This work was supported by grant funding provided by the American Association of Plastic Surgeons Academic Scholarship and the Department of Defense W81XWH2110135&#160;to AHH. Aesthetic Surgery Education and Research Foundation grant to MS. NIH U01DK119099, R01NS042617 and R01DK125835 to CKS.

The protocol follows the guidelines of the institution&#39;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
<ol>
	<li>Use eight week-old C57BL/6 mice of equal gender distribution.</li>
	<li>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.</li>
	<li>Administer 0.5 mg/kg sustained-release (SR) buprenorphine subcutaneously for pain control. 
	NOTE: Additional analgesic drugs administered post-op: Carprofen&#160;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 &#8211; 6 h).</li>
	<li>Position the mouse dorsally and prep the tail with 70% isopropyl alcohol.</li>
	<li>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.</li>
	<li>Mark a 3 mm circumferential excision on the tail 20 mm from the base.</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Inject 0.1 mL of isosulfan blue (1%) subcutaneously proximal to tip of the tail.</li>
	<li>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.</li>
	<li>Dress the tail wound with a sterile&#160;adherent clear dressing.&#160;Check post-op incisions daily to ensure that they are not infected or bleeding and provide wound care for 2 weeks.&#160;</li>
	<li>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.&#160;</li>
</ol>
2. Tail vascular assessment with laser speckle contrast imaging
<ol>
	<li>Anesthetize the mouse as in step 1.2.</li>
	<li>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.</li>
	<li>Evaluate the venous and arterial perfusion for patency. Qualitatively, continuity of flow should be visualized.</li>
</ol>
3. Functional lymphatic evaluation with near infrared laser angiography
<ol>
	<li>Anesthetize the animal as in step 1.2</li>
	<li>Reconstitute indocyanine green (ICG) (25 mg/10 mL) and administer 0.1 mL subcutaneously into the distal mouse tail near the tip.</li>
	<li>Dim the room lights. Place near-infrared laser angiography in buffering setting followed by live capture.</li>
</ol>
4. Focal delivery of nucleic acid cargo to mouse tail using TNT
<ol>
	<li>Anesthetize the animal as in step 1.2.</li>
	<li>Exfoliate the mouse tail using topical skin exfoliation cream.</li>
	<li>Immerse the mouse tail in collagenase solution (10 mg/mL) at 37 &#176;C for 5 minutes.</li>
	<li>Load DNA into the TNT2.0 chip reservoir35.</li>
	<li>Place the TNT2.0 silicone chip device over the desired focal site of delivery on the tail with nanoneedles in contact with the tail.</li>
	<li>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.</li>
	<li>Apply square wave pulse electric stimulation (10 x 10 ms pulses, 250 V, 10 mA).</li>
</ol>

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The authors have no competing conflicts of interest.

Lymphedema is categorized as a primary (congenital) or secondary (iatrogenic lymphatic) injury38,39. Secondary lymphedema comprises 99% of cases39. Secondary lymphedema is most commonly caused by infection (filariasis) or post-oncological treatment with lymphadenectomy or radiation4,39. A translational animal model is challenging for secondary lymphedema, as 70% of animals treated with lymphadectomy and radiation do not acquire lymphedema2,16. In addition, phenotypical lymphedema exhibits a delayed onset (one year) post-lymphatic injury. The mouse tail model of lymphedema overcomes these obstacles, as all mice undergoing focal tail lymphatic excision exhibit lymphedema within days following the procedure21,23. Full thickness focal subcutaneous excision is performed under visualization with the surgical microscope allowing definitive identification of the tissue plane between the tail veins and the subcutaneous tissues and facilitating vessel preservation. We have previously ligated the lymphatic channel with nylon suture, but as persistent lymphedema can be induced with transection of the lymphatic channel only, ligation must be deemed unnecessary. The two lateral lymphatic channels in the mouse tail are in close proximity with the lateral tail veins. Histologically, the swollen tail displays inflammation, intersitial fluid retention, adipose deposition, and fibrosis, similar to clinical lymphedema24,27,28,40.
One pitfall of this model is the risk of injury to the lateral veins and vasculature. Performing the procedure on full thickness skin excision using loupe magnification can lead to inadvertent venous bleeding during dissection. Careful excision under high stereoscopic magnification facilitates greater precision for staying within the a vascular plane between the vessel adventitia and subdermal layer. Another difficulty is that tail necrosis occurs with a frequency as high as 30%30, as vessel injury greatly increases tail necrosis risk. The model marginalizes tail necrosis with (1) the use of a surgical microscope for meticulous dissection and (2) the confirmation of vessel patency by laser speckle imaging41. If vascular injury is identified, the animal should be removed from the study. Other investigators have used intracardiac microsphere injection to assess arterial perfusion22. Laser speckle imaging allows quantification of the blood flow kinetics of veins in addition to arteries41. This minimally invasive technique can provide precise microperfusion data.41
The tail volume is used as a phenotypic outcome variable of the model. Assessing lymphatic function of the tail in the model also is used to assess the experimental effect. We use near infrared laser lymphangiography to evaluate lymphatic function in the mouse tail. This directly visualizes real time lymphatic flow in the live animal. ICG laser lymphangiography is also commonly used clinically during lymphatic microsurgical threapeutic procedures such as lymphovenous anastomosis so it translates well10. Clinically, this facilitates introperative lymphatic mapping and the identification of target lymphatic vessels to connect them into veins in lymphovenous anatomosis to treat lymphedema7,10. One pitfall of using ICG laser lymphangiography is the ease with which the mouse tail and other materials can become coated with ICG, resulting in non-speciic fluoresence and hindering proper visualization of the lymphatics. Therefore, we change gloves immediately both after ICG handling and administration to minimize this risk.
TNT was developed initially for in vivo tissue reprogramming33. It is used as a gene transfer platform, more broadly including the rescue of diabetic peripheral neuropathy and the repair of crushed nerves34,36 and utilizes three essential components: (1) a silicone nanochip for nanoneedle based gene transfer; (2) a nucleic acid cargo&#160;(plasmids with ORF or siRNAs); and (3) a standard power supply. TNT facilitates direct, transcutaneous, non-viral gene delivery with a rapid focused electric field. It has been used to decrease limb ischemia by increasing neovascularization in a mouse model33.&#160;More recently, TNT2.0 has been used to label wound-site exosomes35. Using TNT in the mouse tail lymphedema model offers an exciting future for the delivery of gene-based therapies.
A translational limitation of the mouse tail lymphedema model has been the spontaneous resolution of lymphedema21,22, as tail swelling resolves after 20-30 days in some experimental models21. In the model, tail swelling volume, as measured by the commonly used truncated cone equation37, has been sustained for 15 weeks without exhibiting resolution. Perhaps the technique enhancements have maximized persistence of lymphedema. The technique modifications include complete dissection under microcopic magnification, laser speckle evaluation of tail vasculature to ensure rigor for lymphatic origin of lymphedema, functional assessment with ICG laser lymphangiography, and TNT2.0 for therapeutic gene delivery. The modified mouse tail model of lymphedema is a reproducible and clinically translatable animal model of lymphedema.

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 worldwide2,3,4. 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 lymphedema2,4,5. 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 bypass7,8,9,10,11,12,13,14. 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 injury15,16 and most individuals who experience iatrogenic insult with radiation and surgery do not develop lymphedema4,6,17. Although large animal models, including canine, sheep, and pig have been described18,19,20, 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 dissection21,22,23,24,25,26. 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 lymphedema24,27,28,29. 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 vectors31,32. 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 field33,34,35,36. The model includes using TNT2.0 to allow for focal gene delivery of potential gene-based therapeutics to the lymphatic injury site of the mouse tail35.

<table><tbody><tr><td>Tegaderm Film</td><td></td><td>1626W</td><td></td></tr><tr><td>Surgical Microscope</td><td>Leica, Wetzlar, Germany</td><td>MSV266</td><td></td></tr><tr><td>Adherent Dressing (Tegaderm)</td><td>3M, St. Paul, Minn.</td><td>1626W</td><td></td></tr><tr><td>Laser speckle (Pericam PSI System )</td><td>Perimed AB, Stockholm, Sweden)</td><td>PSIZ</td><td></td></tr><tr><td>Near-infrared laser (LUNA)</td><td>Stryker (Formerly Novadaq Technologies, Toronto, Canada)</td><td>LU3000</td><td></td></tr><tr><td>C57BL/6 mice</td><td>Jackson Laboratories</td><td>000664</td><td></td></tr><tr><td>Micro-Adson Forceps - 1x2 Teeth</td><td>Fine Science Tools (USA) Inc.</td><td>11019-12</td><td></td></tr><tr><td>V-Hook</td><td>Fine Science Tools (USA) Inc.</td><td>18052-12</td><td></td></tr><tr><td>Scalpel SS NO15</td><td>Fischer Scientific</td><td>29556</td><td></td></tr><tr><td>Disposable Needle 30GX1</td><td>Fischer Scientific</td><td>305128</td><td></td></tr><tr><td>Operating Scissors</td><td>Fischer Scientific</td><td>12-460-796</td><td></td></tr><tr><td>Surgi-Or Jeweler&#039;s Forceps, Sklar 4-1/2 in</td><td>Fischer Scientific</td><td>50-118-4255</td><td></td></tr><tr><td>Spring Scissors - Straight/Sharp-Sharp/8mm Cutting Edge</td><td>Fine Science Tools (USA) Inc.</td><td>15024-10</td><td></td></tr><tr><td>Cardiogreen</td><td>Sigma</td><td>I2633-25MG</td><td></td></tr><tr><td>IsosulfanBlue (Lymphazurin)&amp;nbsp; 50 mg/5ml</td><td>Mylan</td><td>67457-220-05</td><td></td></tr></tbody></table>

a murine tail lymphedema model

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.

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 sustained to week 15. Tail volume can be used as an outcome variable to assess the effect of therapeutic interventions for lymphedema in the model. In Figure 3, high resolution laser speckle for assessment of tail vasculature patency can be observed. This adds rigor to the model to ensure welling is secondary to lymphatic dysfunction rather than venous injury. The effect of interventions can then potentially translate to lymphedema treatment with greater confidence. Figure 4 shows a functional lymphatic assessment performed via near infrared laser lymphangiography. This additional outcome variable allows for a functional lymphatic effect of interventions. Figure 5 demonstrates the focal delivery of genetic cargo transcutaneously at the surgical site using tissue nanotransfection technology (TNT2.0). TNT2.0 facilitates point of care delivery of potential candidate gene-based therapeutics in this lymphedema model.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61848/61848fig1v2.jpg" /> 
Figure 1:&#160;Mouse tail model for sustained lymphedema. (A) A 3 mm wide full-thickness skin excision is performed on a murine tail 20 mm from the base under the surgical micrscope. Care is taken to preserve the vasculature. (B) A schematic of the cross-section of the mouse tail. DV=dorsal vein, LV=lateral veins, A=ventral caudal artery, CV=caudal vertebra, T=tendon and muscle, yellow arrows shows the lymphatics. (C) Following administration of isosulfan blue into the tail tip to localize the lymphatics, the lymphatics (yellow arrow) exhibit blue color. The lymphatics are disrupted while preserving the adjacent lateral veins (white arrow). <a href="https://www.jove.com/files/ftp_upload/61848/61848fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61848/61848fig2v2.jpg" /> 
Figure 2:&#160;Progressive swelling of the mouse tail lymphedema model. (A) Following full-thickness skin excision and lymphatic transection, the mouse tail exhibits progressive swelling that is sustained over 15 weeks. The bracket denotes 20 mm from the base of the tail to the start of the surgical full thickness skin excision. (B-C) Quantification of the change in tail volume over 15 weeks represented as (B) bar graphs, each dot representing an animal, n=15, or as (C)&#160;line graph. Data represented as &#177; SEM. <a href="https://www.jove.com/files/ftp_upload/61848/61848fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61848/61848fig3v2.jpg" /> 
Figure 3:&#160;High resolution laser speckle contrast imaging to confirm mouse tail perfusion in the lymphedema mouse tail model. Laser speckle is used to assess mouse tail vasculature postoperatively to validate swelling of lymphatic etiology and minimize tail necrosis. (A) A mouse tail with injured lateral veins (black arrow) detected by laser speckle. (B) Intact lateral tail vein (black arrow) post-lymphedema surgery detected by laser speckle. (n=5) resolution 0.02 mm; Color coded bar indicates perfusion (blue: low, red: high) as measured in arbitrary relative units. <a href="https://www.jove.com/files/ftp_upload/61848/61848fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61848/61848fig4v2.jpg" /> 
Figure 4:&#160;Assessment of lymphatic function using near infrared laser lymphangioraphy in the mouse tail model. Indocyanine green (ICG) injected into the tip of the mouse tail localizes to the lymphatics. Preoperatively, the lymphatics are intact along the mouse tail. Postoperatively, there is no ICG transit beyond the surgical site, confirming that swelling is caused by lymphatic dysfunction. Yellow arrow indicates the base of the tail. <a href="https://www.jove.com/files/ftp_upload/61848/61848fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61848/61848fig5v2.jpg" /> 
Figure 5:&#160;Focal delivery of genetic cargo using tissue nanotransfection technology (TNT). (A) Illustration of TNT delivery. (B) Plasmids are loaded into the TNT2.0 reservoir. The positive and negative electrical probes are attached and a brief, square wave pulse electric stimulation is delivered (10 x 10 ms pulses, 250 V, 10 mA), facilitating focal, non-viral, transcutaneous transfection. (C) Efficiency of genetic cargo delivery using TNT2.0 as observed through fluorescein amidite (FAM) labeled DNA delivery to the murine tail. Mouse tails were sectioned two days after TNT treatment and assessed through fluorescence microscopy. White dotted lines indicate the epithelium of the skin of murine tail. White arrows indicate the FAM labelled DNA.&#160;<a href="https://www.jove.com/files/ftp_upload/61848/61848fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Murine Tail Lymphedema Model at JoVE.com

A Murine Tail Lymphedema Model

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.

Lymphedema is extremity swelling caused by lymphatic dysfunction. The affected limb enlarges because of accumulation of fluid, adipose, and fibrosis. ...

Nanyang Technological University

Research

JoVE Journal

Medicine

5.6K Views.  Indiana University School of Medicine. 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.

Video: A Murine Tail Lymphedema Model

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Immunology and Infection

Pathological Analysis of Lung Metastasis Following Lateral Tail-Vein Injection of Tumor Cells

Metastasis, the primary cause of morbidity and mortality for most cancer patients, can be challenging to model preclinically in mice. Few spontaneous metastasis models are available. Thus, the experimental metastasis model involving tail-vein injection of suitable cell lines is a mainstay of metastasis research. When cancer cells are injected into the lateral tail-vein, the lung is their preferred site of colonization. A potential limitation of this technique is the accurate quantification of the metastatic lung tumor burden. While some investigators count macrometastases of a pre-defined size and/or include micrometastases following sectioning of tissue, others determine the area of metastatic lesions relative to normal tissue area. Both of these quantification methods can be exceedingly difficult when the metastatic burden is high. Herein, we demonstrate an intravenous injection model of lung metastasis followed by an advanced method for quantifying metastatic tumor burden using image analysis software. This process allows for investigation of multiple end-point parameters, including average metastasis size, total number of metastases, and total metastasis area, to provide a comprehensive analysis. Furthermore, this method has been reviewed by a veterinary pathologist board-certified by the American College of Veterinary Pathologists (SEK) to ensure accuracy.

Intravenous injection of cancer cells is often used in metastasis research, but the metastatic tumor burden can be difficult to analyze. Herein, we demonstrate a tail-vein injection model of metastasis and include a novel approach to analyze the resulting metastatic lung tumor burden.

Metastasis, the primary cause of morbidity and mortality for most cancer patients, can be challenging to model preclinically in mice. Few spontaneous ...

Cancer Research

Analysis of Lymph Node Volume by Ultra-High-Frequency Ultrasound Imaging in the Braf/Pten Genetically Engineered Mouse Model of Melanoma

Tyr::CreER+,BrafCA/+,Ptenlox/lox&#160;genetically engineered mice (Braf/Pten mice) are widely used as an in vivo model of metastatic melanoma. Once a primary tumor has been induced by tamoxifen treatment, an increase in metastatic burden is observed within 4-6 weeks after induction. This paper shows how Ultra-High-Frequency UltraSound (UHFUS) imaging can be exploited to monitor the increase in metastatic involvement of the inguinal lymph nodes by measuring the increase in their volume.
The UHFUS system is used to scan anesthetized mice with a UHFUS linear probe (22-55 MHz, axial resolution 40 &#181;m). B-mode images from the inguinal lymph nodes (both left and right sides) are acquired in a short-axis view, positioning the animals in dorsal recumbency. Ultrasound records are acquired using a 44 &#181;m step size on a motorized mechanical arm. Afterward, two-dimensional (2D) B-mode acquisitions are imported into the software platform for ultrasound image post-processing, and inguinal lymph nodes are identified and segmented semi-automatically in the acquired cross-sectional 2D images. Finally, a total reconstruction of the three-dimensional (3D) volume is automatically obtained along with the rendering of the lymph node volume, which is also expressed as an absolute measurement.
This non-invasive in vivo technique is very well tolerated and allows the scheduling of multiple imaging sessions on the same experimental animal over 2 weeks. It is, therefore, ideal to assess the impact of pharmacological treatment on metastatic disease.

Melanoma is a very aggressive disease that quickly spreads to other organs. This protocol describes the application of ultra-high-frequency ultrasound imaging, coupled with 3D rendering, to monitor the volume of the inguinal lymph nodes in the Braf/Pten mouse model of metastatic melanoma.

Tyr::CreER+,BrafCA/+,Ptenlox/lox&#160;genetically engineered mice (Braf/Pten mice) are widely used as an in vivo model of metastatic melanoma. Once a ...

Assessment of Acute Wound Healing using the Dorsal Subcutaneous Polyvinyl Alcohol Sponge Implantation and Excisional Tail Skin Wound Models.

Wound healing is a complex process that requires the orderly progression of inflammation, granulation tissue formation, fibrosis, and resolution. Murine models provide valuable mechanistic insight into these processes; however, no single model fully addresses all aspects of the wound healing response. Instead, it is ideal to use multiple models to address the different aspects of wound healing. Here, two different methods that address diverse aspects of the wound healing response are described. In the first model, polyvinyl alcohol sponges are subcutaneously implanted along the mouse dorsum. Following sponge retrieval, cells can be isolated by mechanical disruption, and fluids can be extracted by centrifugation, thus allowing for a detailed characterization of cellular and cytokine responses in the acute wound environment. A limitation of this model is the inability to assess the rate of wound closure. For this, a tail skin excision model is utilized. In this model, a 10 mm x 3 mm rectangular piece of tail skin is excised along the dorsal surface, near the base of the tail. This model can be easily photographed for planimetric analysis to determine healing rates and can be excised for histological analysis. Both described methods can be utilized in genetically altered mouse strains, or in conjunction with models of comorbid conditions, such as diabetes, aging, or secondary infection, in order to elucidate wound healing mechanisms.

Here, two murine wound healing models are described, one designed to assess cellular and cytokine wound healing responses and the other to quantify the rate of wound closure. These methods can be used with complex disease models such as diabetes to determine mechanisms of various aspects of poor wound healing.

Wound healing is a complex process that requires the orderly progression of inflammation, granulation tissue formation, fibrosis, and resolution. Murine ...

Blocking Lymph Flow by Suturing Afferent Lymphatic Vessels in Mice

Lymphatic vessels are critical in maintaining tissue fluid balance and optimizing immune protection by transporting antigens, cytokines, and cells to draining lymph nodes (LNs). Interruption of lymph flow is an important method when studying the function of lymphatic vessels. The afferent lymphatic vessels from the murine footpad to the popliteal lymph nodes (pLNs) are well-defined as the only routes for lymph drainage into the pLNs. Suturing these afferent lymphatic vessels can selectively prevent lymph flow to the pLNs. This method allows for interference in lymph flow with minimal damage to the lymphatic endothelial cells in the draining pLN, the afferent lymphatic vessels, as well as other lymphatic vessels around the area. This method has been used to study how lymph impacts high endothelial venules (HEV) and chemokine expression in the LN, and how lymph flows through the adipose tissue surrounding the LN in the absence of functional lymphatic vessels. With the growing recognition of the importance of lymphatic function, this method will have broader applications to further unravel the function of lymphatic vessels in regulating the LN microenvironment and immune responses.

A protocol to block lymph flow by surgical suturing of afferent lymphatic vessels is presented.

Lymphatic vessels are critical in maintaining tissue fluid balance and optimizing immune protection by transporting antigens, cytokines, and cells to ...

A Revised Method for Inducing Secondary Lymphedema in the Hindlimb of Mice

Animal models are of paramount importance in the research of lymphedema in order to understand the pathophysiology of the disease but also to explore potential treatment options. This mouse model allows researchers to induce significant lymphedema lasting at least 8 weeks. Lymphedema is induced using a combination of fractioned radiotherapy and surgical ablation of lymphatics. This model requires that the mice get a dose of 10 Gray (Gy) radiation before and after surgery. The surgical part of the model involves ligation of three lymph vessels and extraction of two lymph nodes from the mouse hindlimb. Having access to microsurgical tools and a microscope is essential, due to the small anatomical structures of mice. The advantage of this model is that it results in statistically significant lymphedema, which provides a good basis for evaluating different treatment options. It is also a great and easily available option for microsurgical training. The limitation of this model is that the procedure can be time consuming, especially if not practiced in advance. The model results in objectively quantifiable lymphedema in mice, without causing severe morbidity and has been tested in three separate projects.

This animal model enables researchers to induce statistically significant secondary lymphedema in the hindlimb of mice, lasting at least 8 weeks. The model can be used to study the pathophysiology of lymphedema and to investigate novel treatment options.

Animal models are of paramount importance in the research of lymphedema in order to understand the pathophysiology of the disease but also to explore ...

Evaluating Antigen-Specific CD8+ T Cells During Lymph Node Metastasis

Draining Lymph Node Metastasis Model for Assessing the Dynamics of Antigen-Specific CD8+ T Cells During Tumorigenesis

Tumor antigen-specific CD8+ T cells from draining lymph nodes gain an accumulating importance in mounting anti-tumor immune response during tumorigenesis. However, in many cases, cancer cells form metastatic loci in lymph nodes before further metastasizing to distant organs. To what extent the local and systematic CD8+ T cell responses were influenced by LN metastasis remains obscure. To this end, we set up a murine LN metastasis model combined with a B16F10-GP melanoma cell line expressing the surrogate neoantigen derived from lymphocytic choriomeningitis virus (LCMV), glycoprotein (GP), and P14 transgenic mice harboring T cell receptors (TCRs) specific to GP-derived peptide GP33-41 presented by the class I major histocompatibility complex (MHC) molecule H-2Db. This protocol enables the study of antigen-specific CD8+ T cell responses during LN metastasis. In this protocol, C57BL/6J mice were subcutaneously implanted with B16F10-GP cells, followed by adoptive transfer with naive P14 cells. When the subcutaneous tumor grew to approximately 5 mm in diameter, the primary tumor was excised, and B16F10-GP cells were directly injected into the tumor draining lymph node (TdLN). Then, the dynamics of CD8+ T cells were monitored during the process of LN metastasis. Collectively, this model has provided an approach to precisely investigate the antigen-specific CD8+ T cell immune responses during LN metastasis.

Author Spotlight: A Model to Study the Systemic and Local Dynamics of CD8+ T Cells During LN Metastasis

The experimental design presented here provides a useful reproductive model for the studies of antigen-specific CD8+ T cells during lymph node (LN) metastasis, which excludes the perturbation of bystander CD8+ T cells.

Tumor antigen-specific CD8+ T cells from draining lymph nodes gain an accumulating importance in mounting anti-tumor immune response during tumorigenesis. ...

Popliteal Vascular Lymph Node Resection in the Rabbit Hindlimb for Secondary Lymphedema Induction

Lymphedema is a common condition often associated with cancer and its treatment, which leads to damage to the lymphatic system, and current treatments are mostly palliative rather than curative. Its high incidence among oncologic patients indicates the need to study both normal lymphatic function and pathologic dysfunction. To reproduce chronic lymphedema, it is necessary to choose a suitable experimental animal. Attempts to establish animal models are limited by the regenerative capacity of the lymphatic system. Among the potential candidates, the rabbit hindlimb is easy to handle and extrapolate to the human clinical scenario, making it advantageous. In addition, the size of this species allows for better selection of lymphatic vessels for vascularized lymph node resection.
In this study, we present a procedure of vascular lymph node resection in the rabbit hindlimb for inducing secondary lymphedema. Anesthetized animals were subjected to circumferential measurement, patent blue V infiltration, and indocyanine green lymphography (ICG-L) using real-time near-infrared fluorescence, a technique that allows the identification of single popliteal nodes and lymphatic channels. Access to the identified structures is achieved by excising the popliteal node and ligating the medial and lateral afferent lymphatics. Special care must be taken to ensure that any lymphatic vessel that joins the femoral lymphatic system within the thigh without entering the popliteal node can be identified and ligated. 
Postoperative evaluation was performed at 3, 6, and 12 months after induction using circumferential measurements of the hindlimb and ICG-L. As demonstrated during follow-up, the animals developed dermal backflow that was maintained until the 12th month, making this experimental animal useful for novel long-term evaluations in the management of lymphedema. In conclusion, the approach described here is feasible and reproducible. Additionally, during the time window presented, it can be representative of human lymphedema, thus providing a useful research tool.

Here, the surgical induction of stable acquired lymphedema in the rabbit hindlimb is described. This experimental animal can be used to further investigate the effect of lymphedema treatment by microsurgical techniques.

Lymphedema is a common condition often associated with cancer and its treatment, which leads to damage to the lymphatic system, and current treatments are ...

Experimental Metastatic Melanoma Mouse Model: A Platform to Study the Metastatic Potential of Melanoma Cells in the Lungs

Source: Joshua J. Timmons&#160;et al. <a href="https://www.jove.com/v/54039"> Injection of Syngeneic Murine Melanoma Cells to Determine Their Metastatic Potential in the Lungs</a>. J. Vis. Exp. (2016)
In this video, melanoma cells are intravenously injected through the tail vein in the mouse model. The melanoma cells further travel through the circulatory system, adapting to the distant microenvironment and causing metastasis in organs like lungs.

Source: Joshua J. Timmons&#160;et al.  Injection of Syngeneic Murine Melanoma Cells to Determine Their Metastatic Potential in the Lungs. J. Vis. Exp. ...

JoVE Encyclopedia of Experiments

Skin Cancer

Animal model studies

Harvesting an Inguinal Draining Lymph Node from a Murine Model

Source: Rezende, R. M. et al., <a href="https://app.jove.com/v/59335">Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy.</a>&#160;J. Vis. Exp.&#160; (2019)
This video demonstrates the surgical isolation procedure of the inguinal draining lymph node from a mouse model. Incision of the abdominal region provides access to the inguinal lymph node, which is surgically harvested. The harvested translucent inguinal lymph node can be used for immunological studies.

Source: Rezende, R. M. et al., Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy.&#160;J. Vis. Exp.&#160; ...