Ischemic Tissue Injury in the Dorsal Skinfold Chamber of the Mouse: A Skin Flap Model to Investigate Acute Persistent Ischemia

Summary

The window of the murine dorsal skinfold chamber presented visualizes a zone of acute persistent ischemia of a musculocutaneous flap. Intravital epi-fluorescence microscopy permits for direct and repetitive assessment of the microvasculature and quantification of hemodynamics. Morphologic and hemodynamic results can further be correlated with histological and molecular analyses.

Abstract

Despite profound expertise and advanced surgical techniques, ischemia-induced complications ranging from wound breakdown to extensive tissue necrosis are still occurring, particularly in reconstructive flap surgery. Multiple experimental flap models have been developed to analyze underlying causes and mechanisms and to investigate treatment strategies to prevent ischemic complications. The limiting factor of most models is the lacking possibility to directly and repetitively visualize microvascular architecture and hemodynamics. The goal of the protocol was to present a well-established mouse model affiliating these before mentioned lacking elements. Harder et al. have developed a model of a musculocutaneous flap with a random perfusion pattern that undergoes acute persistent ischemia and results in ~50% necrosis after 10 days if kept untreated. With the aid of intravital epi-fluorescence microscopy, this chamber model allows repetitive visualization of morphology and hemodynamics in different regions of interest over time. Associated processes such as apoptosis, inflammation, microvascular leakage and angiogenesis can be investigated and correlated to immunohistochemical and molecular protein assays. To date, the model has proven feasibility and reproducibility in several published experimental studies investigating the effect of pre-, peri- and postconditioning of ischemically challenged tissue.

Introduction

Coverage of exposed tendon, bone and implant material in reconstructive surgery relies on the use of flaps. A flap is a block of tissue that is transferred on its vascular pedicle that guarantees arterial inflow and venous outflow. Despite broad expertise and the availability of a variety of flaps to be transferred, ischemia-induced complications ranging from wound breakdown to total tissue loss are still encountered. Whereas conservative treatment and healing by secondary intention can be expected after minor tissue necrosis, significant flap necrosis usually requires surgical revision, including debridement, wound conditioning and secondary reconstruction. This increases morbidity, prolongs hospital stay and consequently leads to increased health care costs.

Flaps with an undefined pattern of vasculature or randomly perfused areas in the distal zone most remote from the arterial inflow are particularly prone to ischemic damage. Accordingly, numerous experimental and clinical studies have evaluated the development of necrosis in both, axial pattern flaps (defined blood supply) and random pattern flaps (undefined blood supply)^1-3. The main findings are commonly based on macroscopic evaluation of the size of the necrotic area. In order to assess the causes and mechanisms of tissue necrosis more in detail, several studies focused on the analysis of microcirculation. Different techniques have been used to measure tissue perfusion, including the analysis of tissue oxygen tension using polarographic electrodes^4-5, as well as the measurement of blood flow using laser Doppler flowmetry^6-7, dye diffusion⁸, and microspheres^9-10. These techniques, however, only allow for measuring indirect parameters of tissue perfusion and do not enable any morphological analysis of the microhemodynamic processes within an individual area of interest of a flap.

Sandison is known to be the first who has used a transparent chamber for prolonged in vivo studies, which he performed in rabbits¹¹. In 1943 — approximately 20 years later — Algire was the first to adapt such a transparent chamber to be applicable in mice in order to study the behavior of micro-implants of tumor cells¹². Due to the fact that mice are so-called loose skin animals and after some technical refinements over the following years, Lehr and co-workers were able to adapt such a dorsal skinfold chamber developing a smaller and lighter titanium chamber. This chamber enabled evaluation using intravital fluorescence microscopy, a technique that allows direct and repetitive visualization of a number of morphologic and microcirculatory features and their changes over time under different physiological and pathophysiological conditions, such as ischemia-reperfusion injury¹³.

In the investigation of perfusion of skin, muscle and bone flaps under normal and pathological conditions two trends occurred: First, the “acute” flap models that do not use the dorsal skinfold chamber such as the pedicled ear flap in the mouse¹⁴, the laterally based island skin flap in the hamster¹⁵ and the pedicled composite flap in the rat¹⁶. Second, the “chronic” flap model where the combination of a flap with a dorsal skinfold chamber permits repetitive microcirculatory analyses over several days with intravital fluorescence microscopy. It consists of a randomly perfused musculocutaneous flap that is integrated in the skinfold chamber of the mouse ¹⁷. Its width-to-length ratio was chosen that a situation of acute persistent ischemia consistently results in ~50% flap tissue necrosis 10 to 14 days after flap elevation. This reproducible extent of tissue necrosis allows further evaluation of both, protective (i.e., development of less necrosis) and detrimental factors (i.e., development of more necrosis) on flap pathophysiology. During the last years, several experimental publications demonstrating the effect of different pre-, peri- and post-conditioning procedures, including the administration of tissue-protective substances^18-24 and the local application of physiologic stressors such as heat²⁵ and shockwaves²⁶, have emerged.

The quantitative analyses of necrosis, microvascular morphology and microcirculatory parameters can further be correlated to immunohistochemical analyses and protein assays. Different proteins and molecules including vascular endothelial growth factor (VEGF), nitric oxide synthases (NOS), nuclear factor kappa B (NF-κB) and heat shock proteins (HSP-32: heme-oxygenase 1 (HO-1) and HSP-70) have been shown to play a role in tissue protection. Based on this chamber flap model, two modifications have been developed in order to analyze neovascularization and microcirculation during skin graft healing²⁷ and angiogenic developments in a pedicled flap with axial pattern perfusion²⁸. We present a reproducible and reliable model that includes an ischemically challenged musculocutaneous flap in the mouse skinfold chamber. This model allows visualization and quantification of the microcirculation and hemodynamics by intravital epi-fluorescence microscopy.

Protocol

NOTE: Prior to implementation of the presented model, the corresponding animal protection laws must be consulted and permission must be obtained from the local authorities. In this work, all experiments were performed in conformity with the guiding principles for research involving animals and the German legislation on protection of animals. The experiments were approved by the local animal care committee.

1. Animal Preparation and Surgical Elevation of the Flap

1. Keep animals in single cages at room temperature of 22–24 °C and at a relative humidity of 60–65% with a 12-hr day-and-night cycle. Allow mice free access to drinking water and standard laboratory chow. Always maintain sterile conditions during survival surgery.
2. Anesthetize the mouse by intraperitoneal (i.p.) injection of 0.1 ml saline solution per 10 g bw containing 90 mg/kg bw ketamine hydrochloride and 25 mg/kg bw dihydroxylidinothiazine hydrochloride. Anesthesia is necessary for animal preparation, surgery and subsequent microscopy. Confirm proper anesthetization by checking the response to a painful stimulus. During the time of anesthesia, apply vet ointment on eyes to prevent dryness.
3. After induction of sufficient anesthesia, depilate the back with an electric shaver. Hereafter, apply depilation cream to the shaved area to remove the remaining hair.
4. During residence time of the depilation cream of ~7–10 min, prepare instruments, including skin forceps and micro forceps, scissors and micro scissors, suture material and a marker pen. Disinfect and prepare the titanium chamber by applying the two screws with the nut at the chamber’s base and by fixing self-adhesive foam (Fig. 1 A-C) around the window of the chamber’s counterpart to guarantee tightness of the chamber system.
5. Remove all depilation cream from the back by washing it off with tepid water. Then dry and disinfect the skin with an alcohol-containing solution.
6. Place the mouse in prone position and define the midline of the back. Grasp the skin centrally and lift it up to create a fold (Fig. 2A). By trans-illumination using any kind of light source, decide where to provisorily place the titanium frame (Fig. 2B). Make sure that the distal branches of the lateral thoracic artery (LTA) cranially and the deep circumflex iliac artery (DCIA) caudally course through the center of the chamber’s window (Fig. 2B-D). Once the positioning of the chamber is done, perforate both layers of the skin at the very bottom of the fold for later fixation of the titanium frame. Now remove the titanium frame, place the mouse in prone position and redefine the midline of the back if necessary.
7. Outline the flap perpendicular to the spine starting at the midline with a width-to-length ratio of 15 x 11 mm to result in a laterally based and randomly perfused flap. Then outline an additional skin area of 2 mm extending to the contralateral side (Fig. 2C, D). This area is picked with the forceps and later sutured to the titanium frame without interfering with the visible flap area.
8. Following final marking, incise the flap. In doing so, transect both the LTA and DCIA and elevate the flap (Fig. 2E). There is no need for coagulation or ligation of the transected vessels.
9. Place the chamber’s screw through the previously made skin perforations (Fig. 2F) and fixate the skinfold both anteriorly and posteriorly within the chamber’s frame of the elevated flap to the backside part of the frame using the holes of the chamber’s frame and a 5-0 interrupted sutures (Fig. 2G).
10. Suture the vertical limbs of the flap back to the surrounding skin by means of 5-0 interrupted sutures (Fig. 2H, I).
11. Excise a hemi-circular area of skin lateral to the small skin strip of 2 mm, i.e., on the other side of the flap to observe the flap’s vasculature. This allows direct view through the observation window of the chamber that has a surface of 90 mm².
12. Remove the gelatin-like loose areolar tissue from the striated muscle using microsurgical instruments and optical magnifying lens or microscope in order to improve the image quality during microscopy. Try not to damage vessels within the muscular tissue layers.
13. Finally, seal the chamber by mounting the counterpart of the frame that has previously been endued with self-adhering strips of foam anteriorly, cranially and posteriorly, bedew the flap with topical bisbenzimide and saline (Fig. 2J). Thereafter, apply the observation window with a cover glass using a snap ring. Try not to entrap any air blisters under the glass (Fig. 2K, L). The skin will adhere to the cover glass by adhesion forces.

2. Intravital Epifluorescence Microscopy

1. Wait 24 hr after surgical preparation for the first microscopic analysis for ideal optical quality. This analysis represents the baseline of microscopy and is repeated at day 3, 5, 7 and 10 after surgery. Each time, anesthetize the mouse as described in step 1.2. Place the animal in a lateral decubital position on a custom-made Plexiglas carrier. That way, the muscular tissue layers of the flap adhering to the cover glass are facing up-wards.
2. Inject 0.05 ml of 5% green fluorescence dextran (molecular weight 150,000) and 0.05 ml 1% highly fluorescent Rhodamine family dye in a tail vein or the retro-bulbar venous plexus. The green fluorescent dextran stains the non-cellular components of the blood if applied intravenously (i.v.). The fluorescent Rhodamine stains leukocytes and platelets that can be distinguished from other cellular and non-cellular components. Finally, bisbenzimide stains nuclear components that emit more or less fluorescence according to the state of condensation.
3. Subsequently, place the mouse under the microscope. Ensure the microscope includes a LED system — LED beam combinations for 470 & 425 nm, 365 & 490 nm, and 540 & 580 nm as well as the corresponding filter sets (62 HE BFP/GFP/HcRed, range 1: 350–390 nm excitation wavelength, split 395 nm / 402–448 nm; range 2: 460–488 nm, split 495 nm / 500–557 nm; range 3: 567–602 nm, split 610 nm / 615 nm to infinite. 20 Rhodamine, range: 540–552 nm, split 560 nm, emission 575–640 nm).
4. Record the stills and motion-pictures using a charge-coupled device video camera and save them on a computer hard disk for further off-line analysis.
5. Use different objectives (2.5X, NA (numerical aperture) = 0.06 for panorama views of the chamber; 5X, NA = 0.16; 10X, NA = 0.32; 20X, NA = 0.50; 50X, NA = 0.55) for recordings of the regions of interest.
6. Virtually, divide the flap surface that is visible through the chamber’s window into three areas of equal height, i.e., a proximal, a central, and a distal area most remote from the vascular inflow (Fig. 3F). For image acquisition, proceed the following way at each observation time point:
   1. Scan the tissue image by image within the chamber’s window from proximal to distal using the 5X objective.
   2. In each area of the flap, select second- or third-order arterioles and their accompanying venules with easily identifiable branching patterns. Using the 5X, 10X, or 20X objective, make printouts of the arteriolovenular bundles in order to easily re-localize the bundles throughout the whole observation period.
   3. With the 20X objective and the 50X objective, further record 5–6 capillary fields and “apoptotic” fields per area of the flap respectively. All images recorded with the 10X and 20X objectives use the blue light (fluorescent dextran) and green light (Rhodamine) filter, whereas images recorded with the 5X and 50X objective use blue light (fluorescent dextran) and white light (Bisbenzimide) filter, respectively.

3. Analysis of Recorded Data

NOTE: With the use of a computer-assisted image analysis system quantify all recorded parameters off-line as follows²⁹.

1. Measurement of the area of non-perfused, respectively necrotic tissue (mm²).
   1. Use the complete scanned chamber window image recorded with the 5X objective (from 2.6.1) and the fluorescent dextran to measure non-perfused respectively necrotic tissue. Use “AreaBo” function to measure the non-perfused dark area: Border the non-perfused area and calculate the area in mm² by using the software’s Area measurement algorithms.
2. Measurement of microvascular diameter in arterioles, capillaries and venules (µm).
   1. Load video or pictures from the recorded AV-bundles or the capillary fields into the software. For best results use the stills or videos with the highest magnification where definite borders of the vessel are clearly visible.
   2. Use the “DiamPe” function of the software to make sure the measurements are performed perpendicular to the vessel’s wall. To do so mark two points on the vessel’s wall and with the third click mark the perpendicular diameter of the vessel. The software will perform the measurement in µm.
   3. Repeat the measurements on the same vessels for all recordings at the same place. Measure multiple capillaries to accumulate average diameter.
3. Analysis of red blood cell (RBC) velocity in arterioles, capillaries and venules (mm/sec).
   1. For arteriolar red blood cell (RBC) velocity (mm/s) measurements use the “VeloLSD” function of the software and choose the same vessels in the fluorescent dextran window for which the diameters have been measured.
   2. Analyze it with the computer-assisted image analysis system using the line shift method, which is on the basis of the measurement of the shift (mm) of an individual intravascular gray level pattern over time (sec).
4. Measurement of volumetric blood flow in vessels (pl/sec).
   1. Calculate volumetric blood flow (pl/sec) in arterioles, venules and capillaries from RBC velocity and vessel cross-sectional area (π * r²) according to the equation of Gross and Aroesty, that is, Q = V * π * r², assuming a cylindrical vessel shape³⁰.
5. Measurement of functional capillary density of RBC-perfused capillaries (nutritive perfusion: cm/cm²).
   1. Load a recorded fluorescent capillary field with 20X objective magnification into the software. Use the “DensLA” function to measure functional capillary density of RBC-perfused microvessels.
   2. Trace the perfused capillaries with the cursor and mark the perfused vessels. Do not mark non-perfused capillaries, arterioles or venules. The software will measure functional capillary density of the perfused capillaries in cm/cm². Repeat the steps for the other recorded capillary fields.
6. Measurement of tortuosity of the microvessels (microvascular remodeling), early signs of angiogenesis such as bud and sprout-formation and newly formed microvessels.
   1. Load recorded videos or frames of fluorescent capillary fields into the software. Identify gyrose vessels and use the “TorqIx” function of the software. Trace the definite vessel flow with the course and end with a right click. The software will draw a straight line for the direct way and will set it in relation with the traced path.
      NOTE: Watch out for signs of angiogenesis such as buds, sprouts and newly formed capillaries, which usually emanate perpendicularly from the pre-existing capillaries. Measure the density of these newly formed vessels as described in step 3.5.
7. Identification of characteristics for apoptotic cell death, such as nuclear condensation, fragmentation and/or margination (cells/mm²).
   1. Load Bisbenzimide-recorded videos (50X objective) into the software. Use the “DenseNA” function. Mark apoptotic cells according to the mentioned characteristics such as nuclear condensation, fragmentation and margination with a left click.
   2. After marking all characteristic cells throughout the video, click “N/A“ in the software, which will automatically count all marked cells and divide it throughout the area. The result will be apoptotic cells/mm².
8. Analysis of leukocyte adherence to the vascular endothelium representing inflammation. Intermittent adherence of leukocytes (rolling leukocytes; adherence < 30 sec: number of rollers/mm² endothelial surface) and firm adherence of leukocytes (sticking leukocytes; adherence > 30 sec: number of stickers/mm² endothelial surface).
   1. Analyze the inflammatory response by counting the number of Rhodamine labeled leukocytes that adhered to the endothelial lining of post-capillary venules for a time period of ≥30 sec (“stickers”) and the intermittently adhering leukocytes (<30 sec, “rollers”). Venular leukocyte adherence is expressed as cells per mm² endothelial surface.
9. Measurement of intravascular and interstitial grey levels as a parameter for microvascular leakage (macromolecular extravasation E=E1/E2).
   1. Assess macromolecular leakage as a parameter of microvascular permeability after an i.v. injection of a fluorescent dextran. Densitometrically determine multiple grey levels in the tissue directly adjacent to the capillary vessel wall (E1), as well as in the marginal cell-free plasma of the vessel (E2). Then calculate macromolecular extravasation (E) as the ratio of E1/E2 ³¹.

4. Postoperative Care

1. Return the animal in a separate cage to avoid the company of other animals until fully recovered. Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbency. Monitor animals daily for bleeding, local and systemic signs of infection and the position of the chamber.
2. Evaluate the general state of the animal by observing motor activity, body weight, signs of pain, tolerance to the dressing and auto-mutilation.
3. Keep the mice one per cage to avoid mutual manipulation of the chamber. At any sign of pain, apply Buprenorphine at a dose of 0.05–0.1 mg/kg bw, subcutaneously in 8 hr intervals.

5. Euthanasia and Explantation of the Skinfold Chamber

1. At day 10, sacrifice animals using an overdose of an anesthetic drug (150 mg/kg Pentobarbital).
2. Remove the chamber and sample the flap tissue for immunohistochemical and molecular protein assays. Sample tissues for conventional histology (formalin) and quantifiable protein assays (liquid nitrogen or dry ice).

Results

Necrosis

The main endpoint of this model — tissue necrosis following flap elevation (i.e., induction of acute persistent ischemia) — is repeatedly measured and illustrated macroscopically as shown in Figure 3 over a period of 10 days. Final demarcation of flap necrosis usually occurs between day 5 and 7 after surgery and is characterized by a red fringe, i.e., zone of vasodilation and microvascular remodeling, developing betw...

Discussion

In order to decrease ischemic complications and thereby improve the clinical outcome, more detailed knowledge of pathophysiologic processes in critically perfused flap tissue is required. The development of new animal models that mimic acute persistent ischemia is therefore mandatory. Accordingly, we were able to develop an easily reproducible and reliable model allowing for repetitive morphological, dynamic and functional real-time evaluation of various parameters of muscle and skin vasculature that can be correlated wi...

Materials

Name	Company	Catalog Number	Comments
Name of Reagent/ Equipment	Company	Catalog Number	Comments/Description
C57Bl/6 mice 6-8w 20-22g	Charles River
depilation cream	Veet		any depilation cream
titanium chamber	Irola	160001	Halteblech M
slotted cheese head screw	Screws and More	842210	DIN84 M2x10
hexagon full nut	Screws and More	93422	DIN934 M2
snap ring	Schaefer-Peters	472212	DIN472 J12x1,0
cover glass	Volab		custom-made cover glass 11,8mm in diameter
fixing foam	tesamoll	05559-100	tesamoll Standard I-Profile
ketamine hydrochloride	Parke Davis		Ketavet®
dihydroxylidinothiazine hydrochloride	Bayer		Rompun®
Buprenorphin	Essex Pharma		Temgesic®
Saline 0,9%
desinfection alcohol
Vicryl 5-0	Ethicon	V 490 H
Ethilon 5-0	Ethicon	EH 7823 H
1ml syringes
surgical skin marker with flexible ruler	Purple surgical	PS3151	any surgical skin marker and flexible ruler
pointed scissors
Micro-Scissors
normal scissors
2 clamps
fine anatomic forceps
micro-forceps
hex nuter driver	wiha	1018
screwdriver	wiha	685
snap ring plier	Knipex	4411J1	12-25mm
wire cutter	Knipex	70 02 160	Wire cutter is used to cut screws short; 160mm
trans-illumination light	IKEA	501.632.02	LED light Jansjö; any light
magnification glasses
intravital microscope	Zeiss	490035-0001-000	Scope.A1.Axiotech
LED system	Zeiss	423052-9501-000	Colibri.2
LED module 365nm	Zeiss	423052-9011-000
LED module 470nm	Zeiss	423052-9052-000
LED module 540-580nm	Zeiss	423052-9121-000
Filter set 62 62 HE BFP + GFP + HcRed	Zeiss	489062-9901-000	range 1: 350-390nm excitation wavelength split 395 / 402-448nm; range 2: 460-488nm, split 495nm / 500-557nm; range 3: 567-602nm, split 610nm / 615-infinite
Filter set 20 Rhodamine	Zeiss	485020-0000-000	540-552nm, split 560, emission 575-640nm
2,5x objective NA=0,06	Zeiss	421020-9900-000	A-Plan 2,5x/0.06
5x objective NA=0,16	Zeiss	420330-9901-000	EC Plan-Neofluar 5x/0.16 M27
10x objetive NA=0,30	Zeiss	420340-9901-000	EC Plan-Neofluar 10x/0.30 M27
20x objective NA=0.50	Zeiss	420350-9900-000	EC Plan-Neofluar 20x/0.50 M27
50x objective NA=0,55	Zeiss	422472-9960-000	LD Epiplan-Neofluar 50x/0.55 DIC 27
ZEN imaging software	Zeiss		ZenPro 2012
CapImage	Dr. Zeintl
Fluorescein isothiocyanate-dextran	Sigma-Aldrich	45946
bisBenzimide H 33342 trihydrochloride	Sigma-Aldrich	B2261	harmful if swallowed; causes severe skin burns and eye damage, may cause repiratory irritat
Rhodamine 6G chloride	Invitrogen	R634	harmful if swallowed; may cause genetic defects; may cause cancer; may damage fertility or the unborn child
Pentobarbital	Merial		Narcoren®