All animal activities described here were conducted in accredited facilities and approved by the Institutional Animal Care and Use Committee (IACUC) of the Baden-Württemberg Regional Council in Karlsruhe, Germany (35-9185.81/G-62/23). Experimental animals were managed according to institutional standards, German laws for animal use and care, the directives of the European Community Council (2010/63/EU), and the ARRIVE guidelines. Male Sprague Dawley rats with an order weight of 400 grams were used after being acclimatized for one week. The details of the reagents and the equipment used in this study are listed in the Table of Materials.

1. Anesthesia and analgesia

1. Anesthetize the rat with the medication following the institutional protocols. Isoflurane is recommended for induction of sedation followed by an i.p. injection of 100 mg/kg body weight ketamine and 4 mg/kg body weight xylazine. Additional analgesia can be achieved with a s.c. injection of 5 mg/kg body weight carprofen. For details, please refer to Studier-Fischer et al.¹³.
2. Apply ophthalmic ointment to the eyes to prevent dryness.
3. Guarantee proper analgetic depth using the toe pinch test with surgical forceps.
4. Regularly reassess and adjust the depth of anesthesia during surgery.

2. Procedure preparation

1. Prepare the scrub table with all the required materials and instruments, including releasable microvascular clamps and the applicator (Figure 2A-C), blunt overholt clamp, fine preparation scissor, and forceps.
   1. Prepare the surgical preparation hooks by folding cannulas at an angle of 135° at 1 cm to the tip and connect them to plastic perfusion tubes via Luer-lock ready to apply tension using a surgical mosquito clamp (Figure 2D,E).
2. Prepare a rodent surgical exposure apparatus, including Y-shaped fixation rods and a heating pad, as specified in Studier-Fischer et al.¹³.
3. Ensure proper oxygenation is achieved via the inhalation of 100% oxygen using a neonatal face mask (Figure 2F).

Figure 2: Experimental and animal setup. (A) Surgical instruments and materials required. (B,C) Releasable microvascular clamp and applicator. (D,E) A folded cannula connected to a perfusion tube is used as a surgical preparation hook. (F,G) Rat model oxygenated with a face mask and shaven. (H) Cutaneous incision over complete abdominal length. (I–M) resection of the xiphoid and hemostasis. (N–Q) Hepatic mobilization and dissection of the falciform ligament (arrow 1). (R) Application of preparation hooks and metal stands for exposure of organs after laparotomy. (S) Full visceral exposure of abdominal major vessels using blunt hooks (arrow 2), silicone vessel loops (arrow 3), and surgical compress (arrow 4). (T,U) Abdominal aorta and caval vein. (V) Atraumatic preparation instruments. (W) Humidified cotton swap (arrow 5). (X) Humidified compress in forceps (arrow 6) and blunt overholt clamps (arrow 7). Please click here to view a larger version of this figure.

3. Surgical preparation

1. Shave the surgical access site for a midline laparotomy (Figure 2G). Perform a cutaneous incision over the desired abdominal length, i.e., about 7 cm (Figure 2H), advance with the laparotomy by dissecting the fascia, and stitch the surgical preparation hooks with attached plastic tubes and surgical mosquito clamps through the skin. Expose the surgical situs using the surgical preparation hooks, applying tension to the tissue.
2. Ensure that the peritoneum is dissected cranially only until a few millimeters below the beginning of the sternum, leaving some of the peritoneum intact. Place a part of a surgical compress below the sternum (Figure 2I) and resect the xiphoid with strong material scissors. Apply pressure to the resection area using the surgical compress to achieve sufficient hemostasis of this well-perfused region (Figure 2I-M).
3. Mobilize the liver dorso-caudally to expose the falciform ligament (Figure 2N) and dissect the ligament (Figure 2O-Q).
4. For hemostatic control, depending on where the clamping of the major vessels should be performed, perform a full visceral exposure of abdominal major vessels (Figure 2R) and the vessels slung with silicone vessel loops (Figure 2S-U). Only atraumatic preparation instruments should be used (Figure 2V), such as humidified cotton swaps (Figure 2W), humidified compress in forceps, and blunt overholt clamps (Figure 2X).

4. Preparation and clamping of abdominal aorta for arterial ischemia

1. Perform a left medialization of the upper abdominal organs using atraumatic preparation instruments to gain access to the left adrenal artery (Figure 3A-D).
2. Identify the pulsating site typically medial to the cranial extension of the left adrenal artery, indicating the course of the aorta (Figure 3E). Advance through the soft tissue using overholt clamps for blunt dissection in order to access the abdominal aorta (Figure 3F,G).
3. Tunnel the abdominal aorta at the most cranial end using blunt overholt clamps (Figure 3H-L) and sling the aorta using a silicone vessel loop (Figure 3M-T).
4. Apply a suitable aneurysm microvascular clamp using the silicone loop to slightly luxate the aorta ventrally and guide the aneurysm microvascular clamp along the silicone loop to guarantee isolated aortic clamping (Figure 3U). Depending on the research question, the microvascular clamp can be released again.

Figure 3: Preparation and clamping of the abdominal aorta. (A) Exposure of visceral organs. (B–E) Left medialization of upper abdominal organs using atraumatic preparation instruments to gain access to the left adrenal artery. (F,G) Blunt dissection medial of the left adrenal artery at the pulsating site (gray arrow) in order to access the abdominal aorta. (H–L) Tunneling of the abdominal aorta using blunt overholt clamps. (M–T) Slinging the aorta using a silicone vessel loop. (U) Application of a releasable aneurysm microvascular clamp using the silicone loop as guidance. (V–Z) Visualization of the celiac artery (orange) in reference to the aorta (red) and the silicone vessel loop. Please click here to view a larger version of this figure.

5. Preparation and clamping of suprahepatic abdominal caval vein for venous congestion

1. Mobilize the liver to the right using atraumatic preparation instruments, sharply dissect the hepatic ligaments, and further lateralize the liver (Figure 4A-C).
2. Open the retrohepatic space at the left crus of the diaphragm using blunt overholt clamps (Figure 4D-G).
3. Tunnel the caval vein using blunt overholt clamps (Figure 4H-K) and sling the caval vein using silicone vessel loops (Figure 4L-O).
4. Apply a suitable aneurysm microvascular clamp using the silicone loop to slightly luxate the caval vein ventrally and guide the aneurysm microvascular clamp along the silicone loop to guarantee isolated clamping of the caval vein (Figure 4R).

Figure 4: Preparation and clamping of suprahepatic abdominal caval vein. (A) Exposure of cranial visceral organs. (B) Tissue-protective mobilization of the liver and sharp dissection of hepatic ligaments using atraumatic preparation instruments. (C) Lateralization of the liver. (D–G) The opening of the retrohepatic space and preparation at the left crus of the diaphragm. (H–K) Tunneling of the caval vein (blue) using blunt overholt clamps. (L–O) Slinging the caval vein using silicone vessel loops. (P,Q) Exertion of tension to tentatively restrict caval blood flow. (R) Application of a releasable aneurysm microvascular clamp using the silicone loop as guidance. Please click here to view a larger version of this figure.

6. Clamping of the abdominal aorta and suprahepatic abdominal caval vein for combined malperfusion

1. Perform the steps above until both the aorta and caval vein are slung with silicone vessel loops. Advance with the application of the aneurysm microvascular clamp for both vessels again using the silicone loop for guidance. Clamping the aorta first and minimizing the time required until subsequent caval clamping to only a few seconds is recommended.
   NOTE: Depending on the desired scenario and research purpose, malperfusion can be continued over or released after a defined timespan, and animals can either be euthanized by sharp cardiectomy (following institutionally approved protocols) for non-survival applications, or receive stepwise abdominal closure using surgical sutures in case of planned follow-ups and survival experiments. For the present study, the animals were euthanized.