Inducing Ischemia-reperfusion Injury in the Mouse Ear Skin for Intravital Multiphoton Imaging of Immune Responses

Summary

This protocol describes the induction of an ischemia-reperfusion (IR) model on mouse ear skin using magnet clamping. Using a custom-built intravital imaging model, we study in vivo inflammatory responses post-reperfusion. The rationale behind the development of this technique is to extend the understanding of how leukocytes respond to skin IR injury.

Abstract

Ischemia-reperfusion injury (IRI) occurs when there is transient hypoxia due to the obstruction of blood flow (ischemia) followed by a subsequent re-oxygenation of the tissues (reperfusion). In the skin, ischemia-reperfusion (IR) is the main contributing factor to the pathophysiology of pressure ulcers. While the cascade of events leading up to the inflammatory response has been well studied, the spatial and temporal responses of the different subsets of immune cells to an IR injury are not well understood. Existing models of IR using the clamping technique on the skin flank are highly invasive and unsuitable for studying immune responses to injury, while similar non-invasive magnet clamping studies in the skin flank are less-than-ideal for intravital imaging studies. In this protocol, we describe a robust model of non-invasive IR developed on mouse ear skin, where we aim to visualize in real-time the cellular response of immune cells after reperfusion via multiphoton intravital imaging (MP-IVM).

Introduction

Ischemia-reperfusion injury (IRI) occurs when there is a transient hypoxia due to the obstruction of blood flow (ischemia) followed by a subsequent re-oxygenation of the tissues (reperfusion). In the skin, ischemia-reperfusion (IR) is thought to be one of the contributing factors to the pathophysiology of pressure ulcers, where prolonged bed rest predisposes long-term hospital patients to injury. In these patients, both the skin and the underlying muscles are constantly exposed to weight pressure exerted over areas of bony prominence, resulting in localized injuries that, if left untreated, may become necrotic¹.

The damages involved in an IRI are twofold. During ischemia, the occlusion of blood vessels leads to a drastic drop of oxygen delivery to the tissues. This results in a decrease of ATP and pH, which inactivates ATPases involved in cellular metabolism. In turn, cellular calcium levels spike, and stressed or damaged cells undergo apoptosis or necrosis². The release of intracellular contents or damage associated molecular patterns (DAMP), like HMGB1, contributes to the inflammatory response³. The second insult occurs during reperfusion. Although oxygen and pH levels are restored during reperfusion, this results in the generation of reactive oxygen species (ROS), which leads to the oxidation of intracellular lipids, DNA, and proteins. Consequently, pro-inflammatory mediators are activated, which sets off a secondary inflammatory response that involves the recruitment of immune cells to the inflammatory site². While the cascade of biochemical events leading up to the inflammatory response has been well described, the spatial and temporal regulation of the immune cell activities are not well understood.

Here, we describe a robust IR model on mouse ear skin using simple magnet clamping. Coupled with multiphoton intravital imaging (MP-IVM), we established a model to study the in vivo inflammatory responses that occur after reperfusion takes place. The rationale behind the development and use of this technique is to try to understand how both interstitial and infiltrating cells respond to IR in real time.

Existing models of IR using the clamping technique on the skin flank are highly invasive, as they require the surgical implantation of steel plates in the skin flank, making them less-than-ideal for immunological studies⁴. A similar non-invasive clamping technique has been described in the mouse skin flank^5,6. However, because of the incorporation of the intravital imaging component in this method, we instead chose the ear skin as the targeted IR site, as it circumvents movements due to breathing and offers stability during imaging^7,8. Moreover, leukocyte subsets that span the interstitium are identical between the ear skin and the skin flank, although the numbers and proportions may vary slightly⁹. Thus, the ear skin represents an ideal imaging site.

In addition, most data retrieved from these IRI models are limited to macroscopic evaluations (grading of ulcers) and microscopic analyses of endpoint inflammatory indicators¹⁰. Using this model, real-time visualization of the cellular response of neutrophils after reperfusion in the skin of a fluorescent reporter mouse is enabled. A previously published intravital ear imaging model is utilized⁸ with additional modifications (Figures 1, 2).

Protocol

All experiments dealing with live animals were conducted in accordance to all relevant animal use and care guidelines and regulations.

1. Choice of Fluorescent Reporter Mice

1. Use 6- to 12-week-old LysM-eGFP¹¹ mice (no preference for either males or females).
   Note: The use of various cell-specific fluorescent reporter mice enables the visualization of different immune cells in vivo. In this strain, circulating neutrophils (GFP^hi cells), circulating monocytes (GFP^lo cells), and dermal macrophages (GFP^lo cells) can be visualized. With the imaging parameters used, only the bright signals from GFP-positive neutrophils will be detected.
   Note: A list of immune-cell-specific fluorescent reporter mouse strains suitable for this type of skin imaging study can be found in Reference 8.
   Note: It is highly recommended that albino mice be used for imaging, as pigmented mice are more prone to photodamage. This is because the pigmented ear skin is much more sensitive to laser-induced speckling (indicative of tissue burning). As a result, neutrophil recruitment and accumulation may be observed even during the steady state^8,12.
2. Keep the mice in specific pathogen-free (SPF) conditions with 12-h light-dark cycles.

2. Mouse Anesthesia

1. Anesthetize the mouse with an intra-peritoneal injection of ketamine-xylazine (8 µL g^-1 bodyweight), composed of a mixture of 15 mg mL^-1 ketamine and 1 mg mL^-1 xylazine dissolved in sterile water.
2. Place the mouse on a heating pad to maintain its body temperature at 37 °C throughout the preparation procedure. Check for sufficient anesthesia by observing an absence of a toe pinch reflex.
   Note: After the first hour, subsequent quarter doses of anesthetic will need to be administered subcutaneously and will last for approximately 0.5 h each. The twitching of whiskers or the tail may also indicate that the anesthesia is wearing off and that a top-up is required.
3. Use ophthalmic lubricant on the eyes to prevent dryness while under anesthesia.

3. Depilation

1. Carefully apply depilatory cream to the upper two-thirds of the dorsal mouse ear using cotton-tip applicators.
2. Wait for 2 - 3 min before removing the cream using wet cotton-tip applicators in a thorough but gentle manner.
   Note: Do not allow hair removal cream to stay on the mouse ear for too long, as it may induce inflammation^13,14.

4. Induction of Ischemia and Reperfusion Injury

1. Use gold-plated, N42-grade neodymium magnets, 12 mm diameter x 2 mm thick, and with a Gauss rating of approximately 3,000 to induce ischemia in the mouse ear skin.
   Note: In this case, the dimpled face of the magnets denotes its north pole.
2. Slot the magnets into their individual plastic guides.
   Note: The plastic guide serves to ease the placement and separation of the high-strength magnets. Due to their strong magnetic force yet low resistance to breakage, do not place individual magnets in close proximity to each other or to other metals. Breakage and splintering may occur if they pull towards each other.
3. Position the first (dorsal) magnet such that only the edge is in contact with the second (ventral) magnet (Figure 3a)
   Note: This prevents the magnets from snapping together before they have been properly positioned.
4. Position both magnets such that the ventral magnet lies flat on the ear (Figure 3a).
   Note: Before ischemia is induced, ensure that the mouse is kept at 37 °C and that sufficient anesthesia is maintained (see step 2.2).
5. Once ready, carefully let the magnets come together (Figure 3a).
   Note: For imaging purposes, clamp only half of the ear so that an IR and non-IR region can be observed.
6. After 1.5 h of ischemia, remove the magnets by twisting the magnets away from each other using the plastic guides, allowing reperfusion to take place.
   Note: Care must be taken to prevent the ears from creasing when the magnets are placed. Incomplete ischemia is evident if the macroscopically visible major blood vessels re-perfuse immediately (i.e., blood fills the vessels immediately) after the magnets are removed. Although reperfusion does not occur immediately after the removal of magnets, blood vessel occlusion is only transient. As such, it is imperative to prepare the mouse ear for imaging as swiftly as possible.

5. Injection of Blood Vessel Labeling Agents

1. Immediately after magnet removal, administer intravenously (via retro-orbital or tail vein injection) Evans blue (10 mg mL^-1 in PBS or saline; 1 µL g^-1 bodyweight) or another blood vessel labeling agent of choice.
   Note: Before injection, ensure that sufficient anesthesia is still maintained by performing gentle toe pinching.

6. Placement of the Ear on the Imaging Platform

1. Cut 2 pieces of masking tape 1.5 cm in length and 1.8 cm in width.
2. Allow the adhesive sides to stick together whilst leaving about 1 mm of adhesive along its width.
3. Cut the masking tape in two, lengthwise, to accommodate its placement within the slit on the ear platform.
4. Insert this masking tape about halfway through the slit, such that the adhesive side faces up.
5. Position the mouse on the heating pad, such that the ear to be imaged is next to the masking tape strip.
6. By using two PBS-moistened cotton-tip applicators, gently press the ear against the adhesive strip.
7. Using the strip as a guide, bring the mouse ear through the slit while simultaneously adjusting the mouse closer towards the stage.
8. To remove the sticky tape, first add a drop of PBS to reduce the adhesiveness of the tape.
9. Separate the mouse ear from the masking tape as gently as possible using a fine paintbrush.
10. Flatten the ear against the ear platform by gently rolling a moist cotton-tip applicator over the ear.
11. Put a drop of PBS underneath the coverslip (which is kept in position on the coverslip holder using grease; Figure 2) and gently place it over the ear. Top up with more PBS if necessary.
    Note: The coverslip holder increases the stability during imaging.
12. Insert the rectal temperature probe and connect the wires to the heating system according to manufacturer's instructions.
    Note: Set the temperature of the body heating pad to 37 °C and the ear stage platform to 35 °C.

7. Multiphoton Microscope Setup and Imaging Parameters

Note: This protocol uses a single beam, multiphoton microscope with a tunable (680 - 1,080 nm) Ti:Sa laser (3.3 W at 800 nm; pulse length of 140 fs; 80 MHz repetition rate) with a 20X water objective (NA = 1.0) for intravital imaging studies.

1. Open the imaging software.
2. Align the laser according to the manufacturer's instructions.
3. Adjust the excitation wavelength to 950 nm.
   Note: GFP and Evans blue can be simultaneously excited at 950 nm.
4. To preview, use the following settings: 500 µm² scanfield, 505 x 505 pixel resolution, and a scan frequency of 800 Hz in a single line scan. Click "Preview."
5. Toggle the attenuator (laser) power. Ensure that all essential signals are picked up without exposing the imaging field to excessive amounts of laser power, which may induce heat damage.
   Note: Start at a low attenuator power and increase if the signal is dim. As neutrophils will be absent from the interstitium at early time points post-reperfusion, macrophages may be used as a gauge to determine the minimal power required, as the former are dimmer than neutrophils.
   Note: If this step is to be done for the first time, adjust the settings in the non-ischemic zone, where intact vascular integrity is expected and will facilitate the setting of the attenuator power. In subsequent experiments, these settings do not require much modification unless the laser power is unstable.
6. Adjust the PMT settings.
   Note: Check with the manufacturer for the maximal optimal gain voltage for the PMT. Setting the PMT voltage beyond the recommended threshold will result in a higher signal-to-noise ratio. The general recommendation is to set the PMT gain voltage to the recommended threshold and to increase the attenuator power as necessary should the signal be too dim.
7. Select an imaging region that is in close proximity to the edge of the ischemia, characterized by massive Evans blue leakage.
8. Collect GFP and Evans blue signals using 525/50 bandpass (BP) and 655/40 BP filters, respectively. For second harmonic generation (SHG) signals of collagen fibers within the dermal compartment, use a 475/42 BP filter.
9. Create a folder to save the images in a format that is compatible for the available image analysis software.
10. To acquire, use the following settings: 500 µm² scanfield, 505 x 505 pixel resolution, and a scan frequency 400 Hz in a single line scan.
11. A 100-µm z-stack with a step size of 4 µm can be acquired repeatedly over time, at 1 min intervals, to monitor neutrophil infiltration.
    Note: Especially for leukocytes (e.g., neutrophils) with a higher migratory velocity, an interval that is longer than 1 min may result in difficulties during cell tracking analysis. In this case, the user can either reduce the thickness of the acquisition stack or increase the scan frequency.
    Note: While the required acquisition stack of the ischemic zone may appear smaller as a result of the compression (alternatively characterized by a higher SHG intensity), subsequent reperfusion will result in massive inflammation that causes the ear to swell. Significant drifts in the Z-direction are expected. As such, the acquisition of a large z-stack is necessary to accommodate the drift.
12. Throughout the imaging, top up the PBS and water regularly to keep the ear moist and the objective lens immersed in water.
    Note: A typical experiment usually lasts for 2 - 4 h. Depending on the experimental design, and coupled with a well-controlled anesthesia regime, extending the duration of imaging is possible.

8. Terminating the Experiment

1. Euthanize the mouse by carbon dioxide asphyxiation according to the institution's Institutional Animal Care and Use Committee (IACUC) procedures.
   Note: Always euthanize the mouse by approved methods as determined by the institution's rules, regulations, and guidelines. If repeated imaging is needed, keep the mouse on the heating pad until the anesthesia wears off. Once the mouse has regained sufficient consciousness and is mobile, return the mouse to its cage.

9. Image Analysis

Note: Data generated from the imaging experiment can be visualized by different software packages.

1. Open the imaging analysis software.
2. Under Surpass Mode, import the file (Open → select any file in the desired folder → Click "open").
3. Edit pseudo colors if default colors do not apply (Edit → Display adjustment → Under the channel tab, select the desired color from the color palette).
4. Adjust the brightness, contrast, and background (Edit → Display Adjustment → Toggle Max and Min values).
5. Ensure that the file dimensions are correct (Edit → Image properties → Under the geometry tab, check the voxel size. In this protocol, X = 0.99, Y = 0.99, and Z = 4).
   Note: Derive X and Y by dividing the scanfield dimension by the pixel resolution. Z is the thickness between each slice.
   Note: The output of these data sets can be presented as maximum projection movies.
   Note: Tracking analysis is required to fully characterize the cellular activities and interactions in order to understand their function in vivo. Detailed instructions of cell tracking using the spot function available in the software can be found in the protocol in Reference 15.
   Note: There are many ways of quantifying leukocyte migration, details of which can be found in the review article in Reference 16.

Results

This protocol uses a custom-built ear skin imaging platform, as shown in Figure 1. Several features of this platform are specifically designed to facilitate imaging while maintaining physiological settings. Placing the ear on the heated brass platform not only maintains the ear at a physiological temperature of 35 °C, but it also isolates the ear from inevitable movements due to breathing. The addition of a metal clip on the brass platform creates a gap to prevent the cov...

Discussion

Significance

IR is one of the leading causes of skin pressure ulcers. The early stages (I and II) of pressure ulcers describe the condition of the human skin (as compared to the underlying subcutaneous tissues and muscles). However, an understanding of the immunological etiology is still lacking. Here, we present a simple and robust IR model on mouse ear skin in order to address this gap. We simulate ischemia by clamping the mouse ear between two magnets and subsequently study the downstream immu...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Mice strains
Lysozyme-GFP C57BL/6	Thomas Graf, Center for Genomic Regulation
C57BL/6-C2J	Jackson Laboratories	000058	To be crossed with Lysozyme-GFP to generate albino Lysozyme-GFP for skin imaging
Reagents
PBS
Viaflex 0.9% (wt/vol) saline	Baxter Healthcare	F8B1323
Ketamine (100 mg mL−1 ketamine hydrochloride	Parnell		Ketamine is a controlled drug and all relevant local regulations should be followed
Ilium Xylazil-20 (20 mg mL−1 xylazine hydrochloride)	Troy Laboratories		Xylazil-20 is a controlled drug and all relevant local regulations should be followed.
Evans blue (10 mg mL−1 in PBS or saline)	Sigma-Aldrich	46160
Ultrapurified water
Equipment
Insulin syringe with needle	BD	328838
Transfer pipettes	Biologix Research Company	30-0135
3 M paper masking tape	3M	2214
Deckglaser microscope cover glass (22 mm × 32 mm)	Paul Marienfeld	101112
Curved splinter forceps	Aesculap, B. Braun Melsungen	BD312R
Veet hair removal cream	Reckitt Benckiser
Medical cotton-tipped applicators	Puritan Medical Products Company	806-WC
C-fold towels	Kimberly-Clark	20311
Kimwipes delicate task wipes	Kimtech Science	34155
Gold-plated, N42-grade neodymium magnets, 12 mm in diameter and 2 mm thick	first4magnets	F656S
Plastic guide, 10 cm by 1.5 cm (polyvinyl chloride material)			fold in half lengthwise, bind with masking tape and slot magnet in
High vacuum grease	Dow Corning
Microscope
TriM Scope II single-beam two-photon microscope	LaVision BioTec
Tunable (680–1,080 nm) Coherent Chameleon Ultra II One Box Ti:sapphire laser (≥3.3 W at 800 nm; pulse length of 140 fs, 80 MHz repetition rate)	Coherent
Water-dipping objectives (20×, NA = 1.0)	Olympus	XLUMPLFLN20xW
Miscroscope filter and mirror sets (for imaging GFP, SHG, Evans Blue)
495 long-pass	Chroma	T495LPXR
560 lomg-pass	Chroma	T560LPXR
475/42 band-pass	Semrock	FF01-475/42-25
525/50 band-pass	Chroma	ET525/50m
655/40 band-pass	Chroma	NC028647
Skin-imaging stage platform (refer to diagram for assembly)
A metal base plate (126 mm × 126 mm × 1 mm)
A brass platform for the ear (79 mm × 19 mm; 1 mm thickness at side, 0.5 mm thickness in the middle; Figure 1) with slit (1.7 mm × 1 mm; 1.5 mm away from long edge)
Two plastic blocks (10 mm in height)—for heat insulation
Curved holder, for positioning the control thermistor on the ear platform
Interface cable CC-28 with DIN connector and thermistors, one for the temperature control and the other for the temperature monitor	(Warner Instruments (Harvard Apparatus)	640106	connect the interface cable to both resistive heater blocks set at 35 °C
Resistive heater blocks RH-2	(Warner Instruments (Harvard Apparatus)	640274	Resistive heater blocks can heat the brass ear platform up to over 100 °C within minutes. Ensure that the control thermistor has been properly secured in the holder in order to avoid overheating.
Temperature controller TC-344B for the ear platform	(Warner Instruments (Harvard Apparatus)	640101
Temperature controller TR-200 for mouse heating pad	Fine Science Tools	21052-00	Unit is no longer for sale. Ask manufacturer for alternatives
Power supply for TR-200	Fine Science Tools	21051-00	Unit is no longer for sale. Ask manufacturer for alternatives
Heating pad	Fine Science Tools	21060-00	Unit is no longer for sale. Ask manufacturer for alternatives.
Animal rectal probe	Fine Science Tools	21060-01	Unit is no longer for sale. Ask manufacturer for alternatives. After connecting the rectal probe and heating pad to the temperature controller TR-200, set the temperature to 37 °C
Coverslip holder
2 plastic rods, 1 cm in diameter, 10 cm in length
1 plastic adaptor with holes drilled to accommodate rods (refer to diagram)
3 plastic tightening screws for keeping plastic rods in place
1 metal plate, 6 cm x 2.5 cm, with a 2 cm square cut at 1 end, 2 mm edge away from short edge
1 pair of nut and bolt for attaching metal plate to plastic rod
1 acrylic base (4 cm x 5 cm x 1.5 cm) with magnet to hold coverslip holder on skin-imaging stage platform. 1 rod is permanently fixed onto base.
Imaging analysis software
Imaris v8.1.2	Bitplane