Intravital Microscopy to Study Platelet-Leukocyte-Endothelial Interactions in the Mouse Liver

Summary

Intravital microscopy is a powerful tool that provides insight into both the temporal and spatial relationships of rapid and/or sequential processes. Herein, we describe a protocol to assess both protein-protein interactions and platelet-neutrophil-endothelial interactions in liver sinusoids in a murine model of experimental sepsis (endotoxemia).

Abstract

Inflammation and thrombosis are complex processes that occur primarily in the microcirculation. Although standard histology may provide insight into the end pathway for both inflammation and thrombosis, it is not capable of showing the temporal changes that occur throughout the time course of these processes. Intravital microscopy (IVM) is the use of live-animal imaging to gain temporal insight into physiologic processes in vivo. This method is particularly powerful when assessing cellular and protein interactions within the circulation due to the rapid and sequential events that are often necessary for these interactions to occur. While IVM is an extremely powerful imaging methodology capable of viewing complex processes in vivo, there are a number of methodological factors that are important to consider when planning an IVM study. This paper outlines the process of conducting intravital imaging of the liver, identifying important considerations and potential pitfalls that may arise. Thus, this paper describes the use of IVM to study platelet-leukocyte-endothelial interactions in liver sinusoids to study the relative contributions of each in different models of acute liver injury.

Introduction

Inflammation and thrombosis are complex processes that occur primarily in the microcirculation. This protocol outlines the surgical preparation that allows for imaging of the liver microvasculature in vivo. Although standard histology may provide insight into the end pathways for both inflammation and thrombosis, it cannot show the temporal changes that occur throughout the process. Furthermore, this method is particularly powerful when assessing the transient cellular and protein interactions that occur within the microvascular circulation due to its ability to capture, via videomicroscopy, the often rapid and sequential interactions occurring within biological systems. This paper describes the use of intravital microscopy to study platelet-leukocyte-endothelial interactions in liver sinusoids to study the relative contribution of each in different models of acute liver injury.

While this surgical preparation and imaging modality have the potential to be adapted to a host of inflammatory and pathological models, two models of vascular inflammation are outlined here: an endotoxemia model of murine sepsis and acetaminophen (APAP)-induced liver injury. The injection of endotoxin (lipopolysaccharide; LPS) to induce an experimental model of murine sepsis began as early as the 1930s with its isolation and subsequent exploration as a key molecule in the progression of sepsis¹. While this model is limited in that LPS is only a single component of Gram-negative bacteria, this is a widely accepted and utilized method that is relatively simple and quick to perform. Furthermore, it rapidly and accurately replicates the physiological manifestations of sepsis¹. Given the role intestinal endotoxin absorption plays in the development of liver disease and inflammation², this is a superb model for studying platelet-leukocyte-endothelial interactions in the mouse liver.

In addition to an LPS model of sepsis, APAP overdose provides an excellent model for the study of pathological cell-cell interactions in the liver. APAP overdose can lead to acute liver failure, marked by thrombocytopenia and platelet accumulation in the liver, subsequently blocking leukocyte-mediated liver recovery³. While the surgical preparation and imaging methodology for this model can be adapted for a number of biological questions and to study various pathological processes, both the LPS model of sepsis and APAP-induced liver injury are excellent models for the study of platelet-leukocyte-endothelial interactions in vivo.

IVM has some inherent advantages over standard histological techniques. While standard histological methods allow the study of whole or sectioned tissue samples and the appreciation of proteins and tissue architecture, these methodologies come with limitations. By design, these processes require tissue processing, which has the potential to distort or mask what is found in the living system. Tissue must be fixed during histological preparation, introducing the potential for fixation artifacts or enhanced autofluorescence. Fixation can also lead to intracellular changes in tissue, and the potential for improper fixation may lead to tissue degradation. Furthermore, histologic methods lack the potential for directly studying the temporal aspects of protein or cellular interactions and have the potential of missing ephemeral or infrequent interactions.

Conversely, fluorescence intravital microscopy allows one to avoid many of the complications and limitations inherent in standard histology. IVM avoids fixation and, thereby, the artifacts or tissue degradation that can occur during histological preparation. By design, it also allows for the imaging of tissues within the living biological system, and, as such, tissue isolation and sectioning are not required. Furthermore, it allows for the imaging and study of transient or infrequent processes, which can be difficult or impossible to capture using histology. This IVM method can also be used to capture and identify sequential processes (e.g., platelet- or leukocyte-initiated interactions resulting in platelet-leukocyte aggregates bound to vascular endothelium). Finally, this method can be adapted to a host of imaging systems. Depending upon the needs of the study and the desired dataset, after surgical preparation, the externalized liver can be placed on almost any imaging system desired. We have successfully applied this protocol to imaging using widefield fluorescence microscopy, spinning disk microscopy, laser scanning confocal microscopy using a resonance head scanner, as well as two-photon microscopy. However, there is no reason to believe that this method should be limited to the aforementioned microscopy systems.

This methods paper outlines a protocol for IVM, which was used previously to study platelet-leukocyte-endothelial interactions in the mouse liver. This protocol has been used to compare temporal platelet-endothelial adhesion of platelets, either labeled with fluorescently conjugated antibodies or genetically modified for endogenous fluorescence⁴. This protocol has been used to evaluate transient platelet-leukocyte-endothelial interactions in the liver sinusoids in an endotoxemic model of liver inflammation and to co-localize P-selectin and recombinant protein. In addition, this protocol made it possible to determine whether the differences seen in neutrophil density using standard histology were a result of differences in red blood cell velocities (as a surrogate for volumetric flow rate)⁵. Finally, this method has been used to evaluate platelet recruitment by Kupffer cells in the liver sinusoids in an acute model of APAP-induced liver injury⁶. This body of work would not have been possible using standard histological methods. As stated, this protocol can be adapted for a variety of imaging systems, and, with proper surgical preparation, fluorescent antibody choice, and imaging settings, this protocol is highly reliable and reproducible for the study of liver pathology.

Protocol

All animal protocols were approved by the Institutional Animal Care and Use Committee of Baylor College of Medicine and the Research & Development Committee of the Michael E. DeBakey Veterans Affairs Medical Center. All experiments are terminal, with euthanasia performed at the end under a surgical plane of anesthesia. See the Table of Materials for details related to all materials, reagents, and equipment used in this protocol.

1. Preparation of antibodies and dyes

1. Antibody and dye selection
   NOTE: The number of simultaneous colors observed will be dependent on the specific microscope setup. In the system used here, four different wavelengths can be visualized simultaneously.
   1. Visualizing vessels:
      1. Based on the experimental design, label the vasculature with either an endothelium-specific (e.g., CD31) antibody and/or a molecule that is uniformly distributed in the plasma (e.g., dextran or albumin).
         NOTE: This article describes the use of Texas Red-labeled dextran (150 kDa; 250 µg/mouse). The benefit of this large molecular weight protein is in its ability to stay within the vasculature during acute inflammation. Care must be taken when using antibody labeling to avoid antibodies that have the potential to interfere with endothelial function.
   2. Visualizing platelets:
      1. To identify platelets, use either genetically modified mice (e.g., R26R-EYFP^f/f/PF4-Cre) or platelet-specific (e.g., anti-GPIbβ) antibodies for IVM. For in-depth information regarding the use of fluorescent labeling of platelets, refer to a previous publication⁴.
         NOTE: Here, DyLight488-labeled anti-GPIbβ antibodies (X488; 6 µg/mouse) are used. Care must be taken when using antibody labeling to avoid interference with platelet function or adhesion.
   3. Visualizing leukocytes:
      1. Similar to imaging platelets, visualize the leukocytes of interest with either genetically modified mice (e.g., LysM-EGFP for granulocytes) or leukocyte-specific antibodies (e.g., Ly6G for murine neutrophils). To follow this protocol, use Brilliant Violet 421 (BV421)-labeled anti-Ly6G antibodies (3 µg/mouse). In acute (<6 h) experiments, label neutrophils with 3 µg of anti-Ly6G (clone 1A8) per animal.
         NOTE: When using anti-Ly6G antibody (clone 1A8), it should be noted that this antibody has been used to deplete neutrophils (0.05-1 mg antibody/mouse)^7-9 after at least 24 h and/or with repeated injections. Anti-Ly6G (1A8) antibodies have also been reported to affect neutrophil adhesion and transmigration at high (but not low) shear stress in certain, but not all, situations ^9,10. Therefore, the antibody dosage should be chosen carefully for each case.
   4. Additional proteins:
      1. Depending on the number of lasers and filter sets/detectors available on a particular imaging system, add more wavelengths to identify more objects simultaneously while taking care to avoid spectral overlap.
         NOTE: This protocol describes the imaging of a 4th object, P-selectin (clone Psel.KO2.3), using a PerCP-eFluor 710-labeled anti-P-selectin antibody (4 µg/mouse). Although the majority of the descriptions and images in this paper are for LPS-induced acute liver injury, other injury models may be of interest and may call for alternative labels. See the discussion for a description of antibodies used for APAP-induced injury⁶.
2. Removal of preservatives and sterilization
   NOTE: Although there are commercially available antibodies and dyes ready for injection into animals, most of them will have carriers or preservatives (e.g., sodium azide) that may be harmful to animals or add confounding variables. Antibodies and dyes containing carriers or preservatives should be dialyzed against Dulbecco's phosphate-buffered saline without calcium or magnesium (DPBS^-/-; or other physiologic buffers).
   1. Pour 500 mL DPBS^-/- (~1,000-fold volume of the antibody/dye to be dialyzed) into a beaker with a magnetic stir bar. Attach a 7,000 MWCO cassette to a float buoy and place it in the dialysis buffer for at least 5 min to precondition the membrane. Remove the cassette and blot the plastic edges dry with lint-free tissue. Avoid touching the dialysis membrane.
   2. Inject the antibody/dye into the cassette through one of the four ports using an 18-20 G needle. Place the cassette with the antibody/dye into the dialysis buffer. Place the beaker on a magnetic stirrer and dialyze for at least 1 h at 4 °C. Change the dialysis buffer.
   3. Repeat the dialysis and changing of the dialysis buffer in step 1.2.2 for a minimum of two more times (at least three exchanges).
   4. Remove the cassette and blot the plastic edges dry with lint-free tissue. Avoid touching the dialysis membrane. Aspirate the antibody/dye from the cassette through an unused port using an 18-20 G needle.
   5. In a tissue culture hood, sterilize the preparations using a 0.2 µm filter prior to use in animals. Store unused portions in sterile tubes at 4 °C.
3. Controls
   1. While the exact controls necessary for each experiment will vary depending upon the design of the study, be sure to include proper sham cohorts, vehicle-treated and non-vehicle-treated groups, as well as isotype controls in order to exclude non-specific labeling and biological effects when analyzing the study results.

2. Intraperitoneal endotoxin injection

NOTE: To evaluate platelet-neutrophil-endothelial interactions in the liver sinusoids in an experimental model of murine sepsis, endotoxin injection can be used.

1. Weigh the mice to determine the amount of endotoxin to inject (5 mg/kg; Escherichia coli serotype O111:B4; endotoxin content 1 x 10⁶ EU/mg).
   NOTE: Endotoxin potency and effect are species- and strain-dependent and may vary from lot to lot¹¹. Therefore, to maximize the consistency of results, experiments should be conducted with the same lot of endotoxin¹².
2. Draw up endotoxin diluted with normal saline (0.9% sodium chloride) into a syringe with a 27 G needle attached. In the non-dominant hand, hold the mouse by the scruff using the thumb and index finger. Using the other hand, put traction on the tail to expose the abdomen and place the tail between the fifth digit and the palm of the non-dominant hand.
3. Clean the abdomen with 70% ethanol. Insert the needle into the lower-left (or right) abdomen and inject the endotoxin.
   NOTE: Care should be taken to avoid inserting the needle too deeply because there is a risk of organ injury that may confound the experiments (e.g., injury to the intestines, bowel, kidneys, etc.).
4. Perform steps 2.1-2.3 in control animals using normal saline (or vehicle).
   NOTE: APAP-induced liver injury can be accomplished using a similar methodology as above, using APAP rather than endotoxin⁶. See the discussion for details related to the APAP-induced liver injury model.

3. Induction and maintenance of anesthesia of the animals and euthanasia

1. Place the animals under a surgical plane of anesthesia. Maintain the anesthesia using inhaled isoflurane via a nosecone or a tracheostomy tube. Assess and adjust the level of anesthesia by monitoring for movement after a toe pinch.
2. Perform euthanasia at the end of all experiments by exsanguination under a surgical plane of anesthesia followed by bilateral pneumothoraces.

4. Catheterization of the jugular vein and trachea

1. Perform a tracheotomy to facilitate breathing and the placement of vascular catheters for the injection of labeled antibodies/dyes.
   NOTE: A surgical dissection microscope or loupes are helpful in catheterization.
2. Shave the fur from the neck and abdomen; remove excess hair. Place the animal on a warming pad to maintain the body temperature at 37 °C, using a rectal probe to monitor core temperature. Scrub the neck with betadine followed by 70% ethanol.
3. Make a vertical midline incision in the neck and expose the trachea using blunt dissection.
   1. Tracheostomy
      1. Prepare a tracheostomy tube PE90 tube with a 21 G blunt needle inserted (Figure 1A).
         NOTE: Adding a ~45° bevel to the end of the tracheostomy tube may help facilitate its insertion into the trachea. Alternatively, an 18-20 G peripheral intravenous catheter may also be substituted for PE90 tubing.
      2. Once the skin and fascia are incised, reflect the salivary glands away from the center.
      3. Using blunt dissection, separate the sternothyroid muscle to give better access to the trachea.
         NOTE: Take care to avoid the supplying vessels and the carotid arteries that are adjacent to the trachea.
      4. Make a small (~1-2 mm) horizontal incision between the tracheal rings on the ventral/anterior side of the trachea (Figure 1B).
         NOTE: The use of Vannas scissors will facilitate proper dissection. Ensure that only the anterior face of the trachea is incised; complete transection of the trachea will make it difficult to cannulate the trachea and risks damaging nearby vascular structures.
      5. Pass a tracheostomy tube into the hole. Ensure that the tip of the tube is not inserted past the carina.
      6. Secure the tracheostomy tube within the trachea by tying a #4-0 silk suture around the trachea and tube, using the tracheal rings as support (Figure 1C).
   2. External jugular cannulation
      1. Prepare a jugular catheter (Figure 2) using PE50 tubing (OD 0.97 mm, ID 0.58 mm) with PE10 tubing (OD 0.61 mm, ID 0.28 mm) inserted into one end (Figure 2 inset) and a 22 G or 23 G blunt needle inserted into the other. Use adhesive to ensure that the tubing does not slip or leak. Attach a syringe filled with sterile normal saline to flush the catheter.
      2. Expose the external jugular vein (EJV) after salivary gland reflection. Using blunt dissection, free the EJV from the surrounding connective tissue (Figure 3A).
      3. Using #4-0 silk sutures, ligate the cranial end of the visualized EJV and place a loose loop around the caudal end of the EJV (Figure 3B). Clamp the ends of each suture together with hemostats and gently retract the two hemostats away from each other to help expose the EJV.
         NOTE: The optimal region to cannulate is between the branches of the anterior jugular vein cranially and the internal jugular vein caudally. To ease suture placement, fold a piece (~15 cm) of suture in half and pass the bent end underneath the EJV with one pair of forceps while raising the EJV with a separate pair of forceps to prevent rolling of the vessel. Then, cut the suture to make two separate lengths.
      4. Using Vannas scissors, make a small (~1 mm) incision in the anterior surface of the EJV. Insert a bent 30 G blunt needle (augmented with a 0.15 mm minutien pin bluntly filed and inserted into the needle tip) into the EJV through the incision and gently lift it up to facilitate passage of the catheter containing normal saline into the EJV.
         NOTE: Ensure that the catheter and hub do not have any air bubbles before insertion and flushing into the mouse. An air embolus may be lethal to the animal.
      5. Once the catheter is within the EJV, loosen the caudal loop just enough to allow passage of the catheter deeper into the vein. Check for placement of the catheter by aspirating blood and flushing the catheter to check for leaks (Figure 3C).
      6. Using the caudal loop, secure the catheter within the EJV. Using the cranial suture, secure the catheter to the cranial end of the EJV to minimize accidental decannulation (Figure 3D).
      7. Close the neck skin incision with #4-0 silk suture.

5. Exteriorization of the liver

1. Clean the abdomen with a betadine scrub followed by 70% ethanol.
2. Make a ~1 cm vertical incision in the skin of the upper abdomen beginning at the base of the sternum. Using blunt dissection, separate the skin from the abdominal muscles. Cauterize the large feeding vessels of the skin and muscles to minimize bleeding during the following steps (Figure 4A).
3. Make a ~1-2 cm horizontal incision of the skin followed by the abdominal muscle, bisecting the vertical incision in step 5.2 approximately at the top of the visible liver; use a cautery tool to assist in hemostasis of the skin (Figure 4B).
   NOTE: The size of the abdomen incision is critical for optimal imaging of the liver. If the opening is too small, there is a risk of liver ischemia due to overstretching or impingement of the hepatic artery and/or portal vein. If the opening is too large, then other organs, such as the intestines, may also be exteriorized and interfere with imaging.
4. Cut an approximately 1 cm circle in the center of a 2 in x 2 in gauze and moisten it with warm normal saline. Position the gauze so the opening is centered over the incision. Gently pull the edges of the incision apart and apply light, upward pressure on the abdomen to exteriorize the liver so that the liver rests easily on top of the moistened gauze (Figure 4C).
5. Place the mouse onto the imaging tray in the prone position, with the liver centered over the coverslip (Figure 4D).
   NOTE: In a control animal, the liver should appear maroon. An inverted, laser scanning confocal microscope with a two-way resonance head scanner is used for IVM in this protocol. Therefore, a 3D printer was used to create a custom IVM tray (Figure 5). The .stl files created for 3D printing are provided (Supplemental File 1); adjustments may be necessary to fit individuals' needs.
6. Transfer the animal to the inverted microscope.

6. Liver intravital microscopy imaging

NOTE: This protocol uses a laser scanning confocal microscope with a two-way resonance head scanner for intravital microscopy. To stabilize the resonance head frequency, turn on the system for at least 30 min prior to imaging. Different systems may have different capabilities. Discussion of these is outside the scope of this article. Contact the equipment representatives for the technical details of specific systems.

1. To maintain a surgical plane of anesthesia, connect the animal to the isoflurane delivery system.
   NOTE: Depending on the experimental protocol, mechanical ventilation may be necessary (vs. spontaneous ventilation).
2. Place a radiant warming pad over the animal, using the rectal probe to maintain a core temperature of 37 °C.
   NOTE: In this protocol, a rectangular piece of foam that fits around the IVM tray acts as an insulator and a spacer on which to place the radiant warming pad (Figure 4D).
3. Set the objective turret to the desired magnification, applying immersion oil as necessary. To follow this protocol, use the 60x/NA1.30 silicone oil objective with 1x optical zoom.
4. Once the focal plane is identified, identify ≥10 random fields of view, avoiding the areas near the edges of the liver.
   NOTE: As the liver exhibits autofluorescence in the fluorescein isothiocyanate (FITC) channel, use epifluorescence to quickly identify the general focal plane.
   1. Depending on the system, identify the edges of the visible liver using epifluorescence in the FITC channel and record the (x,y) stage coordinates.
   2. Use the (x,y) boundaries of the liver and a random number generator to identify random (x,y) coordinates. For an acceptable field of view, look for (a) the presence of vessels dispersed throughout the field of view; (b) ≥80% of the vessels in the same focal plane; and (c) ≥1 field of view away from any liver edges (example in Figure 6). If the focus of image analysis is on platelet-leukocyte-endothelial interactions in the hepatic sinusoids, exclude fields with vessels whose diameters are ≥15 µm (murine sinusoids are typically ≤10 µm in diameter).

7. Image analysis

NOTE: ImageJ/FIJI was used to process all of the images in this paper. FIJI (fiji.sc) is an open-source version of Image J (imagej.nih.gov) that has more functionality through the use of additional plugins and macros¹³.

1. Open the multichannel, time-lapsed files in ImageJ/FIJI.
   NOTE: Depending on the platform, additional programs/plugins may be required. ImageJ/FIJI has a Bio-Formats Importer that can open multiple different types of files, such as the .oir files by Olympus. Here, the files have been converted to .tif because it is a more widely used format.
2. Make sure that the correct scale is set; look for the pixel/µm ratio and the time interval in the metadata/properties.
3. Use different filters to remove background noise (e.g., a median filter with a 2 pixel radius used in this protocol). Ensure that the processing is similar amongst all the images to minimize variation in post-processing. As counts and lengths are being compared in these experiments, instead of mean fluorescence intensity, filter to improve clarity and the identification and measurement of cell-cell interactions.
   NOTE: The use and magnitude of filters will be dependent on the size of the target(s) of interest and the signal-to-noise ratio in the images.
4. As there is heterogeneity between fields of view in vascular density, measure the total vascular area in each field using the selection brush (Figure 7).
   1. Right-click the oval selection tool and pick the selection brush tool.
   2. Adjust the size of the brush by double-clicking the selection brush tool icon.
   3. Trace over the vessels to highlight/select them. To delete parts of the selection, use [alt]-left click and add parts by using [shift]-left click.
   4. Press [M] to measure the selected area.
      NOTE: If the image scale has been properly set, then the output should be in µm², which is converted to mm².
5. Count the number of platelets and leukocytes in the field. Consider cells to be adhered cells if they moved <1 cell body over 30 s. Calculate the number of cells (platelets and/or leukocytes) per mm².
6. To estimate the relative blood flow within each vessel, measure the mean red blood cell (RBC) velocity as a surrogate using the MTrackJ plugin in ImageJ/FIJI (imagescience.org/meijering/ software/mtrackj)¹⁴, as described previously⁵. Identify RBCs as round or disc-shaped filling defects in the dextran channel. Save the tracks for record keeping, asynchronous analysis, and verification.
   NOTE: An example of this is shown in Figure 8 and Supplemental Video S1.

Results

Assessing the effect of vimentin rod domain in leukocyte adhesion to inflamed endothelium
Leukocyte P-selectin glycoprotein ligand-1 (PSGL-1) binding to endothelial and platelet P-selectin occurs during the acute phase of sepsis-induced liver injury inflammation. However, the recombinant human rod domain of vimentin (rhRod) has been shown to bind to P-selectin and block leukocyte adhesion to both endothelium and platelets. This protocol was utilized in a mouse model of sepsis to visualize the real-ti...

Discussion

The purpose of this methods paper is to outline the necessary steps required to reliably capture high-resolution intravital images and videos of the mouse liver under homeostatic conditions and following the administration of endotoxin or APAP. While this protocol has allowed for the consistent production of data on platelet-leukocyte-endothelial interactions in the liver, there are a number of critical steps required for success, as well as potential pitfalls that are important to avoid when using this imaging paradigm....

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Surgical Supplies
2" x 2" non-woven sponges	McKesson Med. Surg	92242000	For liver isolation
#4-0 silk braided suture with needle	SOFSILK	N/A	4-0 Softsilk coated braided black, nonabsorbable: C-1 cutting needle
#4-0 silk braided suture without needle	Ethicon	N/A	4-0 Black braided silk, nonabsorbable
21 G blunt needle (0.5 inch)	SAI Infusion Technologies	B21-50	This is used to attach to the end of the tracheostomy tube to allow for connection to the ventilator. An alternative source is Instech
23 G blunt needle (0.5 inch)	SAI Infusion Technologies	B23-50	This is used for the vascular catheter to allow for connection to a syringe. An alternative source is Instech
Dissecting Scissors (Pointed Tip)	Kent Scientific	INS600393-G	Micro Dissecting Scissors; Carbide Blades; Straight; Sharp Points; 24 mm Blade Length; 4 1/2" Overall Length
McPherson-Vannas Micro Scissors (Vannas)	Kent Scientific	INS600124	These are useful for creating the openings in the trachea and vessels
Polyethylene tubing 10	Instech	BTPE-10	This is used to make the intravascular portion of the catheter. An alternative source is BD Intramedic
Polyethylene tubing 50	Instech	BTPE-50	This is used to make the extravascular portion of the catheter.  An alternative source is BD Intramedic
Polyethylene tubing 90	Instech	BTPE-90	This is used to make the tracheostomy tube.  An alternative source is BD Intramedic
USP grade sterile normal saline	Coviden	8881570121	Hospira 0.0% Sodium Chloride Injection, USP
Microscopy Supplies
Isoflurane delivery system and ventilator	Kent Scientific	Somnosuite	Combination rodent ventilator and volatile anesthetic delivery system
Foam spacer for warming pad during microscopy	N/A	N/A	This spacer should be cut from high quality foam, should fit around the liver microscope tray and specific height dimensions are dependent upon the microscope system
Laser scanning confocal microscope system with resonance head scanner	Olympus	FV3000	Although we describe the use of an Olympus FV3000 using a resonance head scanner, this protocol with work with most imaging systems
Liver Microscope Tray	N/A	N/A	The liver microscope tray was designed for an inverted microscope
Antibodies & Related Reagents
Brilliant Violet 421/anti-mouse Ly6G antibody	BioLegend	127628	3 µg/mouse. To label neutrophils
BV421/F4/80 antibody	BioLegend	123132	0.75 mg/kg. To label Kupffer cells
Dulbecco's phosphate buffered saline w/o calcium or magnesium	Gibco/ThermoFisher Scientific	14190144	Used as dialysate to remove sodium azide from antibodies
DyLight649/anti-GPIbβ antibody	emfret Analytics	X649	3 µg/mouse. To label platelets
DyLight488/anti-mouse GPIbβ antibody	emfret Analytics	X488	6 µg/mouse. To label platelets
Endotoxin from Escherichia coli serotype O111:B4	Sigma-Aldrich	L3024	5 mg/kg; Potency of endotoxin may vary from lot to lot. Therefore, the same lot should be used for a series of experiments to minimize variation due to endotoxin lot
PerCP-eFluor 710/anti-mouse P-selectin antibody	Invitrogen	46-0626-82	4 µg/mouse. To label P-selectin
Slide-a-Lyzer 7,000 MWCO cassette	Thermo Scientific	66370	Used to dialyze antibodies to remove sodium azide
Texas Red-labeled dextran	Sigma-Aldrich	T1287	~150 kDa; 250 µg/mouse
TRITC/bovine serum albumin	Sigma-Aldrich	A2289	500 µg/mouse. Dilute to a stock concentration of 50 mg/mL (5%) in normal saline. Used to label the vasculature. It may leak into the interstitial space more readily than high molecular weight dextran during inflammation