A Murine Tail Lymphedema Model

Summary

Lymphedema is extremity swelling caused by lymphatic dysfunction. We describe a chronic murine tail model of lymphedema and the novel use of tissue nanotransfection technology (TNT) for genetic cargo delivery to the tail.

Abstract

Lymphedema is extremity swelling caused by lymphatic dysfunction. The affected limb enlarges because of accumulation of fluid, adipose, and fibrosis. There is no cure for this disease. A mouse tail model that uses a focal full thickness skin excision near the base of the tail, resulting in tail swelling, has been used to study lymphedema. However, this model may result in vascular comprise and consequent tail necrosis and early tail swelling resolution, limiting its clinical translatability. The chronic murine tail lymphedema model induces sustained lymphedema over 15 weeks and a reliable perfusion to the tail. Enhancements of the traditional murine tail lymphedema model include 1) precise full thickness excision and lymphatic clipping using a surgical microscope, 2) confirmation of post-operative arterial and venous perfusion using high resolution laser speckle, and 3) functional assessment using indocyanine green near infrared laser lymphangiography. We also use tissue nanotransfection technology (TNT) for novel non-viral, transcutaneous, focal delivery of genetic cargo to the mouse tail vasculature.

Introduction

Lymphedema is extremity swelling caused by lymphatic dysfunction. The affected limb enlarges because of the accumulation of fluid, adipose, and fibrosis¹. Lymphedema affects 250 million people worldwide^2,3,4. It is estimated that 20-40% of patients who undergo treatment for solid malignancies, such as breast cancer, melanoma, gynecological/urologic tumors, or sarcomas, develop lymphedema^2,4,5. Morbidity from lymphedema includes recurrent infections, pain, and deformity⁶. There is no cure for this progressive, life-long disease. Current therapies are variaby effective⁷ and include compression, complete decongestive therapy by physical therapists, excisional procedures, and microsurgical operations, including vascularized lymph node transfer and lymphovenous bypass^{7,8,9,10,11,12,13,14}. The ideal treatment for lymphedema has yet to be discovered.

Studying the mechanism and therapy of lymphedema has been limited. There is an average delayed onset of one year following the lymphatic injury^15,16 and most individuals who experience iatrogenic insult with radiation and surgery do not develop lymphedema^4,6,17. Although large animal models, including canine, sheep, and pig have been described^18,19,20, the mouse tail model has been the most widely applied because of ease, cost, and reproducibilty. Mouse models for investigating lymphedema include a tail model, diptheria-toxin mediated lymphatic ablation, and axillary or popliteal lymph node dissection^{21,22,23,24,25,26}. Most tail models use a focal, full thickness skin excision with lymphatic channel clipping that is performed near the base of the tail²², resulting in tail swelling and histological features similar to human lymphedema^24,27,28,29. However, the standard murine tail model typically spontaneous resolves in as few as 20 days and is accompanied by periodic tail necrosis³⁰. The lymphedema mouse tail model extends a sustained lymphedema beyond 15 weeks, demonstrates confirmed arterial and venous patency, and allows functional lymphatic dysfunction assessment.

An murine tail model of lymphedema allows for evaluation of novel therapeutics to treat lymphedema. Gene-based strategies have been used in the mouse model mediated by viral vectors^31,32. We also use a novel tissue nanotransfection technology (TNT) for genetic cargo delivery to the lymphedematous mouse tail. TNT facilitates direct, transcutaneous gene delivery using a chip with nanochannels in a rapid focused electric field^33,34,35,36. The model includes using TNT_2.0 to allow for focal gene delivery of potential gene-based therapeutics to the lymphatic injury site of the mouse tail³⁵.

Protocol

The protocol follows the guidelines of the institution's animal research ethics committee. All animal experiments were approved by the Indiana University School of Medicine Institutional Animal Care and Use Committee. Animals were housed under a 12-hour light-dark cycle with food and water ad libitum.

1. Surgical Disruption of Mouse Tail Lymphatics

1. Use eight week-old C57BL/6 mice of equal gender distribution.
2. Place a mouse under general anesthesia in an induction chamber with 3-4% isoflurane in 100% oxygen followed by maintenance sedation at 1-3% during the procedure.
3. Administer 0.5 mg/kg sustained-release (SR) buprenorphine subcutaneously for pain control.
   NOTE: Additional analgesic drugs administered post-op: Carprofen once every 24 h for at least 48 h and Bupivacaine once either after the incision was made or before closing the incision, applied by dripping onto skin edges (lasts up to 4 – 6 h).
4. Position the mouse dorsally and prep the tail with 70% isopropyl alcohol.
5. Measure the tail diameter prior to the procedure at 5 mm increments starting 20 mm from the base of the tail using a caliper. These measurements will be used to calculate volume using the truncated cone equation³⁷.
6. Mark a 3 mm circumferential excision on the tail 20 mm from the base.
7. Perform a meticulous 3 mm full-thickness skin excision with a sterile surgical blade (size 15), leaving all the underlying vasculature intact under surgical microscopic magnification. Incise the superior circumferential mark (20 mm from tail base) first through the dermis followed by a circumferential full thickness incision 3 mm distlal to the first incision.
   1. Make a perpendicular full thickness vertical incision to connect the two incisions. Use a toothed fine pickup to grasp a leading edge and use microscissors to carefully dissect deep within the avascular plane to the dermis and superficial to the vein adventitia.
8. Inject 0.1 mL of isosulfan blue (1%) subcutaneously proximal to tip of the tail.
9. Identify the two lymphatic channels adjacent to the lateral tail veins under the surgical microscope. The lymphatics will appear blue because of isosulfan injection. Transect the lymphatics using straight microsurgical scissors. Use the scissors to carefully dissect a plane between the lateral vein and the lymphatic. Then pass the tip of one scissor blade between the lymphatic vessel and the lateral vein and close the blades to transect the lymphatic vessel.
10. Dress the tail wound with a sterile adherent clear dressing. Check post-op incisions daily to ensure that they are not infected or bleeding and provide wound care for 2 weeks. 
11. House the animals singly to prevent any further injury to the tail and to prevent the animals from biting each other, which would lead to surgical complications. 

2. Tail vascular assessment with laser speckle contrast imaging

1. Anesthetize the mouse as in step 1.2.
2. To use laser speckle contrast imaging to visualize tail vascularity, set the width to 0.8 cm, height to 1.8 cm, point density to high, frame rate to 44 images/second, time to 30 seconds, and color photo to 1 per 10 seconds.
3. Evaluate the venous and arterial perfusion for patency. Qualitatively, continuity of flow should be visualized.

3. Functional lymphatic evaluation with near infrared laser angiography

1. Anesthetize the animal as in step 1.2
2. Reconstitute indocyanine green (ICG) (25 mg/10 mL) and administer 0.1 mL subcutaneously into the distal mouse tail near the tip.
3. Dim the room lights. Place near-infrared laser angiography in buffering setting followed by live capture.

4. Focal delivery of nucleic acid cargo to mouse tail using TNT

1. Anesthetize the animal as in step 1.2.
2. Exfoliate the mouse tail using topical skin exfoliation cream.
3. Immerse the mouse tail in collagenase solution (10 mg/mL) at 37 °C for 5 minutes.
4. Load DNA into the TNT_2.0 chip reservoir³⁵.
5. Place the TNT_2.0 silicone chip device over the desired focal site of delivery on the tail with nanoneedles in contact with the tail.
6. Place a positive electrical probe in the reservoir. Attach the negative probe to a 30 G needle and insert the needle subcutaneously into the tail to the site of delivery.
7. Apply square wave pulse electric stimulation (10 x 10 ms pulses, 250 V, 10 mA).

Results

The technique for the mouse tail model for sustained lymphedema is shown in Figure 1. The figure exhibits the relevant anatomy of the mouse tail model. Figure 2 demonstrates the progressive swelling and sustained persistant lymphedema in the mouse tail after lymphedema induction. The mouse tail volume, as calculated by the truncated cone equation, peaks at week 4 and plateaus to week 6 followed by gradual improvement that is sust...

Discussion

Lymphedema is categorized as a primary (congenital) or secondary (iatrogenic lymphatic) injury^38,39. Secondary lymphedema comprises 99% of cases³⁹. Secondary lymphedema is most commonly caused by infection (filariasis) or post-oncological treatment with lymphadenectomy or radiation^4,39. A translational animal model is challenging for secondary lymphedema, as 70% of animals treated ...

Materials

Name	Company	Catalog Number	Comments
Surgical Microscope	Leica, Wetzlar, Germany	MSV266
Adherent Dressing (Tegaderm)	3M, St. Paul, Minn.	1626W
Laser speckle (Pericam PSI System )	Perimed AB, Stockholm, Sweden)	PSIZ
Near-infrared laser (LUNA)	Stryker (Formerly Novadaq Technologies, Toronto, Canada)	LU3000
C57BL/6 mice	Jackson Laboratories	000664
Micro-Adson Forceps - 1x2 Teeth	Fine Science Tools (USA) Inc.	11019-12
V-Hook	Fine Science Tools (USA) Inc.	18052-12
Scalpel SS NO15	Fischer Scientific	29556
Disposable Needle 30GX1	Fischer Scientific	305128
Operating Scissors	Fischer Scientific	12-460-796
Surgi-Or Jeweler's Forceps, Sklar 4-1/2 in	Fischer Scientific	50-118-4255
Spring Scissors - Straight/Sharp-Sharp/8mm Cutting Edge	Fine Science Tools (USA) Inc.	15024-10
Cardiogreen	Sigma	I2633-25MG
IsosulfanBlue (Lymphazurin)  50 mg/5ml	Mylan	67457-220-05