.9%

Alcohol swabs

Adhesive tape

Absorbent spears

Thor-labs stage connector for clamp connection

Rectal thermometer

Portable Oximeter with blood pressure monitor

Vetbond tissue glue (3M)

Cotton swabs

Suture: 3–0 silk

Fiber optic illuminator (Cole Parmer) for surgery

Surgical tools for abdominotomy and cannulation procedure

Thermal cautery instrument (e.g., Geiger Medical Technologies Inc)

PE-10 and PE-90 tubing (Intramedic)

Heparin

Isoflurane vaporizer (Molecular Imaging Products, cat. no. AS-01–0007; http://www.mipcompany.com/)

Customized abdominal suction window (see annotation to step 14, below)

Micromanipulator (Thorlabs; http://www.thorlabs.com)

Suction regulator and tubing

12-mm glass coverslips

High vacuum grease (e.g., Dow Corning)

Hypromellose (0.3%) lubricant eye gel (e.g., GenTeal Tears, Alcon Laboratories, Inc.)

Two-photon microscope with motorized and heated stage

Additional reagents and equipment for intraperitoneal injection of mice(Donovan & Brown, 2006).

Anesthetizing the mouse

1. Anesthetize mouse using intraperitoneal injection (Donovan & Brown, 2006)of ketamine (100 mg/kg) and xylazine (20 mg/kg).

2. Place the mouse on a heated microscope stage to obtain a rectal temperature of approx. 37 degrees Celsius.

3. Tape the mouse's limbs to the heated stage and place it in a supine posture. To flatten the neck and head, wrap 3–0 silk suture around the incisors and tape it to the stage. Light up the operation area with a fiber-optic illuminator.

4. After obtaining appropriate anesthetic (confirmed by paw pinch), dissect the trachea with small scissors and forceps, then make a short horizontal incision in the anterior trachea with fine scissors. PE-90 tubing with a beveled edge should be inserted into the trachea and secured with Vetbond or 3–0 silk suture.

5. At this step, cannulation of the carotid artery can be performed using PE-10 heparinized tubing. To perform cannulation, first remove skin from the front part of the neck while using sharp forceps and scissors to expose the right carotid artery. The blood flow is stopped using silk suture from the head side and a clamp. Afterwards, an incision into the carotid artery is made and PE-10 tubing is inserted and knotted using silk suture. Cannulation is essential when any substance needs to be administrated during the intravital session.

Surgery and placement of the liver window

• 6.Reposition the mouse to the supine position. Position a capped needle or similar sized instrument under the front shoulders to elevate the mouse's right abdominal cavity. Reattach the mouse to the heated stage with new tape.
• 7.Give a pre-warmed physiological saline i.e. injection(Donovan & Brown, 2006) (0.5 to 1.0 ml based on the body weight).

Note: The purpose of this injection is to keep the mouse hydrated and restore surgical and insensible blood loss that might occur during the experiment. At 2 hours, a repeat injection of 0.5 mL saline can be administered, followed by hourly injections.

• 8.Using a paw pinch, reassess the level of anesthetic every 15 minutes.
• 9.Wet the mouse fur overlaying the right abdominal cavity with an alcohol swab. Alternatively, you can shave the mouse fur using little clippers.
• 10.Remove mouse fur and underlying subcutaneous tissue from the right abdominal cavity using fine scissors and forceps. Dissect carefully until the liver lobes are visible. Any bleeding vessels should be cauterized with adjusted heat.
• 11.Attach the liver suction window (Figure. 1 and 2) to the strut and adjust the height and horizontal plane with a micromanipulator. Seal a 12-mm glass coverslip onto the suction window using vacuum grease.

Figure. 1. Schema for liver intravital microscopy.

(A) The overall design of the quantitative liver intravital imaging (qLIM) is shown here. A catheter is inserted into the carotid artery to allow intravascular administration of fluorescent antibodies. The mouse is anesthetized as described in basic protocol I. A heated stage and a temperature-controlled enclosure surround the qLIM microscope stage to keep the mouse’s temperature stable. To immobilize a small section of the right lobe of the liver against a cover glass, gentle vacuum suction is given to the liver window device. A NIR laser is used to accomplish qLIM imaging. Excitation fluorescence is represented by red in the objective, while emission fluorescence is represented by green. A pressure transducer is linked to the carotid artery catheter to monitor blood pressure during imaging. Blood oxygen saturation, breath rate, breath distension, heart rate, and pulse distention are all monitored with pulse oximetry.

Figure 2: Detailed steps involved in quantitative liver intravital imaging.

(A) General anesthetic procedure using i.p. injections of ketamine (100 mg/kg) and xylazine (20 mg/kg) cocktail with an optional addition of atropine. Mouse needs to be under investigator’s supervision to monitor the anesthetic progress. (B) Anesthetized mouse is placed supinely on heat pad with the temperature of approximately 37°C and becomes immobilized by taping. The neck and head are flatten using 3–0 silk suture and the operation area is lightened up with a fiber-optic illuminator. (C-E). Skin is removed from front part of a neck and using sharp forceps and scissors right carotid artery is exposed. The blood flow is stopped using silk suture from the head side and clamp. Precisely made incision in the artery allows to insert PE-10 heparinized tubing and release the clamp. Tubing must be secured by double knot using 3–0 silk suture. Trachea is exposed and incision between 4^th and 5^th tracheal cartilages is made using ultra fine scissors and tracheal junction is inserted into the created window. PE-90 tubing connected to mini-vent station with silicone tubing needs to be inserted into the window and immobilized using tissue bond glue. (F-G). The procedure of exposing the right liver lobe is done by removing skin and surrounding tissues by gentle incisions followed by cauterization of the bleeding sites. After exposing the lobe, clamp with 12mm cover glass is horizontally placed 5 mm above the lobe being ready for the suction. The suction is made by putting the clamp to the lobe under the adjusted pressure in one-movement manner. After suction the clamp is lifted by 2–3 mm for better manipulation of the objective focus. (H) Mouse supinely placed on the heat pad is moved to the microscope site.

Note: The right lobe of the liver was chosen for imaging. Also, to avoid touching the surface of the liver with surgical instruments, extreme caution must be utilized.

• 12.Fill the liver window with 20 to 25 mmHg suction and lower it onto the right lobe of the liver; the liver will then enter the abdominal window and be stabilized for imaging. Administer lubricant eye gel to generate a meniscus between the suction window coverslip and the microscope objective by lowering the two-photon microscope objective to just above the glass coverslip.

Note: As little suction as possible should be used to allow the liver to enter the liver window. The suction window must be totally horizontal (i.e., imaging through a tilted coverslip) to avoid tangential imaging.

• 13.Scan to the deepest liver depth using z-steps (up to 50 z steps) from the top of the right liver lobe.
• 14.Adjust laser power, excitation laser wavelength, photomultiplier tubes (detector) voltage/gain/offset, and appropriate filter cubes for two-photon microscopy to optimally excite and collect images for the fluorophore of choice (Figure 1 and 2 exhibits the steps shown in basic protocol I).

SUPPORT PROTOCOL 1

MONITORING VITAL SIGNS OF THE MOUSE DURING LIVE LIVER IMAGING

Laboratory mice have physiological traits such as reduced respiratory volumes and fast heart rates that distinguish them from other animals (Hedrich & Bullock, 2004). For maintaining the animal’s safety, evaluating the degree of anesthesia, and keeping the physiological relevance of the procedure, accurate and continuous monitoring of vital signs such as heart rate, blood pressure, body temperature, and blood oxygen saturation is necessary.

Core body temperature

1. Rectal thermometers have traditionally been the most extensively employed in mouse liver intravital imaging for determining core body temperature.

Note: A two-photon microscope with a thermostat-controlled heated stage and an air-heated imaging chamber allows the mouse core body temperature to be maintained during the imaging session.

Heart rate and arterial blood pressure

• 2.Noninvasive tail cuff systems or fluid-filled catheter systems that can be linked to an arterial catheter can be used to monitor hemodynamic parameters such as heart rate and arterial blood pressure.

Note: For more information on arterial pressure monitoring in mice, see Zhao et al’s overview(Zhao et al., 2011).

Blood oxygen saturation

• 3.Pulse oximetry, a simple non-invasive method for monitoring the proportion of oxygen-saturated hemoglobin in blood, can be used to detect blood oxygen saturation in mice during intravital liver imaging.

Note: During intravital liver imaging, commercially available sensors developed for mouse various body regions (neck, thigh, and foot) allow for simple monitoring of blood oxygen saturation.

BASIC PROTOCOL 2

VISUALIZATION OF BLOOD AND BILE TRANSPORT USING INTRAVITAL MICROSCOPY

The liver performs a wide variety of metabolic and regulatory functions. One of the key functions of the liver is bile formation secretion and transport(Alrefai & Gill, 2007; Boyer, 2013; Cai & Boyer, 2021). This is achieved by the active transport of bile acids synthesized in the hepatocytes and captured from the sinusoidal blood across hepatocytes into bile canaliculi (Figure 3A). In the adult liver, hepatocytes have three distinct membrane domains: sinusoidal/basal, lateral, and canalicular/apical. The lateral plasma membrane domains of hepatocytes fuse alongside the bile canaliculi to form tight junctions (zonula occludens) that separate the apical/canalicular domain from the sinusoidal/basal surface by forming the blood bile barrier (BBlB) (Figure 3A)(Pradhan-Sundd & Monga, 2019a). Loss of BBlB has been associated with impaired liver development and disease (Pradhan-Sundd & Monga, 2019b). However, the direct visualization of the BBlB was not achieved due to the lack of live imaging tools.

Figure 3: qLIM enables stable visualization of blood and bile flow in murine liver.

(A) Schematic showing the blood and bile flow in the liver. (B-B”) qLIM imaging showing the real-time localization of Texas Red–dextran (TXR–dextran) and carboxyfluorescein (CF) in hepatic sinusoidal blood vessel and bile duct of the liver of control mouse respectively. TXR-Dextran and CF shows complete exclusion at any given time point in a control liver. (C-C”) Real time localization of CF at three different time point shows its trafficking from hepatocyte to bile canaliculi within 1–2 minutes. CF intensity significantly reduces at around 30 minutes due to degradation.

Recently, we have shown that carboxyflurescein (CF) can be used to analyze the bile trafficking in using different models of acute and chronic liver injury(Pradhan-Sundd et al., 2017, 2018b). After being internalized by hepatocytes, carboxyflurescein diacetate (CFDA) gets hydrolyzed by esterase into fluorogenic CF, which emits green fluorescence at 517nm (Liu et al., 2011). In WT, the uptake of CFDA and hydrolysis into CF occurs within 1 minute after injection (Figure 3B-C”). In liver injury models with impaired blood bile barrier, the movement of CF to hepatocyte and eventually to the bile canaliculi gets affected. Thus, CF can be used as a surrogate marker of bile flow(Pradhan-Sundd et al., 2018b; Vats et al., 2020). Here, we describe how to acquire, analyze and quantify the transport of CF from blood to hepatocyte, and finally to bile canaliculi as well as blood flow through hepatic sinusoids. Follow the Basic Protocol 1 to successfully perform mouse thoracotomy and place imaging window on the right lobe of the liver. Additional materials and methods are described below.

Materials

See Basic Protocol 1

Carboxyflurescein diacetate (Sigma, Cat #21879), (Molecular weight 377)

Texas red dextran (TXR-dextran) (Thermo Fisher, Cat#D1830), (MW 70,000)

Image analysis software

Note: Image analysis software can be purchased commercially (NIS-elements/ Olympus) or obtained for free via freeware (e.g., ImageJ, a free image-processing tool from the NIH with an extensive library of plug-ins; http://rsb.info.nih.gov/ij/). Image analysis can also be conducted using image processing tool box in MATLAB (MathWorks Inc).

Protocol steps and annotations:

1. Follow the methods described in basic protocol I to prepare the mouse for surgery and intravital imaging (Figure 1, 2).

2. Inject the intravascular fluorescent dyes through the carotid artery catheter (Fig 1). Intravascular fluorescent dyes used in this assay are 200 μg of Texas Red dextran, or 100 μg of Carboxyflurescein (CF). Texas red dextran is used to visualize the blood flow through the liver sinusoids whereas carboxyflurescein (Molecular weight 377)(Massou et al., 2000) is used to visualize bile canaliculi. Other dyes might be used for staining different cell types as described here(Brzoska et al., 2020).

3. Perform microscopy using two-photon excitation microscope (Figure 2H).

Image Acquisition

• 4.Acquire video/ images every 30 secs till up to 20–30 minutes.

Note: In WT mice, the uptake of CF in hepatocytes occurs within 1 minute after injection (Figure 2B, supplemental video S1) and CF appears as linear green pattern outlining its secretion into biliary canaliculi through the apical surface (Figure 2B’). At around 2 min, CF was localized prominently to biliary canaliculi having been secreted by the hepatocytes(Figure 2C-C’, supplemental video S2). For around 20 minutes CF shows predominant bile canalicular localization (Figure 2D supplemental video S3). Movies were processed using Nikon’s NIS Elements software. By 30 min, CF decreased and started to disappear from the biliary canaliculi suggesting its clearance from the biliary compartment. The sinusoidal vessels containing TXR-dextran were visible as red fluorescence (Figure 2B-D red channel).

Image Processing

• 5.Perform spectral unmixing is using NIS Elements (or equivalent) to separate out tissue autofluorescence from the FITC fluorescence and to reduce bleed through among different channels of bicolor images.
• 6.Apply a median filter with a kernel size of 3 over each video frame to improve signal-to-noise ratio.
• 7.Enhance signal contrast in each channel of a multicolor image can be further by adjusting the maxima and minima of the intensity histogram of that channel. The colors of channels might be changed using the software post-experimental analysis tools. To increase the quality of the movie, noise reduction/denoising/frame rate adjustment might be performed.

Commentary

Background information

Intravital microscopy is a highly effective tool for imaging biological processes in living animals. The capacity to visualize organs in live animals at cellular and subcellular resolution has aided key discoveries in various fields such as hemostasis, metabolism, tumor biology, immunology, aging and neuroscience (Brzoska et al., 2020; Dunn & Ryan, 2017; Liew & Kubes, 2016; Matsumoto et al., 2018; Ricard & Debarbieux, 2014; Weigert et al., 2013). Interestingly, compared to imaging organs with dynamic movements (such as lung), liver imaging is relatively straight forward and does not require mechanical ventilation. For qLIM imaging we have adapted the method from (Bennewitz et al., 2014; Brzoska et al., 2020) which is primarily based on a modified approach first described by of Looney et al 2011. In this method gentle vacuum pressure is applied to stabilize a small area of the mouse organ (Looney et al., 2011) which does not alter normal perfusion and thus allows imaging at a physiological level.

Using this modified qLIM method we have successfully shown that CF and TXR-dextran can be used to not only image blood and bile trafficking in real time (as described in Basic Protocol II) but also to understand the effects of acute and chronic liver injury in hepatic BBlB regulation. (Pradhan-Sundd et al., 2017, 2018b; Vats et al., 2020; Vats et al., 2021). For example, in a control mouse liver, CF and TXR-dextran shows complete separation at any given time point of imaging (Figure 4A, supplemental video S4). However, in chronic liver injury models, CF and TXR-dextran fails to localize correctly due to the loss of BBlB integrity. As shown in Figure 4B, C and D, impaired BBlB function can appear as either complete colocalization of CF and TXR dextran in hepatic sinusoids (Figure 4B, supplemental video S5) seen in DDC diet (3,5-diethoxycarbonyl-1,4-dihydrocollidine) induced liver injury(Pradhan-Sundd et al., 2018a) or localization of CF to hepatocytes and impaired trafficking to bile canaliculi as seen in mice fed with CDE diet (choline-deficient, ethionine-supplemented) (Figure 4C, supplemental video S6). Additionally, hepatocyte damage or apoptosis can also cause mislocalization of CF and TXR-dextran as seen in Figure 4D, (supplemental video S7). We have also shown that qLIM can be useful to analyse hepatic blood circulation in hematological diseases with sinusoidal vasoocclusion (Vats et al., 2020).

Figure 4: Intravital imaging of blood bile barrier in steady state and during liver injury.

(A) Real time imaging of control mouse liver shows completely exclusive localization TXR-dextran and CF in the hepatic sinusoids and bile duct respectively. In a control mouse CF travels from hepatic sinusoids to hepatocytes and then to bile canaliculi. Whereas TXR-dextran flows through the hepatic sinusoids. (B-D) Mislocalization of CF can be used to measure blood bile barrier integrity in models of liver injury. Impaired localization of (B) CF from blood to hepatocyte, (C) hepatocyte to bile duct and (D) complete loss of BBlB integrity with mislocalization and leakage of both CF and TXR-dextran can be seen utilizing intravital imaging in mouse liver indicative of BBlB defects.

Critical Parameters and troubleshooting for qLIM

Mouse surgery and anesthesia

Surgical preparation is the key to optimal liver imaging. Great care must be taken to avoid touching the surface of the liver during the surgery and placement of the liver suction window. After the liver window is placed, it is difficult to remove and replace without causing any tissue injury.

Depth of anesthesia is also a critical parameter. Mice must be deeply anesthetized with ketamine+ xylazine+ isoflurane during the imaging to obtain quality movies. This can be periodically assessed with a paw pinch, and the isoflurane flow rate can be adjusted.

Microscope set-up

The multitude of biological questions that can be answered by live imaging requires a variety of different imaging set-ups to acquire the desired data. In order to test how blood is circulating through the vasculature, video-rate imaging is required to capture the events of interest. In contrast, imaging CF trafficking may require a timepoint every 30 sec. These considerations should be considered when choosing a microscope and again during acquisition.

The laser power that reaches the sample has the ability to easily cause injury to the liver tissue. It is therefore critical to have the appropriate settings in place before beginning an experiment. The amount of laser light that reaches the sample, the number of frames averaged, and photomultiplier tube (PMT) gain should all be optimized to allow for the best signal-to-noise ratio while keeping the laser power and dwell time to a minimum. Similarly, to avoid photobleaching liver tissue should not be overexposed to laser for a prolonged time period. Ambient light should be carefully screened from the microscope and PMTs. For in vivo imaging, monitoring of the mouse should be done in between timepoints and with as little intervention as possible.

Liver tissue viability while imaging

In intravital liver imaging, each preparation should be tested for viability at the beginning of an experiment. Dextran leak can be used as measures of tissue damage/permeability. 70-kD dextran should not leak from a good surgical preparation. When imaging over an extended period of time, it is important to periodically assess the health of the tissue. If cells slow down or do not move during an imaging session, the imaging set-up should be checked for any of these three critical parameters.

Image Acquisition and Analysis

Live mouse liver imaging experiments require a careful planning and attention to details. Experimental design should take into consideration the type of data to be acquired, how to be acquired and how the data has to be analyzed. Since, intravital imaging generates enormous amount of data, processing power and memory capacity of the analysis platform are crucial for data acquisition and analysis.

Time Considerations

Set-up of suction apparatus, and microscope should take approximately 20 min. Anesthesia, tracheotomy, liver surgery, and placement of the liver suction window should take an additional 30 min. Image acquisition can proceed for several hours, depending on the experimental conditions.

Supplementary Material

vS1

Video Legends:

Movie S1. Visualization of blood and bile trafficking in a control mouse after IV administration of Texas red dextran and CF at 1-minute post administration. The sinusoids in control mouse liver visualized by IV injection of Texas red dextran (red). Hepatocytes and biliary canaliculi are enriched in CF (green). Texas Red Dextran and CF localization appeared to be mutually exclusive at this time point. Original acquisition rate.

vS2

Movie S2. Time series video exhibiting the flow of Texas red dextran and CF through liver sinusoids and bile canaliculi at 2 min post administration in a control mouse. Intravital imaging of a control mouse liver 2 minutes after IV injection of CFDA (green) and Texas red dextran (red). The biliary canaliculi are nicely outlined with CF (green). Texas Red Dextran and CF localization is mutually exclusive at this time. Original acquisition rate.

vS3

Movie S3. Visualization of blood and bile flow in a control liver through liver sinusoids and bile canaliculi using texas red dextran and CF respectively at 20 minutes post administration. Normal liver sinusoids in a control mouse visualized by IV injection of Texas red dextran (red). The bile ducts are visualized with CF (green). CF fluorescence has considerably weakened at this time point as CF-containing bile is emptied from the liver. Original acquisition rate.

vS4

Movie S4. Visualization of blood and bile trafficking in a control mouse after IV administration of Texas red dextran and CF 1 minute prior to imaging. The sinusoids in control mouse liver visualized by IV injection of Texas red dextran (red). Hepatocytes and biliary canaliculi are enriched in CF (green). Texas Red Dextran and CF localization appeared to be mutually exclusive at this time point. Original acquisition rate.

vS5

Movie S5. Visualization of blood and bile trafficking in mice with liver injury induced by DDC diet for 4 days after IV administration of Texas red dextran and CF. DDC diet fed mouse liver sinusoids visualized by IV injection of Texas red dextran (red). Substantial co-localization of CF (green) and Texas red dextran (red) is visible in the hepatic sinusoids. CF (green) fluorescence is absent from both hepatocytes and biliary canaliculi. Original acquisition rate.

vS6

Movie S6. Visualization of blood and bile trafficking in mice with liver injury induced by DDC diet for 6 days after IV administration of Texas red dextran and CF. CDC diet fed mouse liver sinusoids visualized by IV injection of Texas red dextran (red). CF localizes to hepatocytes whereas texas red dextran is confined to sinusoids. Almost no co-localization of CF (green) and Texas red dextran (red) is visible. CF (green) fluorescence is absent from biliary canaliculi. Original acquisition rate.

vS7

Movie S7. Visualization of blood and bile flow in mice with liver injury induced by DDC diet feeding for 14 days through sinusoids and bile canaliculi. DDC diet fed mouse liver sinusoids visualized by IV injection of Texas red dextran (red). CF (green) continues to extensively co-localize with Texas red dextran (red). Some particulate accumulation is also observed in peri-sinusoidal space. Original acquisition rate.

Data availability

The data presented here will be available as per journal guidelines.