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

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 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 12^th 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.

Lymphedema is a chronic condition that deserves special attention, owing to its worldwide incidence, lack of curative and standardized treatment, and serious impact on patients' quality of life^1,2.

In developed countries, lymphedema is mainly acquired and is secondary to breast cancer, owing to the high prevalence of this malignancy; the cumulative incidence of breast cancer-related lymphedema 10 years after surgery can reach up to 41.1%³. However, diseases such as melanoma, gynecological cancers, genitourinary tumors, and head and neck neoplasms are also associated with a high incidence of this disease⁴. Regional lymph node resection, as part of the necessary oncological treatment to increase survival rates, leads to the disruption of functional lymphatic drainage. In some cases, this results in compensatory mechanisms that prevent or delay the onset of lymphedema⁵. However, when chemotherapy and radiotherapy are administered, these mechanisms are not able to compensate for the change, and lymphedema resultantly occurs. This has a negative impact on patients' quality of life, affecting their functional, social, and psychological well-being^6,7.

The need for an effective cure for lymphedema requires understanding of the physiopathology of the lymphatic system, as well as a deep insight into the complex cellular mechanisms and their responses in both normal and dysfunctional lymphatic systems^8,9,10. Such insights can be obtained initially from experimental animal models that can reproduce chronic human diseases¹¹.

Many attempts have been made to replicate lymphedema in experimental animal models; however, most of them have been hindered by some limitations, including the inability to reproduce chronic lymphatic insufficiency in a stable animal model, the costs of the study, and most importantly, the great regenerative capacity of the lymphatic system, which enables it to restore circulation^12,13.

This study presents the experimental approach for surgically inducing stable acquired lymphedema using the rabbit hindlimb. Based on literature review, this animal can be considered optimal for the development of lymphedema because of the consistent anatomy of its hindlimb lymphatic system, which includes a single popliteal node that drains the hindlimb and reaches the main femoral lymphatic system in the leg^14,15.

The specific anatomy of the rabbit's hindlimb allows for the reproduction of the surgical procedures performed in humans to induce secondary lymphedema. Therefore, this procedure can be used for microsurgical training and preclinical research to extrapolate the results to human medicine.

All procedures were approved by the ethical committee of the Jesús Usón Minimally Invasive Surgery Center and the welfare guidelines of the regional government, which are based on European legislation.

1. Presurgical and surgical preparation

1. House nine female New Zealand white rabbits weighing 4-4.5 kg and aged 4 months in separate cages maintained at a temperature of 22-25 °C, with free access to food and water. Make sure that the cages contain a polysulfone tray with a surface area of 3 m² and a height of 40 cm, as well as a bed with wood shavings.
   1. Identify the cages with the project code and animal identification number.
   2. Acclimate the animals for 1 week before surgery to prevent stress-induced problems. Collect preoperative laboratory values of blood samples to ensure that each animal is in good health prior to anesthesia.
2. Ensure all rabbits follow a 12 h fast before each surgical procedure.
   1. After premedication, preoxygenate the rabbits using a face mask (Hall mask) for 5 min with 100% oxygen and a fresh gas flow of 3-5 L/min. Perform the co-induction phase with midazolam (0.3 mg/kg) and propofol (10 mg/kg) intravenously.
3. Intubate the rabbits with 3.0-3.5 endotracheal tubes, with pneumotaponation, connected to a semi-closed circular circuit linked to a ventilator with a flow of fresh gases at 1 L/min for an initial 5 min, and subsequently set at 0.5 L/min.
   1. Perform maintenance anesthesia by inhalation of sevoflurane at a concentration of 3%-3.5% set on the vaporizer.
4. Administer a continuous infusion of Ringer's lactate solution (2-4 mL/kg/h) through the marginal vein of the ear to the anesthetized rabbits throughout the surgical procedure.
   1. Use a protective eye ointment to protect the ocular surface.
5. General anesthesia monitoring: use a rectal thermometer to monitor the temperature at 38.7-39.7 °C, inspect the mucous membrane color, and monitor the O₂saturation at >95% and the heart rate at 180-240 bpm using a rabbit pulse oximeter.
6. Place a thermal support so the animal maintains a constant temperature throughout the procedure.
7. Administer ketorolac (1.5 mg/kg) plus tramadol (3 mg/kg) intravenously for intraoperative analgesia.
8. Administer antibiotics (7.5 mg/(kg∙day) enrofloxacin subcutaneously [s.c.]) before surgery and 5 days after surgery, as well as postoperative analgesia (10 µg/(kg∙day) buprenorphine s.c.) for 5 days.
9. Place the rabbits in the supine position and shave the animal's hindlimb and inguinal areas. Place the animal in dorsal/supine recumbency and clip the hair from the hindlimb and inguinal areas.
10. Perform skin antisepsis by applying 0.5% chlorhexidine and 70% ethanol to the previously shaved skin. Once the area has been disinfected, cover the rabbit with a sterile cloth, except for the left hindlimb.

2. Popliteal vascular lymph node resection surgery (Figure 1)

1. Infiltrate 0.2-0.3 mL of indocyanine green (ICG) intradermally in the second and third interdigital spaces of the left hindlimb. Massage, gently flex, and extend the hindlimb for a few minutes to facilitate the uptake of the dye into the lymphatic vessels. Use the contralateral limb as a control.
2. Use a real-time, near-infrared fluorescence camera to visualize and mark (using a surgical marker) the lymphatic vessels crossing at the knee level and the popliteal lymph node (PLN) on the skin (Figure 2).
3. Inject patent blue V (0.2 mL) into the interdigital area for subsequent identification of the lymphatic vessels and lymph nodes.
4. Once the PLN is identified using a real-time, near-infrared fluorescence camera (Figure 3), create a 2 cm incision in the center of the popliteal fossa, longitudinal to the long axis of the hindlimb through the ischial vein, which is visible through the skin.
   1. To obtain real-time panoramic images of the lymphatic system with the real-time, near-infrared fluorescence camera, use the optical head equipped with a class 1 laser as the excitation light source and a near-infrared sensitive camera, from the ankle to the knee of the hindlimb of the animal.
   2. Visualize the lymphatic vessels above the muscle fascia by resecting the subcutaneous fat⁵. The lymphatic vessels appear blue due to the patent blue V staining in step 2.3.
   3. Use microsurgical forceps to stretch the incision and expose the PLN, including the vascular and afferent lymphatic pedicles. Ensure clear visibility of all the lymphatic and vascular structures (Figure 4).
   4. Identify the PLN, with a diameter of 0.8 mm, under the ischial vein and between the biceps femoris and medial hamstring muscles.
5. Identify the two main lymphatic vessels on the medial aspect of the PLN. These vessels are located parallel to the distal saphenous vein and divide into a network of microvessels as they approach the PLN (Figure 5).
6. Dissect the lymph node pedicle while avoiding damage to the surrounding tissues and vessels (Figure 6).
7. Ligate the medial artery (a branch of the popliteal artery) and the lateral saphenous vein distally and proximally using 10/0 nylon non-absorbable sutures.
8. Identify and cauterize the two groups of afferent lymphatic vessels that directly join the femoral lymphatic system within the thigh, but do not enter the PLN (Figure 7).
   NOTE: The first group corresponds to the medial afferent lymphatic vessels that drain lymph from the upper leg and calf. The second group is composed of lymphatic vessels in the lower extremity musculature. These vessels run along the gastrocnemius muscle, together with the saphenous vein.
9. Confirm complete disruption of the lymphatic system by repeating real-time near-infrared fluorescence imaging.
10. Remove surrounding fatty tissue entirely to avoid possible lymphangiogenesis.
11. Suture the skin incision with 4-0 polyglycolic acid (PGA) absorbable braided sutures (with a 16 mm 3/8 triangular needle) using a continuous intradermal pattern to avoid postoperative auto-mutilation.
12. House the rabbits individually in cages after surgery; keep them under surveillance and at a room temperature between 16 and 22 °C.

3. Postoperative evaluation

1. Perform postoperative assessments at 3, 6, and 12 months after induction.
2. Anesthetize the rabbits following the steps used previously (steps 1.2-1.7).
3. Measure the perimeters of the hindlimbs of the anesthetized rabbits with a tape measure. Take measurements every 2 cm, with the first point being at the ankle and the last at the knee. Calculate the total volume using the truncated cone formula.
4. Use indocyanine green lymphography (ICG-L) for assessing lymphatic function.
   1. Infiltrate 0.2-0.3 mL of ICG intradermally into the second and third interdigital spaces and gently massage for 1 min to facilitate ICG uptake into the lymphatic vessels.
5. Collect images after 15 min using the near-infrared fluorescence system to assess dermal backflow.
6. Once the follow-ups have been completed, euthanize the rabbit following the same anesthetic protocol as that used in the intervention. Once the desired anesthetic plane is achieved, administer intravenous potassium chloride into the auricular vein at an average rate of 2 meq/kg.

Nine rabbits underwent lymphedema induction in this study, however, three rabbits died during the immediate postoperative period and could not be evaluated. Study data were obtained at 3, 6, and 12 months postoperatively by three independent researchers. Circumferential hindlimb measurements and ICG-L were performed under general anesthesia to assess the lymphatic system function and dermal backflow.

The data obtained by ICG-L at 3 months postoperatively showed dermal backflow and the absence of PLNs in the left hindlimbs of the rabbits, compatible with the disruption of normal lymphatic system function. ICG-L results at 6 and 12 months postoperatively showed dermal backflow and dilated lymphatic channels distal to the popliteal fossa during surgical exploration (Figure 8).

No evidence of clinical lymphedema was found upon examination or circumferential hindlimb measurements. The increase in the volume of the left hindlimb was not significant (p < 0.05) compared to that of the right hindlimb (control) (Figure 9). All results obtained from the left hindlimb were compared with those obtained from the right hindlimb of the same animal, which was used as the negative control.

In the described surgical procedure, after PLN resection and ICG application, no compromised blood flow was observed in the hindlimb, no alteration in vital parameters was reported during anesthesia, and no pain or lameness was described. All parameters were assessed according to the guidelines for quantifying pain, stress, and distress in laboratory animals^16,17,18,19.

Figure 1: Experimental surgical procedure for the induction of lymphedema. Detailed protocol for the creation of a vascular lymph node resection in the left hindlimb of the rabbit for the induction of secondary lymphedema. Abbreviations: LN = lymph node; VLN = vascular lymph node. Please click here to view a larger version of this figure.

Figure 2: Preoperative markings of rabbit hindlimbs. Skin marks on the left hindlimb show the measurement points every 2 cm (blue arrow) and a healthy lymphatic system drawn with a surgical marker (red arrow) observed by ICG-L. Scale bar = 2 cm. Abbreviation: ICG-L = indocyanine green lymphography. Please click here to view a larger version of this figure.

Figure 3: Preoperative ICG-L showing the popliteal lymph node and afferent lymphatic vessels from the rabbit hindlimb. Abbreviation: ICG-L = indocyanine green lymphography. Please click here to view a larger version of this figure.

Figure 4: Lymphatic and vascular structures. Popliteal lymph node (asterisk) with an afferent lymphatic vessel (arrow). Please click here to view a larger version of this figure.

Figure 5: Principal lymph vessels. (A) Identification of the two main lymphatic vessels running parallel to the saphenous vein distal to the PLN by surgical microscopy without ICG. (B) Identification of the two main lymphatic vessels running parallel to the saphenous vein distal to the PLN with ICG. Scale bar = 1 cm. Abbreviations: PLN = popliteal lymph node; ICG = indocyanine green. Please click here to view a larger version of this figure.

Figure 6: PLN pedicle. (A) Identification of the PLN pedicle by surgical microscopy without ICG. (B) Identification of the PLN pedicle with ICG. Scale bar = 1 cm. Abbreviations: PLN = popliteal lymph node; ICG = indocyanine green. Please click here to view a larger version of this figure.

Figure 7: Two groups of afferent lymphatic vessels that do not enter the PLN but directly join the femoral lymphatic system within the thigh. Popliteal lymph node (asterisk) with an afferent lymphatic vessel transected (arrows). Scale bar = 1 cm. Please click here to view a larger version of this figure.

Figure 8: Preoperative and postoperative ICG-L. (A) Preoperative ICG-L showing healthy linear lymphatics in the hindlimb.(B) Postoperative ICG-L showing dermal backflow pattern in the rabbit lymphedematous hindlimb. Abbreviation: ICG-L = indocyanine green lymphography. Please click here to view a larger version of this figure.

Figure 9: Circumferential hindlimb measurements. The red column chart corresponds to the right hindlimb measurements, while the green column chart corresponds to the left hindlimb measurements. The increase in volume of the left hindlimb was not significant (p < 0.05) compared to the control right hindlimb. Please click here to view a larger version of this figure.

Resection of the PLN in an experimental animal is a relatively new procedure that can induce secondary lymphedema in the limbs for assessment and study. After lymph node resection, there is a period of alteration of lymphatic system functionality, lymph accumulation, and histological changes of lymphatic vessels that appear dilated. When this lymph accumulation reaches adequate levels, the characteristic dermal backflow of lymphedema, similar to that observed in humans, can be observed using objective techniques such as ICG-L. The development of this technique may create a cost-effective preclinical research animal, which may help prevent and treat this pathophysiological process.

Intraoperatively, patent blue V application can identify the presence or absence of flow through the lymphatic vessels of the hindlimbs⁵. The disadvantage of this technique is that the lymphatic vessels can only be observed intraoperatively. Therefore, a real-time near-infrared fluorescence camera technology is used, which is an excellent method that allows surgeons to identify, map, and quantify lymph flow in the lymphatic channels of the application site without the need for surgery. Nowadays, this technology is considered the gold standard for lymphedema diagnosis.

In this study, all lymphatic vessels that were observed and identified by ICG and patent blue V were patent. After resection of the PLN, the left hindlimbs presented dermal backflow at 3, 6, and 12 months after induction and linear functional lymphatic vessels were not present. This predicted good lymph stasis in the left hindlimb during the total study period. These results suggest the successful creation of secondary lymphedema in an experimental animal over a longer period than in previous studies⁵. However, this technique has its limitations. To obtain reliable results, all vascular and lymphatic structures of the PLN must be ligated and removed, as well as the surrounding fatty tissue, to avoid possible lymphangiogenesis. In addition, because of the physiology of the lymphatic system in rabbits^15,20, no evidence of clinical lymphedema was demonstrated upon examination or circumferential hindlimb measurements, in contrast to other studies using the same experimental animal⁵.

A real-time near-infrared fluorescence camera was used during follow-up. This technology, with its high sensitivity, represents a valuable technique for real-time mapping of lymphatic vessels and lymph nodes, as verified here. Patent blue V is more widely used but is less specific and limited by the fact that lymphatic vessels can only be observed under intraoperative microscopic vision, whereas ICG allows the detection of these same vessels and lymph nodes without the need for any surgical intervention.

The present study demonstrated that the rabbit is a reliable, reproducible, and affordable experimental animal for the surgical induction of lymphedema. The consistent lymphatic system anatomy of the hindlimbs with a unique PLN permits the induction of long-term and stable lymphedema. This makes the rabbit a useful animal for both lymphedema experimentation and research.

The authors have no conflicts of interest to disclose.

This research project was performed at the Jesús Usón Minimally Invasive Surgery Center (CCMIJU), which is part of the ICTS Nanbiosis. The study was performed with the assistance of the following Nanbiosis units: U21, experimental operating room, and U22, animal housing. This work was supported by Hospital de la Santa Creu i Sant Pau. This work has been partially funded by the Junta de Extremadura, the European Regional Development Fund (Grant Number GR21201). The funder played a role in the study design, data collection, analysis, decision to publish, and manuscript preparation. Special thanks are extended to María Pérez for preparing the figures and to the Microsurgery Department of JUMISC for providing constant encouragement.