Isoflurane (2–2.5%) was used as anesthetic for all procedures. Following subcutaneous injection of 0.1 mL of 10% Evans blue (Sigma-Aldrich, Saint Louis, Mich.) into the dorsum of paw, an obliquely oriented incision was made in the midportion of the groin. Lymphatic vessels, inguinal lymph nodes, and popliteal nodes were identified using Evans blue dye staining. The groin fat pad (4 × 2 cm) was completely resected to remove the inguinal lymph nodes. Along the dyed lymph vessel, popliteal lymph node could be found and resected (Figs. 2B, C). One week after lymphadenectomy, rats were treated with different radiation doses, 20, 30, and 40 Gy radiation as a single dose (Varian 2100 EX Linear Accelerator; Medical Imaging Resources, Ann Arbor, Mich.), with the effective field of 3 × 4 cm and a depth of 1.5 cm. Lymphedema formation was quantitatively evaluated by micro-CT imaging per month. The rats were followed up for 4 months.
CT Imaging Study
Rat hind limb volume was measured with micro-CT imaging preoperatively and every month following radiation treatment for 4 months. After general anesthesia with isoflurane was induced, animals were imaged with the NanoSPECT/CT (Bioscan, Washington, D.C.) in a supine position. The 8-cm transaxial field of view (FOV) was capable of imaging both extremities and lower abdomen. A 15-minute scan was performed for all CT images (65 kVp, exposure time of 1000 ms, and 123 μA). Approximate radiation exposure was 22 mGy, which has been shown to not cause any known biological damage to the animal.13 Micro-CT scan images were reconstructed with filtered back-project into a 3D-image volume with pixel size of 0.2 mm in both the transverse and axial directions. All images were saved in Digital Imaging and Communications in Medicine (DICOM) format and later analyzed with appropriate software (PMOD Technologies, Zurich, Switzerland). The imaging analysis was performed by an experienced staff with the following steps. First, after images were loaded into PMOD, images of one limb were rotated and realigned so that the cross-sectional images were perpendicular to the tibia. Second, a spherical mask was placed upon the thigh to delineate the thigh from the abdomen. Once that is done, an automatic segmentation was performed with PMOD segmentation capability (Fig. 3). Volume of the segmented limb was computed automatically and exported from PMOD. Following this, the same procedure was repeated for the other limb. As a result, the volume of each limb was calculated and later used in the statistical analysis to evaluate the lymphedematous extremity. Then, the volume differentiation could be calculated, which was defined as the volume of the lesioned limb minus the volume of the healthy limb and then divided by the volume of the healthy limb multiplied by 100.
Tc99 nanocolloid was performed to evaluate the function of the lymphatic system in the lower extremities in 2 rats in each group. The same NanoSPECT/CT camera was used for the data acquisition by using the SPECT camera to image animals in a 2D planar scintigraphy mode. For each animal, 0.5 mCi of Tc99 nanocolloid was injected subcutaneously in the paw of each side. The tracer volume was 0.1 mL. The image acquisition was performed with 3 separate scans. Immediately after injection, a dynamic planar acquisition was started with 10 seconds per frame for 30 minutes. The 25-cm field of view of the SPECT acquisition covers the lung, the heart, the abdomen, the lower limbs, and the injection sites. After this scan, the animal was taken off the scanner and anesthetized. At the fifth hour post injection, the animal was rescanned with sixty 10-second frames for 10 minutes and then taken off the scanner. Image acquisition was repeated again for the same animal at 24-hour post injection with 20-minute acquisition of 120 10-second frames. The images were again analyzed with PMOD.
The data were analyzed and compared using the Kruskal-Wallis test. Results were expressed as mean ± SD. A P-value of less than 0.05 was used as the criterion for statistical significance.
Lymphedema Rat Model
A total of 71 rats were used, and the mortality and morbidity rate was 38%. The overall success rate of development of lower limb lymphedema was 57.7%. Success rate in the development of lymphedema was highest in group IC (50%) compared to group IB (37.5%) and group IA (0%) (Table 1). As a result of higher total morbidity and mortality rates in groups IB (56.3%) and IC (50%), we removed both inguinal lymph nodes and popliteal lymph nodes that were identified by injection of Evans blue (Fig. 2) and treated with 20 Gy radiation (group II) with minimal morbidity for evaluating the effect of developing hind limb lymphedema (Fig. 3). The results showed that the success rate of development of hind limb lymphedema in group II was higher (22/27, 81.5%) than group I, and the volume differentiation in group II was 7.76% ± 1.94% (P < 0.01). In addition, we followed up the survived rats for 4 months, with the volume differentiation maintaining around 5%.
Lymphoscintigraphy was used to verify the lymphatic drainage every month postoperatively for 3 months. The image was detected after 5 minutes, 15 minutes, 25 minutes, 5 hours, and 24 hours following Tc99 nanocolloid injection. The Tc99 signal was seen in green and red colors. The results showed that the position of lymph nodes could be seen in the healthy limb in 5 minutes. However, after 5 and 24 hours, the Tc99 nanocolloid was accumulated in the lesioned limb in the group II (Fig. 4).
The importance of developing a reliable and consistent animal model to investigate extremity lymphedema is particularly valuable as clinical treatment modalities continue to improve. As awareness of this disease process continues, translating clinical parameters to an animal model will allow clinicians to investigate and optimize treatment strategies. Previous small animal models have been described and investigated.12,14–17 But, inconsistencies in results and methodology make these difficult to implement in the experimental setting. In addition, the quantification methods were inconsistent and inaccurate in previous studies. Therefore, we modified the structural techniques from previous studies.12,18
First, to determine a consistent radiation dose, varying doses of radiation therapy were used (20, 30, and 40 Gy) to treat the inguinal lymph nodes removed in rats. Although 40 Gy would have the highest success rate for developing lower limb lymphedema, this dose proved to be too toxic causing a high rate of morbidity and mortality. In a similar model described by Kanter et al,12 a 5- to 10-mm skin gap was left during wound closure. The inflammatory response induced appreciable acute lymphedema in the rat hind limb after 2 days. In our model, no skin gap is left and direct skin closure is performed. In addition, 45 Gy was used with success in the development of lymphedema in the previous rat hind limb model.12 Although successful in their initial study, we found this dose to be toxic with considerable morbidity. The lower radiation dose (20 Gy) did not create significant toxicity but was unable to initiate lower limb lymphedema with isolated inguinal lymph node removal. Therefore, both inguinal and popliteal lymphadenectomy were performed with the addition of a lower radiation dose. This sequence proved to be successful in creating a lymphedematous extremity.
In addition to the modified technique for improving induction of lymphedema, volume differentiation between limbs determined by micro-CT images proved to be successful and precise in evaluating volumetric parameters in rats. In a previously reported lymphedema model, volume estimations were made by limb circumference measurements and water displacement volumetry,12,15,17 but the studies showed high variability between control and the lymphedema areas. In another study, T2-weighted magnetic resonance imaging was also used for measuring the volume of the fluid accumulation in the subcutaneous tissue, but this methodology has significant associated costs.19 In the presented study, micro-CT was able to accurately and reliably determine volume differentiation between lesioned and healthy limbs with the benefit of decreased costs compared to magnetic resonance imaging. In addition, for specific evaluation of lymphatic flow, lymphoscintigraphy by injected Tc99 nanocolloid was effective at determining the success of lymphadenectomy and progression to lymphedema. These combined imaging techniques will make it possible to determine the physiologic basis of treatment strategies such as VLN transfer.
In this study, we successfully used low radiation doses combined with the removal of inguinal and popliteal lymph nodes to develop lymphedema in the lower limb up to 4 months with minimal donor-site morbidity. The utilization of micro-CT to assess volumetric changes of the lymphedema in the rat model is reproducible and accurate.
1. Warren AG, Brorson H, Borud LJ, et al. Lymphedema: a comprehensive review. Ann Plast Surg. 2007;59:464–472
2. Hadamitzky C, Pabst R. Acquired lymphedema: an urgent need for adequate animal models. Cancer Res. 2008;68:343–345
3. Lin CH, Ali R, Chen SC, et al. Vascularized groin lymph node transfer using the wrist as a recipient site for management of postmastectomy upper extremity lymphedema. Plast Reconstr Surg. 2009;123:1265–1275
4. Yamamoto T, Koshima I, Yoshimatsu H, et al. Simultaneous multi-site lymphaticovenular anastomoses for primary lower extremity and genital lymphoedema complicated with severe lymphorrhea. J Plast Reconstr Aesthet Surg. 2011;64:812–815
5. Chang DW. Lymphaticovenular bypass for lymphedema management in breast cancer patients: a prospective study. Plast Reconstr Surg. 2010;126:752–758
6. Cheng MH, Chen SC, Henry SL, et al. Vascularized groin lymph node flap transfer for postmastectomy upper limb lymphedema: flap anatomy, recipient sites, and outcomes. Plast Reconstr Surg. 2013;131:1286–1298
7. Cheng MH, Huang JJ, Nguyen DH, et al. A novel approach to the treatment of lower extremity lymphedema by transferring a vascularized submental lymph node flap to the ankle. Gynecol Oncol. 2012;126:93–98
8. Chen HC, O’Brien BM, Rogers IW, et al. Lymph node transfer for the treatment of obstructive lymphoedema in the canine model. Br J Plast Surg. 1990;43:578–586
9. Tobbia D, Semple J, Baker A, et al. Lymphedema development and lymphatic function following lymph node excision in sheep. J Vasc Res. 2009;46:426–434
10. Lähteenvuo M, Honkonen K, Tervala T, et al. Growth factor therapy and autologous lymph node transfer in lymphedema. Circulation. 2011;123:613–620
11. Mendez U, Stroup EM, Lynch LL, et al. A chronic and latent lymphatic insufficiency follows recovery from acute lymphedema in the rat foreleg. Am J Physiol Heart Circ Physiol. 2012;303:H1107–H1113
12. Kanter MA, Slavin SA, Kaplan W. An experimental model for chronic lymphedema. Plast Reconstr Surg. 1990;85:573–580
13. Badea CT, Drangova M, Holdsworth DW, et al. In vivo small-animal imaging using micro-CT and digital subtraction angiography. Phys Med Biol. 2008;53:R319–R350
14. Mendez U, Brown EM, Ongstad EL, et al. Functional recovery of fluid drainage precedes lymphangiogenesis in acute murine foreleg lymphedema. Am J Physiol Heart Circ Physiol. 2012;302:H2250–H2256
15. Serizawa F, Ito K, Matsubara M, et al. Extracorporeal shock wave therapy induces therapeutic lymphangiogenesis in a rat model of secondary lymphoedema. Eur J Vasc Endovasc Surg. 2011;42:254–260
16. Pan D, Han J, Wilburn P, et al. Validation of a new technique for the quantitation of edema in the experimental setting. Lymphat Res Biol. 2006;4:153–158
17. Liu Y, Fang Y, Dong P, et al. Effect of vascular endothelial growth factor C (VEGF-C) gene transfer in rat model of secondary lymphedema. Vascul Pharmacol. 2008;49:44–50
18. Ogata F, Azuma R, Kikuchi M, et al. Novel lymphography using indocyanine green dye for near-infrared fluorescence labeling. Ann Plast Surg. 2007;58:652–655
© 2014 American Society of Plastic Surgeons
19. Sommer T, Meier M, Bruns F, et al. Quantification of lymphedema in a rat model by 3D-active contour segmentation by magnetic resonance imaging. Lymphat Res Biol. 2012;10:25–29