Balloon-based Injury to Induce Myointimal Hyperplasia in the Mouse Abdominal Aorta

Summary

This article demonstrates a murine model to study the development of myointimal hyperplasia (MH) after aortic balloon injury.

Abstract

The use of animal models is essential for a better understanding of MH, one major cause for arterial stenosis.In this article, we demonstrate a murine balloon denudation model, which is comparable with established vessel injury models in large animals. The aorta denudation model with balloon catheters mimics the clinical setting and leads to comparable pathobiological and physiological changes. Briefly, after performing a horizontal incision in the aorta abdominalis, a balloon catheter will be inserted into the vessel, inflated, and introduced retrogradely. Inflation of the balloon will lead to intima injury and overdistension of the vessel. After removing the catheter, the aortic incision will be closed with single stiches. The model shown in this article is reproducible, easy to perform, and can be established quickly and reliably. It is especially suitable for evaluating expensive experimental therapeutic agents, which can be applied in an economical fashion. By using different knockout-mouse strains, the impact of different genes on MH development can be assessed.

Introduction

Arterial stenosis in coronary and peripheral arteries has a large effect on the morbidity and mortality of patients¹. One underlying pathological mechanism is myointima hyperplasia (MH), which is characterized by increased proliferation, migration, and synthesis of extracellular matrix proteins from vascular smooth muscle cells (SMC)². SMC are located in the media layer of the vessel and migrate upon stimulation to the surface of the lumen. Stimulatory signals include growth factors, cytokines, cell-cell contact, lipids, extracellular matrix components, and mechanical shear and stretch forces^3,4,5,6. Injuries of the vessel wall, pathological or iatrogenic, cause endothelial cell and smooth muscle cell damage and stimulate inflammatory reactions, and thus lead to MH⁷.

Different animal models are currently available to study arterial injury and myointima hyperplasia. Large animals like pigs or dogs have the advantage of sharing a similar artery and coronary anatomy with humans and are especially suitable for studies investigating angioplasty techniques, procedure, and devices⁸. However, pig models have the drawback of higher thrombogenicity^9,10, while dogs only have a mild response to vessel injury¹¹. In addition, all large animal models require special housing, equipment, and staff, which are connected with high costs and are not always available at an institution. Small animal models include rats and mice. Compared to rats, mice have the advantages of lower cost and the existence of a variety of knock out models. The model described in this video can be combined with ApoE-/- mice fed with a western diet to closely mimic the clinical setting of angioplasty in atherosclerotic vessels¹². Previous models induced vascular injury via wire injury¹³, fluid desiccation¹⁴, spring¹⁵, or cuff injury¹⁶. Since the nature of the injury will greatly impact the development and constitution of MH, using a balloon catheter to induce vessel injury is the best way to mimic the clinical setting.

In this article we describe a novel method to induce MH with a balloon catheter in mice. The use of a balloon catheter (1.2 mm x 6 mm) with a RX-Port (Figure 1A) allows the scraping of the intimal layer and, at the same time, the induction of an overdistension of the vessel. Both of these factors are important triggers for the development of MH. The observation time for this model is 28 days¹⁷.

Protocol

Animals received humane care in compliance with the Guide for the Principles of Laboratory Animals, prepared by the Institute of Laboratory Animal Resources, and published by the National Institutes of Health. All animal protocols were approved by the responsible local authority (''Amt für Gesundheit und Verbraucherschutz, Hansestadt (Office for Health and Consumer Protection) Hamburg'').

1. Catheter Preparation

NOTE: Refer to the Table of Materials for information regarding the catheter.

1. Take catheter from the catheter holder.
2. Pull the guidewire from the lumen through the distal port out.
3. Put in a drop of cyanoacrylate adhesive on the distal end of catheter.
   Caution: Wear latex or nitrile gloves.
4. Place the guidewire at the RX port of the catheter and advance it through the lumen to the distal port. Afterwards, pull the guidewire back slightly to leave a space of ~5 mm to the end.
5. Wait 5 min to allow the adhesive to dry.
6. Move the guidewire, it should be fixed. If it is still mobile, repeat steps 1.3 - 1.6.
7. Fill a 3-mL syringe with 1.5 mL 0.9% saline and connect it to the balloon inflation port.
8. Push the syringe plunger to test balloon inflation. Leave 0.6 mL saline in the syringe.

2. Mouse preparation

1. Obtain male mice at the age of 14 weeks weighing approximately 30 g.
   NOTE: We used animals obtained from the Institute of Laboratory Animals. We used C57BL/6J here.
2. Use an induction chamber to anaesthetize the mouse with 2.0-2.5% isofurane (500 mL/min oxygen flow rate).
3. Place the mouse on its back on a heating pad and maintain anesthesia with a facemask covering the mouth and nose of the mouse. Check for sufficient depth of anesthesia by pinching the hind feet and tail to verify an absence of reflexes.
4. Remove the abdominal hair using a hair trimmer.
5. Spread the hind legs and fix their position using tape.
6. Disinfect the abdominal area using povidone-iodine, followed by 80% ethanol. Repeat this step two more times.
7. Use a surgical drape to ensure surgical site does not get contaminated. Open the skin and muscle layers along the linea alba with a scissor (or scalpel) to expose the abdominal organs.
8. Lay the intestines in a 0.9% saline moisturized glove and wrap to keep them moist.
9. Use two fine forceps to remove the fatty tissue above the abdominal aorta.
10. Using an insulin syringe (30G), inject 250 µL of heparin solution (50 U/mL) into the Inferior Vena Cava (IVC) and wait 3 min for its systemic distribution. Heparin will suppress hemostasis and prevent undesired clotting during surgery.
11. Using two forceps, dissect the infrarenal aorta down to its bifurcation and its outgoing branch vessels.
12. Ligate the side vessels, which are expected to be placed in the clamped area, with a high temperature cauterizer.
13. Stop blood flow by clamping the infrarenal aorta directly beneath the renal arteries.
14. Place a second clamp at a distal position just above the aortic bifurcation.
15. Perform a small horizontal incision using scissors at the midpoint between the clamps along the vessel.
    NOTE: The size of the incision should be equivalent to 1/3 of the circumference of the vessel.
16. Insert a syringe with a (30G) needle into the incision and flush the aorta with 250 µL heparin solution (50 U/mL).
17. Using 10-0 sutures, place one single knot on each side of the incision.
18. Dilate the aorta by inserting a vessel dilator into the incision and spread the vessel slightly. Repeat the dilation 2 to 3 times.
19. Moisturize the balloon-catheter with 0.9% saline.
20. Insert the flattened balloon-catheter into the aorta and advance it towards the proximal clamp on the aorta.
21. When reaching the proximal clamp, carefully open the proximal clamp and inflate the balloon to prevent blood leakage, by injecting ~0.6 mL of saline.
    NOTE: The ratio of the inflated balloon to vessel is 1.5:1.
22. Advance the catheter retrograde for approximately 2 cm.
23. Pull the expanded catheter back, while deflating the balloon slightly by releasing the syringe.
24. Reattach the proximal clamp when the catheter reaches the incision of the aorta. Deflate the balloon completely and remove it.
25. Rinse the aorta with 250 µL heparin solution (50 U/mL) using a 30G syringe.
26. Close the aortic incision using 10-0 sutures. Place interrupted stitches on each lateral side, followed by one or two stitches on the ventral side.
27. Open the distal clamp. In case of bleeding, close the clamp again and place additional stitches.
28. Open the proximal clamp carefully.
29. Place two swabs on the suture to support it and stop any bleeding.
30. Place absorbable hemostats on the suture to sustain it.
    NOTE: An aortic pulse should be visible distally from the incision.
31. Place the intestines back into the abdomen.
32. Rinse the abdominal cavity with sterile 0.9% saline, which has been pre-warmed to 37 °C.
33. Close the abdominal muscle layer using 6-0 running sutures.
34. Close the skin with 5-0 running sutures.
35. Inject 4-5 mg/kg Carprofen subcutaneously before allowing the mouse to wake up. Monitor the animal until it has gained consciousness, maintain sternal recumbency. Keep the animal alone in a cage until complete recovery.
36. Add Metamizole to the drinking water (50 mg/100 mL) as pain medication for 3 days and monitor the animal daily. Usually mice like the sweet taste of metamizole and start drinking immediately after the surgery. If preferred, sustained-release injectable agents can be used instead of metamizole.
    NOTE: The observation period for this model is 28 days.

3. Histopathology

1. Harvest the balloon-injured aorta after 28 days by preparing the mice as described in steps 2.2 to 2.9.
2. Use scissors to remove the balloon-injured aorta (between the bifurcation and 0.3 mm above the renal vessels) and euthanize the mouse by cutting out its heart.
3. Flush the lumen of the vessel with 0.9% NaCl.
4. Fix the harvested vessel in 4% paraformaldehyde (PFA) overnight and dehydrate it in increasing concentrations of ethanol, starting with 70% ethanol for 2 hours, 80% ethanol for 1 hour, 95% ethanol for 2 hours, and 100% ethanol for 5 hours. Then, incubate the samples in xylene for 2 hours 3 times, before infiltrating the samples with paraffin.
   NOTE: Instead of flushing the harvested vessel with 0.9% NaCl, it can be flushed with 4% PFA.
   Caution: PFA and xylene are toxic and should be handled with special care.
5. Embed the sample in paraffin and cut into slices of 5 µm thickness using a microtome.
6. Deparaffinize the slides with xylene 3 times for 5 minutes.
7. Rehydrate tissue slides using a decreasing series of ethanol. Start with 100% ethanol 2 times for 5 min, followed by 3 minutes of 95%, 80% and 70% ethanol.
8. Stain the slides with Masson's trichrome staining as described¹⁸.
9. Dehydrate stained slides in 100% ethanol 2 times for 10 min each. Clear with xylene 2 times for 10 min each and mount in mounting medium.
10. View the slides with a bright field microscope. Use a lens with 5x magnification and a numerical aperture of 0.12 for an overview picture or a lens with 20x magnifications with a numerical aperture of 0.35 for detailed observation.

4. Immunofluorescence Microscopy

1. Rehydrate tissue slides using a decreasing series of ethanol. Start with 100% ethanol 2 times for 5 min, followed by 3 min of 95%, 80% and 70% ethanol.
2. Perform antigen-retrieval by heating the slides in antigen-retrieval solution in a steamer for 20 min.
3. Let the slides cool down to room temperature.
4. After washing the slides for three times with phosphate buffered saline (PBS), apply antigen blocking solution on sections for 30 min.
5. Wash slides three times for 5 min with PBS.
6. Incubate sections with primary antibody diluted in primary antibody diluent.
   NOTE: The right concentration and incubation time should be chosen separately for each antibody.
7. Wash slides three times for 5 min with PBS to remove unbound antibody.
8. Incubate sections with a pre-conjugated secondary antibody diluted in secondary antibody diluent.
   NOTE: The right concentration and incubation time should be chosen separately for each antibody.
9. Remove unbound antibodies by washing the slides for 5 min three times.
10. Counterstain the cell nuclei using 4',6-diamidino-2-phenylindole (DAPI) for 15 min; the final DAPI concentration should be 350 nM.
11. Mount slides in immunofluorescence compatible mounting solution.
    NOTE: Using the wrong mounting solution can obscure the fluorescence signal.
12. View slides with a fluorescence microscope. Use a 40x magnification lens with a numerical aperture of 1.3.

Results

Balloon denudation is a suitable model to study the development of MH in mice. Animals recover well from the surgery and show an excellent physical condition post-operation. We established this model in 50 mice with less than 3% death rate due to the surgical procedure. Figures 1B-C show the key surgical steps. After a skin incision along the linea alba, identify the aorta abdominalis. Place microsurgical clamps (

Discussion

This article demonstrates a murine model to study the development of myointimal hyperplasia and allows the exploration of the underlying pathological processes and the testing of new drugs or therapeutic options.

The most critical step in this protocol is the denudation of the aorta. Special care should be paid during this step as excessive denudation will lead to aneurysm formation and model failure. On the other hand, if denudation is performed insufficiently, too little myointima will devel...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
10-0 Ethilon suture	Ethicon	2814G
3 mL Syringe	BD Medical	309658
37% HCl	Sigma-Aldrich	H1758
5-0 prolene suture	Ethicon	EH7229H
6-0 prolene suture	Ethicon	8706H
Acid Fuchsin	Sigma-Aldrich	F8129-25G	Trichrome staining
Antigen retrieval solution	Dako	S1699
Azophloxin	Waldeck	1B-103	Trichrome staining
Bepanthen Eye and Nose ointment	Bayer	1578675	Eye ointment
Betadine Solution	Betadine Purdue Pharma	NDC:67618-152
C57BL/6J	Charles River	Stock number 000664
Clamp applicator	Fine Science Tools	18056-14	JAW DIMS: 4 x 0.75 mm LENGTH: 13 mm
Collagen 3	abcam	ab7778	Antibody
DAPI	Thermo Fischer	D1306
Donkey anti-Goat IgG AF555	Invitrogen	A21432	Secondary antibody
Donkey anti-Rabbit IgG AF488	Invitrogen	A21206	Secondary antibody
Donkey anti-Rabbit IgG AF488	Invitrogen	A11055	Secondary antibody
Donkey anti-Rabbit IgG AF555	Invitrogen	A31572	Secondary antibody
Ethanol 70%	Th. Geyer	2270
Ethanol 96%	Th. Geyer	2295
Ethanol absolute	Th. Geyer	2246
FAP	abcam	ab28246	Antibody
Forceps fine	Fine Science Tools	11251-20
Forceps standard	Fine Science Tools	11023-10
Glacial Acetic Acid	Sigma-Aldrich	537020
Hair clipper	WAHL	8786-451A ARCO SE
Heparin	Rotexmedica	PZN 3862340	25.000 I.E./mL
High temperature cautery kit	Bovie	18010-00
Image-iT FX Signal Enhancer	Invitrogen	I36933	Blocking solution
Light Green SF	Waldeck	1B-211	Trichrome staining
Microsurgical clamp	Fine Science Tools	18055-04	Micro-Serrefine - 4mm
MINI TREK Coronary Dilatation Catheter 1.20 mm x 6 mm / Rapid-Exchange	Abbott	1012268-06U
Molybdatophosphoric acid hydrate	Merck	1.00532.0100	Trichrome staining
NaCl 0,9%	B.Braun	PZN 06063042 Art. Nr.: 3570160
Needle holder	Fine Science Tools	12075-14
Novaminsulfon	Ratiopharm	PZN 03530402	Metamizole
Orange G	Waldeck	1B-221	Trichrome staining
Paraffin	Leica biosystems	REF 39602004
PBS pH 7,4	Gibco	10010023
PFA 4%	Electron Microscopy Sciences	#157135S
Ponceau S solution	Serva Electrophoresis	33427	Trichrome staining
Primary antibody diluent	Dako	S3022
Prolong Gold Mounting solution	Thermo Fischer	P36930	Mounting solution for immunofluorescence stained slides
Replaceable Fine Tip	Bovie	H101
Resorcin-Fuchsin Weigert	Waldeck	2E-30	Trichrome staining
Rimadyl	Pfizer	400684.00.00	Carprofen
Scissors	Fine Science Tools	14028-10
Scissors Vannas-style	Fine Science Tools	15000-03
Secondary antibody diluent	Dako	S0809
Fast acting Adhesive MINIS 3x1g	UHU	45370	Cyanoacrylate
Slide Rack	Ted Pella	21057
SM22	abcam	ab10135	Antibody
SMA	abcam	ab21027	Antibody
Staining dish	Ted Pella	21075
Surgical microscope	Leica	M651
Tabotamp fibrillar	Ethicon	431962	Absorbable hemostat
Transpore Surgical Tape	3M	1527-1
U-100 Insulin syringe	BD Medical	324825
Vessel Dilator	Fine Science Tools	18603-14
Vitro-Clud	Langenbrinck	04-0001
Weigerts iron hematoxylin Kit	Merck	1.15973.0002	Trichrome staining
Xylene	Th. Geyer	3410