The authors thank Peter Bollen, head of the Biomedical Laboratory for lending the equipment needed to record the footage seen through the microscopes.

Animals were housed in the University of Southern Denmark Animal Care Facility as per institutional guidelines. All procedures involving animal subjects have been approved by The Animal Experiments Inspectorate, Ministry of Environment and Food of Denmark.
1. Pre-surgery irradiation
NOTE: Pre-surgery irradiation takes place 7 days before surgery.
<ol>
	<li>Induce anesthesia.
	<ol>
		<li>Place the mouse in an induction box and set the vaporizer to 3% isoflurane with an oxygen flow rate of 0.8&#8722;1.2 L/min to induce inhalation anesthesia. 
		NOTE: Alternatively, injectable anesthetics can be used but for the short duration of the irradiation inducing inhalation anesthesia was sufficient. For obtaining the results presented in this article, 9-week old female C57BL6 mice were used.</li>
		<li>Make sure the mouse is fully anesthetized by tail or paw pinch test.</li>
	</ol>
	</li>
	<li>Position the mouse for irradiation.
	<ol>
		<li>If fully sedated, move the mouse from the induction box and place it under the source of radiation in supine position and gently fixate the hind limbs with tape. 
		NOTE: The mouse will remain sedated for the short duration of the radiation.</li>
		<li>Place a 1.5 mm thick lead pad to ensure that only the area that undergoes surgery (i.e., the circular area with a diameter of 25 mm around the knee) gets irradiated.</li>
	</ol>
	</li>
	<li>Administer a dose of 10 Gy radiation at a dose rate of 5.11 Gy/min (100 kVp, 10 mA). 
	CAUTION: Safety precautions must be taken when working with radiation. During this experiment, all irradiation was performed in a radiation insulated room, and the source of radiation was only turned on when all personnel had left and sealed the room.</li>
	<li>Place the mouse back in its cage.</li>
</ol>
2. Equipment setup
NOTE: Surgery should be performed in a room dedicated to surgical procedures. The operative surface must be sterile.
<ol>
	<li>Thoroughly clean all operative surfaces with 70% ethanol. Wear hairnet and coveralls. Use sterile surgical instruments and sterile gloves.</li>
	<li>Prepare anesthesia.
	<ol>
		<li>Draw up 1 mL of fentanyl (0.315 mg/mL), 1 mL of midazolam (5 mg/mL), and 2 mL of sterile water. Use different syringes and needles for the different components.</li>
		<li>Mix fentanyl and sterile water by slowly emptying the syringes into a sterile glass tube. When mixed, add midazolam to complete the working solution.</li>
	</ol>
	</li>
	<li>Prepare analgesia.
	<ol>
		<li>Draw up 0.2 mL of buprenorphine (0.3 mg/mL) and 2 mL of saline.</li>
		<li>Mix the volumes by slowly emptying the syringes into a sterile glass tube to complete the working solution.</li>
	</ol>
	</li>
	<li>Turn on the microscope and make sure that the lighting is sufficient, and that the microscope is well adjusted for the operator&#39;s eyes. 
	NOTE: All surgical procedures should be performed under an operating microscope. A magnification range from 4x&#8722;25x is sufficient.</li>
</ol>
3. Preparation
<ol>
	<li>Weigh the mouse pre-surgery by placing the mouse in an empty container on a cleared scale.</li>
	<li>Administer anesthetic.
	<ol>
		<li>Draw up 0.1 mL of anesthetic per 10 g of mouse bodyweight. Inject the anesthetic subcutaneously as a bolus injection.</li>
		<li>Let the mouse rest in a cage with plenty bedding and shelter for approximately 10 min until fully sedated. Examine the anesthetic depth by assessing muscle relaxation and perform paw or tail pinch test.</li>
	</ol>
	</li>
	<li>When fully sedated, shave the hind limb chosen for the procedure using electrical clippers. Make sure to wipe of excess hair.</li>
	<li>Turn on the heating device, such as a heating pad and cover it with a surgical cloth.</li>
	<li>Set the flow of oxygen to 0.8 L/min and connect it with a nosecone. Use 100% oxygen. 
	NOTE: The nosecone is only for oxygen delivery and not anesthesia.</li>
	<li>Apply ophthalmic ointment and inject 0.5 mL of saline subcutaneously, preferably in the scruff of the mouse, to prevent hypovolemia during surgery.</li>
	<li>Position the mouse for surgery.
	<ol>
		<li>Place the mouse on the surgical cloth in supine position. Place the nosecone over the snout.</li>
		<li>Fixate the end of the hindlimbs gently with tape to prevent the mouse from shifting during surgery.</li>
		<li>Sterilize the skin using alcohol/chlorhexidine or alcohol/povidone iodine.</li>
	</ol>
	</li>
</ol>
4. Surgery
NOTE: In this example, the left hind limb (when the mouse is viewed in supine position), has been chosen for the procedure.
<ol>
	<li>Make a circular incision.
	<ol>
		<li>Lift the skin with smooth forceps and clip a small opening approximately 5 mm proximal to the popliteal fossa.</li>
		<li>Slide sharp scissors into the opening and clip towards the knee so that the incision ends just above the knee. Make sure not to puncture the underlying vessels by lifting the skin with forceps while clipping.</li>
		<li>Move the mouse to prone position and continue to clip from the knee towards the popliteal fossa until the circumferential incision is complete.</li>
	</ol>
	</li>
	<li>Dissect the skin below the knee.
	<ol>
		<li>Gently blunt dissect the area below the knee to a couple of millimeters above the ankle, by slowly opening and closing the microscissors while lifting the skin with forceps.</li>
		<li>Carefully snip remaining visible adhesions using microscissors. Use sterile saline regularly to keep the tissue moist during the whole procedure.</li>
	</ol>
	</li>
	<li>Dissect the skin at the proximal rim of the circumferential incision so that it can be retracted with an elastic retractor. 
	NOTE: The retractor allows the operator a better view of the proximal lymph vessel and prevents the proximal rim from shifting during surgery.</li>
	<li>While still in prone position, rotate the hindlimb gently and fixate it with tape, so that the ischiatic vein is visible from the most proximal point of the exposed area to the most distal point.</li>
	<li>Inject approximately 0.01 mL of Patent Blue V subcutaneously between the second and third toe using a 0.5 mL syringe with a 30 G needle. Gently press the paw a couple of times to distribute the Patent Blue V. Visualize the lymph vessels and lymph node through the microscope as the Patent Blue V fills the lymph vessels. 
	NOTE: If the blue color of the lymph vessels fades during the procedure, gently massage the paw to promote uptake, rather than inject more Patent Blue V. Excess use of Patent Blue V may lead to leakage and coloring of the tissue surrounding the lymph vessels which may compromise the procedure.</li>
	<li>Locate the important structures: the popliteal lymph node (PLN), the two lymph vessels distal to the lymph node (DLV1 and DLV2), and the one lymph vessel proximal to the lymph node (PLV). 
	NOTE: All the lymph vessels can be found adjacent to the ischiatic vein. The proximal lymph vessel is usually found medial to the vein, the two distal lymph vessels are found medial and lateral to the vein. The abbreviations of the structures are used in the accompanying video.</li>
	<li>Magnify to clearly visualize the PLV and ligate it with a 10-0 nylon suture using micro-needle holder and microforceps. Press the paw a couple of times to ensure that no Patent Blue V passes proximal to the suture. 
	NOTE: Trimming the fat surrounding the lymph vessel may be necessary.</li>
	<li>Repeat step 4.7 to ligate the two distal lymph vessels. Press the paw several times to ensure that no Patent Blue V passes proximal to the ligature. If the lymph vessels lie to close to the ischiatic vein, try dissecting even further distally. 
	NOTE: In this example, it can be seen that one of the lymph vessels bursts due to the ligature hindering the lymph flow. The lymph vessels will often split from the vein further down.</li>
	<li>Remove the popliteal lymph node.
	<ol>
		<li>Locate the popliteal lymph node and clip a small hole with microscissors to access it and remove it with microforceps and microscissors. 
		NOTE: The lymph node has a smooth pearl-like surface in contrast to the surrounding fat tissue.</li>
		<li>To test if the removed tissue is a lymph node, place it in a test tube filled with water. 
		NOTE: If the tissue is comprised of fat, the tissue will float. If the tissue is a lymph node, it will sink to the bottom.</li>
	</ol>
	</li>
	<li>Remove the inguinal fat pad and lymph node.
	<ol>
		<li>Before removing the inguinal fat pad, use a bipolar coagulator to cauterize the vessels running through the fat.</li>
		<li>Resect the inguinal fat pad using microforceps and microscissors. Gently clip the cauterized vessels running through the fat. Then gently resect the fat tissue in the inguinal area. 
		NOTE: The lymph node located in the fat is rarely colored by Patent Blue V and can be hard to differentiate from the fat. Removing the fat pad in one piece is the best way to ensure the lymph node has been removed.</li>
	</ol>
	</li>
	<li>Rinse the leg thoroughly with sterile saline and confirm through the microscope that any small hairs and particles has been thoroughly removed from the surgical area to avoid wound contamination and infection. Make sure there is no active bleeding.</li>
	<li>Suture the skin edges down to the muscle facia with a 6-0 nylon suture using forceps and needle holder, leaving a gap of 2&#8722;3 mm to constrain the superficial lymph flow. 
	NOTE: The accompanying video shows an example of finished sutures.</li>
	<li>Administer analgesia. Draw up 0.1 mL of analgesia per 30 g of mouse bodyweight. Inject the analgesia subcutaneously as a bolus injection.</li>
	<li>Weigh the mouse for post-surgery for comparison.</li>
	<li>Place the mouse in a cage in a cabinet heated for recovery.</li>
</ol>
5. Postoperative care
<ol>
	<li>Give the mice individual cages to recover after surgery with water and food ad libitum.</li>
	<li>Administer a bolus subcutaneous dose of 0.02 mL of buprenorphine 3x daily for 3 days for analgesia.</li>
	<li>Monitor the animal daily for appropriate wound healing, signs of pain and infection. If signs of infection are present, use antibiotic ointment.</li>
</ol>
6. Post-surgery irradiation
<ol>
	<li>Three days after surgery, repeat the procedure for pre-surgery irradiation (steps 1.1&#8722;1.4).</li>
</ol>

<ol><li>Lawenda, B. D., Mondry, T. E., Johnstone, P. A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymphedema%3A+a+primer+on+the+identification+and+management+of+a+chronic+condition+in+oncologic+treatment%5BTitle%5D)+AND+%22CA%3A+A+Cancer+Journal+for+Clinicians%22%5BJournal%5D)">Lymphedema: a primer on the identification and management of a chronic condition in oncologic treatment.</a> CA: A Cancer Journal for Clinicians. 59 (1), 8-24 (2009).</li>
<li>Greene, A. K. Epidemiology and morbidity of lymphedema. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aGreene%2C+AK+%22Lymphedema%3A+Presentation%2C+Diagnosis%2C+and+Treatment%22">Lymphedema: Presentation, Diagnosis, and Treatment</a>. Greene, A. K., Slavin, S. A., Brorson, H. , Springer International Publishing. New York, NY. 33-44 (2015).</li>
<li>Hespe, G. E., Nores, G. G., Huang, J. J., Mehrara, B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Pathophysiology+of+lymphedema-Is+there+a+chance+for+medication+treatment%3F%5BTitle%5D)+AND+%22Journal+of+Surgical+Oncology%22%5BJournal%5D)">Pathophysiology of lymphedema-Is there a chance for medication treatment?</a> Journal of Surgical Oncology. 115 (1), 96-98 (2017).</li>
<li>Grada, A. A., Phillips, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymphedema%3A+Pathophysiology+and+clinical+manifestations%5BTitle%5D)+AND+%22Journal+of+the+American+Academy+of+Dermatology%22%5BJournal%5D)">Lymphedema: Pathophysiology and clinical manifestations.</a> Journal of the American Academy of Dermatology. 77 (6), 1009-1020 (2017).</li>
<li>Chang, D. W., Masia, J., Garza, R. 3rd, Skoracki, R., Neligan, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymphedema%3A+Surgical+and+Medical+Therapy%5BTitle%5D)+AND+%22Plastic+and+Reconstructive+Surgery%22%5BJournal%5D)">Lymphedema: Surgical and Medical Therapy.</a> Plastic and Reconstructive Surgery. 138 (3 Suppl), 209S-218S (2016).</li>
<li>Carl, H. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Systematic+Review+of+the+Surgical+Treatment+of+Extremity+Lymphedema%5BTitle%5D)+AND+%22Journal+of+Reconstructive+Microsurgery%22%5BJournal%5D)">Systematic Review of the Surgical Treatment of Extremity Lymphedema.</a> Journal of Reconstructive Microsurgery. 33 (6), 412-425 (2017).</li>
<li>DiSipio, T., Rye, S., Newman, B., Hayes, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Incidence+of+unilateral+arm+lymphoedema+after+breast+cancer%3A+a+systematic+review+and+meta-analysis%5BTitle%5D)+AND+%22The+Lancet+Oncology%22%5BJournal%5D)">Incidence of unilateral arm lymphoedema after breast cancer: a systematic review and meta-analysis.</a> The Lancet Oncology. 14 (6), 500-515 (2013).</li>
<li>Douglass, J., Graves, P., Gordon, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Self-Care+for+Management+of+Secondary+Lymphedema%3A+A+Systematic+Review%5BTitle%5D)+AND+%22PLoS+Neglected+Tropical+Diseases%22%5BJournal%5D)">Self-Care for Management of Secondary Lymphedema: A Systematic Review.</a> PLoS Neglected Tropical Diseases. 10 (6), e0004740(2016).</li>
<li>Shih, Y. C. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Incidence%2C+treatment+costs%2C+and+complications+of+lymphedema+after+breast+cancer+among+women+of+working+age%3A+a+2-year+follow-up+study%5BTitle%5D)+AND+%22Journal+of+Clinical+Oncology%22%5BJournal%5D)">Incidence, treatment costs, and complications of lymphedema after breast cancer among women of working age: a 2-year follow-up study.</a> Journal of Clinical Oncology. 27 (12), 2007-2014 (2009).</li>
<li>Ridner, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+psycho-social+impact+of+lymphedema%5BTitle%5D)+AND+%22Lymphatic+Research+and+Biology%22%5BJournal%5D)">The psycho-social impact of lymphedema.</a> Lymphatic Research and Biology. 7 (2), 109-112 (2009).</li>
<li>Gutknecht, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cost-of-illness+of+patients+with+lymphoedema%5BTitle%5D)+AND+%22Journal+of+the+European+Academy+of+Dermatology+and+Venereology%22%5BJournal%5D)">Cost-of-illness of patients with lymphoedema.</a> Journal of the European Academy of Dermatology and Venereology. 31 (11), 1930-1935 (2017).</li>
<li>Hayes, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Prevalence+and+prognostic+significance+of+secondary+lymphedema+following+breast+cancer%5BTitle%5D)+AND+%22Lymphatic+Research+and+Biology%22%5BJournal%5D)">Prevalence and prognostic significance of secondary lymphedema following breast cancer.</a> Lymphatic Research and Biology. 9 (3), 135-141 (2011).</li>
<li>Danese, C. A., Georgalas-Bertakis, M., Morales, L. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+model+of+chronic+postsurgical+lymphedema+in+dogs%27+limbs%5BTitle%5D)+AND+%22Surgery%22%5BJournal%5D)">A model of chronic postsurgical lymphedema in dogs' limbs.</a> Surgery. 64 (4), 814-820 (1968).</li>
<li>Das, S. K., Franklin, J. D., O'Brien, B. M., Morrison, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+practical+model+of+secondary+lymphedema+in+dogs%5BTitle%5D)+AND+%22Plastic+and+Reconstructive+Surgery%22%5BJournal%5D)">A practical model of secondary lymphedema in dogs.</a> Plastic and Reconstructive Surgery. 68 (3), 422-428 (1981).</li>
<li>Huang, G. K., Hsin, Y. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+experimental+model+for+lymphedema+in+rabbit+ear%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">An experimental model for lymphedema in rabbit ear.</a> Microsurgery. 4 (4), 236-242 (1983).</li>
<li>Tobbia, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymphedema+development+and+lymphatic+function+following+lymph+node+excision+in+sheep%5BTitle%5D)+AND+%22Journal+of+Vascular+Research%22%5BJournal%5D)">Lymphedema development and lymphatic function following lymph node excision in sheep.</a> Journal of Vascular Research. 46 (5), 426-434 (2009).</li>
<li>Lahteenvuo, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Growth+factor+therapy+and+autologous+lymph+node+transfer+in+lymphedema%5BTitle%5D)+AND+%22Circulation%22%5BJournal%5D)">Growth factor therapy and autologous lymph node transfer in lymphedema.</a> Circulation. 123 (6), 613-620 (2011).</li>
<li>Honkonen, K. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymph+node+transfer+and+perinodal+lymphatic+growth+factor+treatment+for+lymphedema%5BTitle%5D)+AND+%22Annals+of+Surgery%22%5BJournal%5D)">Lymph node transfer and perinodal lymphatic growth factor treatment for lymphedema.</a> Annals of Surgery. 257 (5), 961-967 (2013).</li>
<li>Wang, G. Y., Zhong, S. Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+model+of+experimental+lymphedema+in+rats%27+limbs%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">A model of experimental lymphedema in rats' limbs.</a> Microsurgery. 6 (4), 204-210 (1985).</li>
<li>Oashi, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+new+model+of+acquired+lymphedema+in+the+mouse+hind+limb%3A+a+preliminary+report%5BTitle%5D)+AND+%22Annals+of+Plastic+Surgery%22%5BJournal%5D)">A new model of acquired lymphedema in the mouse hind limb: a preliminary report.</a> Annals of Plastic Surgery. 69 (5), 565-568 (2012).</li>
<li>Slavin, S. A., Van den Abbeele, A. D., Losken, A., Swartz, M. A., Jain, R. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Return+of+lymphatic+function+after+flap+transfer+for+acute+lymphedema%5BTitle%5D)+AND+%22Annals+of+Surgery%22%5BJournal%5D)">Return of lymphatic function after flap transfer for acute lymphedema.</a> Annals of Surgery. 229 (3), 421-427 (1999).</li>
<li>Cheung, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+experimental+model+for+the+study+of+lymphedema+and+its+response+to+therapeutic+lymphangiogenesis%5BTitle%5D)+AND+%22BioDrugs+%3A+Clinical+Immunotherapeutics%2C+Biopharmaceuticals+and+Gene+Therapy%22%5BJournal%5D)">An experimental model for the study of lymphedema and its response to therapeutic lymphangiogenesis.</a> BioDrugs : Clinical Immunotherapeutics, Biopharmaceuticals and Gene Therapy. 20 (6), 363-370 (2006).</li>
<li>Rutkowski, J. M., Moya, M., Johannes, J., Goldman, J., Swartz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Secondary+lymphedema+in+the+mouse+tail%3A+Lymphatic+hyperplasia%2C+VEGF-C+upregulation%2C+and+the+protective+role+of+MMP-9%5BTitle%5D)+AND+%22Microvascular+Research%22%5BJournal%5D)">Secondary lymphedema in the mouse tail: Lymphatic hyperplasia, VEGF-C upregulation, and the protective role of MMP-9.</a> Microvascular Research. 72 (3), 161-171 (2006).</li>
<li>Tammela, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Therapeutic+differentiation+and+maturation+of+lymphatic+vessels+after+lymph+node+dissection+and+transplantation%5BTitle%5D)+AND+%22Nature+Medicine%22%5BJournal%5D)">Therapeutic differentiation and maturation of lymphatic vessels after lymph node dissection and transplantation.</a> Nature Medicine. 13 (12), 1458-1466 (2007).</li>
<li>Frueh, F. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+models+in+surgical+lymphedema+research--a+systematic+review%5BTitle%5D)+AND+%22Journal+of+Surgical+Research%22%5BJournal%5D)">Animal models in surgical lymphedema research--a systematic review.</a> Journal of Surgical Research. 200 (1), 208-220 (2016).</li>
<li>Jorgensen, M. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Quantification+of+Chronic+Lymphedema+in+a+Revised+Mouse+Model%5BTitle%5D)+AND+%22Annals+of+Plastic+Surgery%22%5BJournal%5D)">Quantification of Chronic Lymphedema in a Revised Mouse Model.</a> Annals of Plastic Surgery. 81 (5), 594-603 (2018).</li>
<li>Frueh, F. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((High-resolution+3D+volumetry+versus+conventional+measuring+techniques+for+the+assessment+of+experimental+lymphedema+in+the+mouse+hindlimb%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">High-resolution 3D volumetry versus conventional measuring techniques for the assessment of experimental lymphedema in the mouse hindlimb.</a> Scientific Reports. 6, 34673(2016).</li>
<li>Biau, D. J., Kerneis, S., Porcher, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Statistics+in+brief%3A+the+importance+of+sample+size+in+the+planning+and+interpretation+of+medical+research%5BTitle%5D)+AND+%22Clinical+Orthopaedics+and+Related+Research%22%5BJournal%5D)">Statistics in brief: the importance of sample size in the planning and interpretation of medical research.</a> Clinical Orthopaedics and Related Research. 466 (9), 2282-2288 (2008).</li>
<li>Korula, P., Varma, S. K., Sunderrao, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Inhibition+of+wound+contraction+by+point-to-point+adherent+splintage%5BTitle%5D)+AND+%22Plastic+and+Reconstructive+Surgery%22%5BJournal%5D)">Inhibition of wound contraction by point-to-point adherent splintage.</a> Plastic and Reconstructive Surgery. 95 (4), 725-730 (1995).</li>
<li>Komatsu, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lymph+Drainage+During+Wound+Healing+in+a+Hindlimb+Lymphedema+Mouse+Model%5BTitle%5D)+AND+%22Lymphatic+Research+and+Biology%22%5BJournal%5D)">Lymph Drainage During Wound Healing in a Hindlimb Lymphedema Mouse Model.</a> Lymphatic Research and Biology. 15 (1), 32-38 (2017).</li>
</ol>

The authors have nothing to disclose.

There are a few critical steps in this protocol. Firstly, it is important that the researchers take safety precautions when working with radioactivity. Secondly, during the surgical part of this protocol, it is important to start the procedure once the mouse has been anesthetized and finish it without unnecessary breaks. This is important to avoid an excessively long surgical period for the animal and to prevent that the anesthesia loses effect during surgery. It is recommended to only administer one bolus injection of a...

Lymphedema is characterized by an accumulation of lymph fluid that leads to localized tissue swelling, which mainly occurs due to impaired or disrupted flow of lymph fluid in the lymphatic vessels1. The lymph flow can be impaired or disrupted by infection, obstruction, injury or congenital defects in the lymphatic system2. These etiologies result in accumulation of lymphatic fluid, which leads to a chronic state of inflammation, resulting in subsequent fibrosis, as well as deposition of adipose tissue3. Lymphedema can be categorized as primary or secondary lymphedema. Primary lymphedema is caused by developmental abnormalities or genetic mutation2,4. Secondary lymphedema occurs due to underlying systemic disease, surgery or trauma2,4. Secondary lymphedema is the most common form of lymphedema in the world2. In developed countries, the most common cause of secondary lymphedema is oncological therapy such as adjuvant radiotherapy and lymph node dissection5. Lymphedema is most frequent among breast cancer patients, but can also develop in patients with gynecologic, melanoma, genitourinary or neck cancer6. It has been suggested that out of all women diagnosed with breast cancer, 21% will develop lymphedema7.
Lymphedema can be stressful to the patient both physically and psychologically. Patients with lymphedema have an increased risk of infection5,8,9, poor quality of life and can develop social anxiety and symptoms of depression10. The complications of chronic lymphedema lead to high cost of care and an increased disease burden9,11. Findings have also suggested that lymphedema might be associated with increased risk of death after breast cancer treatment12. Conservative management such as compression of the affected area, manual lymph drainage and general skincare remain the first line approach. There is currently no curative treatment6. Although progress has been made in the field of surgical and medical therapy, there is still room for improvement. More research, providing insight in the pathophysiology and progression of the disease, is needed to enable clinicians to provide better treatment options for the patients5.
Animal models are being used in preclinical research to understand the pathophysiology of diseases and develop potential treatment options. Several different lymphedema animal models have been established in canines13,14, rabbits15, sheep16, pigs17,18 and rodents19,20,21,22,23,24. The rodent model seems to be the most cost-effective model, when investigating the reconstruction of lymphatic function, due to rodents being easily accessible and relatively low-priced25. The majority of the mice models have focused on inducing lymphedema in the tail of the mice21,22,23. The tail model is very reliable but the exact surgical technique for inducing lymphedema varies significantly in previous published material. This results in fluctuations in duration and robustness of the developed lymphedema presented in known litterature25. Different techniques are also being used for inducing lymphedema in the hindlimb model and they also yield varying results, but the hindlimb model might be easier to understand from a translational perspective. Previous lymphedema models have been hampered by spontaneous lymphedema resolution and therefore a reproducible and permanent lymphedema model is needed25. Researchers have previously tried to increase the dose of radiation, to prevent the spontaneous lymphedema resolution, but this has often led to subsequent severe morbidity25.
This model results in statistically significant lymphedema, without causing severe morbidity, by combining microsurgery with radiation. The model has been revised from a previous surgical model by adding a dose of irradiation that induces lymphedema, without causing severe morbidity26. It also offers a great opportunity for microsurgical training. Having access to microsurgical equipment and a microscope is necessary, due to the small anatomical structures of the mice. The surgical procedure can be performed when the user has been taught basic microsurgical techniques, such as suturing with microsurgical instruments. The operators that performed this procedure all watched tutorial videos by Acland on the preconditions of microsurgical skills (1981) and basic microsuture technique (1985). We recommend practicing the surgical procedure 8&#8722;10 times before using it in research. Practicing the procedure ensures that fewer mistakes are made and that the procedure can be performed more efficiently. When mastered, the surgical procedure can be performed in 45 minutes.

<table><tbody><tr><td>10-0 Nylon suture</td><td>S&amp;amp;T</td><td>12051-10</td><td></td></tr><tr><td>6-0 Nylon suture - Dafilon</td><td>B Braun</td><td>C0933112</td><td></td></tr><tr><td>Coagulator - ICC 50</td><td>ERBE</td><td></td><td></td></tr><tr><td>Cotton tipped applicators</td><td>Yibon medical co</td><td></td><td></td></tr><tr><td>Dissecting forceps</td><td>Lawton</td><td>09-0190</td><td></td></tr><tr><td>Elastic retractors</td><td>Odense University Hospital</td><td></td><td></td></tr><tr><td>Electrical clipper</td><td>Aesculap</td><td>GT420</td><td></td></tr><tr><td>Fentanyl 0,315 mg/ml</td><td>Matrix</td><td></td><td></td></tr><tr><td>Heating pad - PhysioSuite</td><td>Kent Scientific Corp.</td><td></td><td></td></tr><tr><td>Isoflurane 1000mg Attane</td><td>Scan Vet</td><td></td><td></td></tr><tr><td>Isoflurane vaporizer - PPV</td><td>Penlon</td><td></td><td></td></tr><tr><td>Micro jewler forceps</td><td>Lawton</td><td>1405-05</td><td></td></tr><tr><td>Micro Needle holder</td><td>Lawton</td><td>25679-14</td><td></td></tr><tr><td>Micro scissors</td><td>Lawton</td><td>10128-15</td><td></td></tr><tr><td>Micro tying forceps</td><td>Lawton</td><td>43953-10</td><td></td></tr><tr><td>Microfine U-40 syringe 0,5ml</td><td>BD</td><td>328821</td><td></td></tr><tr><td>Microlance syringe 25g</td><td>BD</td><td></td><td></td></tr><tr><td>Microlance syringe 27g</td><td>BD</td><td></td><td></td></tr><tr><td>Midazolam 5 mg/ml (hameln)</td><td>Matrix</td><td></td><td></td></tr><tr><td>Needle holder - Circle wood</td><td>Lawton</td><td>08-0065</td><td></td></tr><tr><td>Non woven swabs</td><td>Selefa</td><td></td><td></td></tr><tr><td>Opmi pico microscope F170</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Patent blue V - 25 mg/ml</td><td>Guerbet</td><td></td><td></td></tr><tr><td>Scissors - Joseph</td><td>BD</td><td>RH1630</td><td></td></tr><tr><td>Siemens INVEON multimodality pre-clinical scanner</td><td>Siemens pre-clinical solutions</td><td></td><td></td></tr><tr><td>Source of radiation - D3100</td><td>Gulmay</td><td></td><td></td></tr><tr><td>Stata Statistical Software: Release 15</td><td>StataCorp LLC</td><td></td><td></td></tr><tr><td>Temgesic - 0,2 mg</td><td>Indivior</td><td></td><td></td></tr><tr><td>Vet eye ointment - viscotears</td><td>Bausch &amp;amp; Lomb</td><td></td><td></td></tr></tbody></table>

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 procedure has previously been used in three separate experiments. All the experiments were made by different lead investigators who all are co-authors of this article. In all three experiments, great care was taken to adhere to the same procedure as described in this protocol. In all three experiments, secondary lymphedema was induced in one hindlimb while the other hindlimb served as a control. Volumes of the hindlimbs were the primary outcome in all three experiments. <strong class...

Watch this Scientific Journal Video about A Revised Method for Inducing Secondary Lymphedema in the Hindlimb of Mice at JoVE.com

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

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 ...

a-revised-method-for-inducing-secondary-lymphedema-hindlimb

Research

JoVE Journal

Medicine

8.0K Views.  University of Southern Denmark. 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.

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

Intra-lymph Node Injection of Biodegradable Polymer Particles

Generation of adaptive immune response relies on efficient drainage or trafficking of antigen to lymph nodes for processing and presentation of these foreign molecules to T and B lymphocytes. Lymph nodes have thus become critical targets for new vaccines and immunotherapies. A recent strategy for targeting these tissues is direct lymph node injection of soluble vaccine components, and clinical trials involving this technique have been promising. Several biomaterial strategies have also been investigated to improve lymph node targeting, for example, tuning particle size for optimal drainage of biomaterial vaccine particles. In this paper we present a new method that combines direct lymph node injection with biodegradable polymer particles that can be laden with antigen, adjuvant, or other vaccine components. In this method polymeric microparticles or nanoparticles are synthesized by a modified double emulsion protocol incorporating lipid stabilizers. Particle properties (e.g.&#160;size, cargo loading) are confirmed by laser diffraction and fluorescent microscopy, respectively. Mouse lymph nodes are then identified by peripheral injection of a nontoxic tracer dye that allows visualization of the target injection site and subsequent deposition of polymer particles in lymph nodes. This technique allows direct control over the doses and combinations of biomaterials and vaccine components delivered to lymph nodes and could be harnessed in the development of new biomaterial-based vaccines.

Lymph nodes are the immunological tissues that orchestrate immune response and are a critical target for vaccines. Biomaterials have been employed to better target lymph nodes and to control delivery of antigens or adjuvants. This paper describes a technique combining these ideas to inject biocompatible polymer particles into lymph nodes.

Generation of adaptive immune response relies on efficient drainage or trafficking of antigen to lymph nodes for processing and presentation of these ...

Bioengineering

Intralymphatic Immunotherapy and Vaccination in Mice

Vaccines are typically injected subcutaneously or intramuscularly for stimulation of immune responses. The success of this requires efficient drainage of vaccine to lymph nodes where antigen presenting cells can interact with lymphocytes for generation of the wanted immune responses. The strength and the type of immune responses induced also depend on the density or frequency of interactions as well as the microenvironment, especially the content of cytokines. As only a minute fraction of peripherally injected vaccines reaches the lymph nodes, vaccinations of mice and humans were performed by direct injection of vaccine into inguinal lymph nodes, i.e.&#160;intralymphatic injection. In man, the procedure is guided by ultrasound. In mice, a small (5-10 mm) incision is made in the inguinal region of anesthetized animals, the lymph node is localized and immobilized with forceps, and a volume of 10-20 &#956;l of the vaccine is injected under visual control. The incision is closed with a single stitch using surgical sutures. Mice were vaccinated with plasmid DNA, RNA, peptide, protein, particles, and bacteria as well as adjuvants, and strong improvement of immune responses against all type of vaccines was observed. The intralymphatic method of vaccination is especially appropriate in situations where conventional vaccination produces insufficient immunity or where the amount of available vaccine is limited.

Prophylactic and therapeutic vaccination often fails to stimulate strong immune responses due to week drainage of the vaccine to lymph nodes and consequently poor involvement of immune cells. By direct injection of vaccine to lymph nodes, so-called intralymphatic injection, vaccine efficacy can be strongly improved and vaccine doses can be reduced.

Vaccines are typically injected subcutaneously or intramuscularly for stimulation of immune responses. The success of this requires efficient drainage of ...

Immunology and Infection

An Alternant Method to the Traditional NASA Hindlimb Unloading Model in Mice

The Morey-Holton hindlimb unloading (HU) method is a widely accepted National Aeronautics and Space Administration (NASA) ground-based model for studying disuse-atrophy in rodents 4-6. Our study evaluated an alternant method to the gold-standard Morey-Holton HU tail-traction technique in mice. Fifty-four female mice (4-8 mo.) were HU for 14 days (n=34) or 28 days (n=20). Recovery from HU was assessed after 3 days of normal cage ambulation following HU (n=22). Aged matched mice (n=76) served as weight-bearing controls.
Prior to HU a tail ring was formed with a 2-0 sterile surgical steel wire that was passed through the 5th, 6th, or 7th inter-vertebral disc space and shaped into a ring from which the mice were suspended. Vertebral location for the tail-ring was selected to appropriately balance animal body weight without interfering with defecation.
We determined the success of this novel HU technique by assessing body weight before and after HU, degree of soleus atrophy, and adrenal mass following HU. Body weight of the mice prior to HU (24.3 &plusmn; 2.9g) did not significantly decline immediately after 14d of HU (22.7 &plusmn; 1.9g), 28d of HU (21.3 + 2.1g) or after 3 days recovery (24.0 &plusmn; 1.8g). Soleus muscle mass significantly declined (-39.1%, and -46.6%) following HU for 14 days and 28 days respectively (p&lt;0.001). Following 3 days of recovery soleus mass significantly increased to 74% of control values. Adrenal weights of HU mice were not different compared to control mice.
The success of our novel HU method is evidenced by the maintenance of animal body weight, comparable adrenal gland weights, and soleus atrophy following HU, corresponding to expected literature values 2, 7, 8. The primary advantages of this HU method include: 1) ease of tail examination during suspension; 2) decreased likelihood of cyanotic, inflamed, and/or necrotic tails frequently observed with tail-taping and HU; 3) no possibility of mice chewing the traction tape and coming out of the suspension apparatus; and 4) rapid recovery and normal cage activity immediately after HU.

We developed an alternant hindlimb unloading model in mice. The primary advantage of our hindlimb unloading tail-ring method over the conventional Morey-Holton tail-traction technique is a simple straightforward procedure that minimizes stress upon the animal.

The Morey-Holton hindlimb unloading (HU) method is a widely accepted National Aeronautics and Space Administration (NASA) ground-based model for studying ...

Biology

Orthotopic Hind Limb Transplantation in the Mouse

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and translational research in the field of transplantation medicine. A vast array of reagents is available exclusively in this setting, including mono- and polyclonal antibodies for both diagnostic and interventional applications. In addition, a vast number of genotyped, inbred, transgenic, and knock out strains allow detailed investigation of the individual contributions of humoral and cellular components to the complex interplay of an immune response and make the mouse the gold standard for immunological research. 
Vascularized Composite Allotransplantation (VCA) delineates a novel field of transplantation using allografts to replace "like with like" in patients suffering traumatic or congenital tissue loss. This surgical methodological protocol shows the use of a non-suture cuff technique for super-microvascular anastomosis in an orthotopic mouse hind limb transplantation model. The model specifically allows for comparison between established paradigms in solid organ transplantation with a novel form of transplants consisting of various different tissue components. Uniquely, this model allows for the transplantation of a viable vascularized bone marrow compartment and niche that have the potential to exert a beneficial effect on the balance of immune acceptance and rejection. This technique provides a tool to investigate alloantigen recognition and allograft rejection and acceptance, as well as enables the pursuit of functional nerve regeneration studies to further advance this novel field of transplantation.

This novel model for orthotopic hind limb transplantation in the mouse, applying a non-suture cuff technique for super-microvascular anastomosis, provides a powerful tool for in&#160;vivo mechanistic immunological research related to vascularized composite allotransplantation (VCA).

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and ...

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 ...

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. ...

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.

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. ...

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 ...

A Modified Surgical Model of Hind Limb Ischemia in ApoE-/- Mice using a Miniature Incision

The purpose of this study is to introduce and evaluate a modified surgical approach to induce acute ischemia in mice that can be implemented in most animal laboratories.&#160;Contrary to the conventional approach for double ligation of the femoral artery (DLFA), a smaller incision on the right inguinal region was made to expose the proximal femoral artery (FA) to perform DLFA. Then, using a 7-0 suture, the incision was dragged to the knee region to expose the distal FA. Magnetic resonance imaging (MRI) on bilateral hind limbs was used to detect FA occlusion after the surgery. At 0, 1, 3, 5, and 7 days after the surgery, functional recovery of the hind limbs was visually assessed and graded using the Tarlov scale. Histologic evaluation was performed after euthanizing the animals 7 days after DLFA. The procedures were successfully performed on the right leg in ten ApoE-/- mice, and no mice died during subsequent observation. The incision sizes in all 10 mice were less than 5 mm (4.2 &#177; 0.63 mm). MRI results showed that FA blood flow in the ischemic side was clearly blocked. The Tarlov scale results demonstrated that hind limb function significantly decreased after the procedure and slowly recovered over the following 7 days. Histologic evaluation showed a significant inflammatory response on the ischemic side and reduced microvascular density in the ischemic hind limb. In conclusion, this study introduces a modified technique using a miniature incision to perform hind limb ischemia (HLI) using DLFA.

A Modified Surgical Model of Hind Limb Ischemia in ApoE-/- Mice using a Miniature Incision

This article demonstrates an efficient surgical approach to establish acute ischemia in mice with a small incision. This approach can be applied by most research groups without any laboratory upgrades.

The purpose of this study is to introduce and evaluate a modified surgical approach to induce acute ischemia in mice that can be implemented in most ...

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; ...

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

Summary

Explore More Videos