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Method Article

A Model of Acute Lung Injury Following Visceral Ischemia-Reperfusion by Supra-Coeliac Aortic Cross Clamping in Rats

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DOI:

10.3791/66896

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August 15th, 2025

 ,  ,  ,  ,  , 
1Department of Vascular Surgery, Rouen University Hospital, 2INSERM U1096, FHU REMOD-VHF, Rouen University Hospital

In This Article

Summary

This study describes a model of acute lung injury following visceral ischemia-reperfusion during supra-celiac aortic cross-clamping via laparotomy in rats. It enables the investigation of both the clinical and biological effects of aortic clamping as well as the impact of ischemia-reperfusion on underlying organs and remote lung injuries.

Abstract

Open surgery for abdominal aortic aneurysms requires transient interruption of blood circulation, causing ischemia-reperfusion of downstream organs. Renal, mesenteric, and splanchnic arteries may be involved. No therapy has proven effective in preventing remote lung injury caused by ischemia-reperfusion after supra-coeliac aortic clamping. To study the pathophysiological pathways of acute lung injury and pharmacological treatments, an experimental model of aortic clamping has been developed in rats.

Subcutaneous analgesia with Buprenorphine (0.05 mg/kg) is administered before the procedure. General anesthesia with spontaneous ventilation is induced using halogen gas (Isoflurane 2.5%) via a mask. A xypho-pubic laparotomy is performed on a heated table (temperature monitored intrarectally). The gastro-splenic block and intestines are retracted to the right flank, and the liver is mobilized to the upper abdomen. The diaphragm and its pillars are identified to allow step-by-step release of the supra-coeliac aorta from its intradiaphragmatic passage. Dissection is performed gently to achieve circumferential control of the aorta between the diaphragm pillars. The underlying celiac trunk and superior mesenteric artery are carefully identified. A microvascular clamp may be used for aortic occlusion above the visceral arteries. Proper clamping is verified by the absence of aortic pulsatility, associated with pallor of the small intestine. The rat remains anesthetized throughout the ischemia duration. At the end of the ischemia duration, the microclamp is removed, and aortic pulsatility, as well as intestinal recoloration, are checked. The abdominal wall is closed with a suture, followed by closure of the rat's skin. At the end of the reperfusion period, the animal is anesthetized in the same manner. It is euthanized by exsanguination during blood sampling, and the lung can then be collected. Lung injury can be assessed through blood gas analysis, lung gravimetry, and histological examination. Lung samples can also be utilized for RT-PCR and Western blot analyses.

Introduction

An aortic aneurysm is a focal and permanent dilation of the aorta1. The evolution of an aneurysm tends towards rupture, associated with massive hemorrhage, which carries significant mortality2. Therefore, the focus of treatment lies in the prevention of this dramatic complication. Aortic aneurysms are most often infrarenal and can be treated with aortic endoprosthesis. An aorto-bi-iliac endograft is a covered bifurcated stent that is introduced in a minimally invasive manner through the femoral arteries using a percutaneous approach. This technique does not require aortic clamping and avoids causing visceral ischemia. It can be performed under local anesthesia. In contrast, more extensive aortic aneurysms-such as thoracoabdominal aneurysms and complex abdominal aortic aneurysms-are more challenging to treat using endovascular techniques. This is due to the involvement of the renal arteries, the superior mesenteric artery, and the celiac trunk within the aneurysmal segment (Figure 1). These major visceral arteries must be reconstructed in addition to the aorta itself. The endovascular, minimally invasive approach therefore, requires a more complex procedure involving the use of fenestrated and/or branched endografts for visceral artery reconstruction. The main limitation of these endografts is the long manufacturing time -- often several months -- making them unsuitable for emergency treatment. In such cases, conventional open surgical repair remains a feasible and reliable option.

The treatment by open repair requires laparotomy and supra-visceral clamping, i.e., temporary visceral ischemia. The complication rate is high during the treatment of these thoracoabdominal aneurysms. Mortality ranges from 5 to 15%3. Pulmonary complications occur in 60% and renal complications in 40%4. These complications, associated with a systemic inflammatory response, can progress to multi-visceral failure5. This morbi-mortality is multifactorial, involving surgical aggression, anesthesia, analgesia, hemodynamic variations, hemorrhagic shock, and ischemia-reperfusion phenomena. Surgical aggression contributes to the manifestation of complications, whether the approach includes thoracotomy, phrenotomy, lumbotomy, or laparotomy, by modifying the ventilatory mechanics, with in particular an impairment of the diaphragmatic function6.

The temporary visceral ischemia reperfusion affects downstream organs, including the digestive tract, splanchnic system, kidneys, and lower limbs7. Pulmonary complications are the most frequent adverse events following thoracoabdominal aortic surgery. Their origin is multifactorial. Supraceliac aortic clamping does not directly cause pulmonary ischemia-reperfusion. However, ischemia-reperfusion of the downstream visceral organs triggers a major systemic inflammatory response. The hypothesis of a causal link between visceral ischemia-reperfusion and the occurrence of remote injury -- particularly lung damage -- is well supported. Pulmonary endothelial dysfunction, a critical component of the alveolar-capillary membrane, appears to be central to the development of Acute Lung Injury (ALI). A massive systemic inflammatory response and the activation of neutrophils play a pivotal role in this process.

ALI is often defined based on histological criteria rather than clinical presentation. ALI refers to an acute inflammatory disorder characterized by the disruption of the pulmonary endothelial and epithelial barriers. The alveolar-capillary membrane, the functional unit of the lung, consists of three components: the microvascular endothelium, the interstitial space, and the alveolar epithelium.

A hallmark of ALI is increased permeability of this barrier, driven by neutrophil infiltration and the release of cytotoxic pro-inflammatory mediators. Alveolar epithelial damage involves both type I and type II pneumocytes. Type I cells, which cover 95% of the alveolar surface, are highly susceptible to injury and play a key role in gas exchange. Type II cells, more numerous but covering a smaller area, are essential for surfactant production, epithelial repair, and ion transport. Following injury, type II cells contribute to the restoration of alveolar integrity through their regenerative capacity and differentiation into type I cells.

Multiple cell death pathways have been implicated in epithelial injury during ALI, with type I cells being particularly vulnerable. ALI is commonly assessed by evaluating pulmonary edema, with or without measures of microvascular permeability to reflect endothelial dysfunction.

The objective of this model is to investigate the link between visceral ischemia-reperfusion and the development of remote lung injury, both at the metabolic and mechanistic levels as illustrated on Figure 2. This specific connection remains only partially understood. Our aortic clamping model represents a visceral ischemia-reperfusion scenario on a healthy aorta, without the presence of an abdominal aortic aneurysm in rat. Above the celiac trunk, there are no major branches in either humans or rats. The aorta above this level corresponds to the supra-cœliac aorta, which gives rise only to small parietal branches supplying the thoracic wall. Therefore, there is no significant difference in the vascular anatomy of the major vessels at this level between humans and rats.

The use of this model allows for extensive exploration. It will help identify early biomarkers in the development of remote tissue injury. Additionally, this model enables the study of potential inhibitors, including inhibitors of circulating mediators or inhibitors of specific remote organs injuries.

Other ischemia-reperfusion models are used to study remote injuries. Cerebral, intestinal, and renal ischemia-reperfusion models are strongly represented8,9,10. There is extensive research on pulmonary damage after intestinal ischemia-reperfusion due to transient occlusion of the superior mesenteric artery (SMA)11,12,13. Ischemia and reperfusion durations are extremely variable, ranging from minutes to several hours14,15. The rationale for the duration of ischemia is to induce sufficient aggression to trigger a significant response leading to the occurrence of remote injuries. A too short duration of ischemia may not cause a significant alteration in the studied metabolic pathway. Regarding reperfusion duration, the rationale lies in the early onset of remote injuries and the potentially transient nature of pathophysiological changes. An ideal reperfusion duration should not be too early or too late to characterize metabolic impairment adequately.

The models are mainly in rodents. Advantages of rodents include their small size, ease of maintenance, short life cycle, and abundant genetic resources16. The use of large animals such as pigs, dogs, or primates does not seem justified in the absence of additional benefits, which aligns with the ethical rules of animal experimentation regarding refinement. Furthermore, due to their anatomical, physiological, and genetic similarity to humans, rodents allow translational studies16.

One of the main limitations of SMA clamping models is that, in clinical practice, the start of the intestinal ischemia period is not identifiable. This makes extrapolation to human clinical situations more uncertain. In contrast, during complex aortic surgery, the ischemia and reperfusion periods of different organs are clearly identified. Moreover, during supra-coeliac aortic clamping, all the underlying viscera are in an ischemic period (digestive tract, liver, spleen, kidneys, pelvic organs, and lower limbs). All of these organs undergo the ischemia-reperfusion phenomenon. Conversely, models of ischemia-reperfusion focused on a specific organ (intestine, kidney, liver, etc.), models of aortic clamping and therefore of ischemia-reperfusion of multiple organs with metabolic pathophysiology are less studied. Similarly, remote injuries such as pulmonary lesions after aortic clamping are poorly studied in terms of metabolism17,18.

Existing aortic clamping models are primarily focused on clamping the abdominal aorta below the renal arteries. The limitation of this model is that it does not induce visceral ischemia-reperfusion. Therefore, it does not allow for the demonstration of a link between visceral ischemia-reperfusion and potential remote injuries18,19. Besides, there is no in vitro model allowing the study of ischemia-reperfusion through aortic clamping, that is why analysis using in vivo models remains necessary.

Therefore, this experimental model of supra-celiac aortic clamping in rats facilitates a comprehensive investigation into the pathophysiological mechanisms and pharmacological interventions targeting remote lung injuries.

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Protocol

The creation of this model complies with the guidelines for animal care and experimentation, as well as French and European legislation. This model has been validated in an experimental animal study that was approved by the Regional University Ethics Committee in Normandy, Rouen, France (n°APAFIS: #18974).

1. Animal model and acclimatization

  1. Animal model adaptation period
    NOTE: The model adheres to the principles of refinement.
    1. House animals in non-overcrowded facilities for one week to allow adaptation before surgery.
    2. House the rats two per cage with ad libitum access to water and food, which will be checked daily by experienced personnel.
    3. Check the cleanliness of the bedding (paper strips) daily and change systematically by experienced personnel twice a week or more if needed.
    4. After a one-week acclimatization period, during which they will have daily contact with the experimenter and exposure to 12-h lighting cycles, procedures will take place in a separate, quiet room with the same experimenter.
  2. Animal model species
    1. Use male Wistar or Sprague Dawley rats to avoid hormonal influences, such as estradiol, on the analysis of visceral ischemia-reperfusion physiopathology from an accredited and certified breeding facility11,20.
    2. Determine the experimental groups and the number of animals per group in advance.
    3. Choose the age, 6-10-week-old rats (350 g) are recommended for use.

2. Anesthesia and analgesia

  1. Analgesia
    1. Weigh each animal on the day of the procedure.
    2. Administer buprenorphine (0.05 mg/kg) subcutaneously at the level of the rat's back skin 1 h before the procedure for analgesic purposes with 25-G needle and 2 mL syringe.
    3. Apply Lidocaine hydrochloride gel 1% to the skin of the abdomen at the level of the future skin incision. Reassess pain regularly (at least twice daily) postoperatively using a scale to assess facial expressions of pain in the rat21.
    4. Administer during the first 24 h, systematic subcutaneous injections of buprenorphine (0.01 to 0.05 mg/kg) every 8 h.
    5. Subsequently, administer buprenorphine injections (0.01 to 0.05 mg/kg) based on the facial pain scale of the National Centre for the Replacement, Refinement, and Reduction of Animals in Research, and repeated every 8 h if necessary, depending on the euthanasia time point21.
  2. General anesthesia
    1. Use halogenated gases (Isoflurane) for general anesthesia22.
    2. Use a ventilated chamber with isoflurane (5 %) at an airflow rate of 2 L/min for 2 min for the induction phase.
    3. For maintenance of anesthesia, switch the rat to a custom-fitted mask for spontaneous ventilation with isoflurane (2-2.5%) at an airflow rate of 2 L/min throughout the surgery.
    4. Clear the upper airways by moving the tongue out of the oral cavity to allow ventilation via the mask.
    5. Continuously monitor anesthesia depth during the procedure and adjust Isoflurane concentration as needed.

3. Rat placement during the procedure

  1. Place the rat in dorsal recumbency with the front and hind limbs immobilized using adhesive tape, while respecting the physiological joint ranges of motion of the rat.
  2. Place the rat on a warming pad set at 37 °C with monitoring of rectal temperature.
  3. Protect the eyes by applying dacryogel. Shave the abdominal fur and thoroughly disinfect the abdomen with 3 iterations of alternating iodine and alcohol scrubbing.

4. Surgical procedure

  1. Condition of the surgical procedure
    1. Perform the procedure under a microsurgical loupe to facilitate delicate maneuvers, although it is not indispensable.
    2. Confirm the depth of anesthesia via a toe-pinch before making the incision.
  2. Incision
    1. Following palpation of the xiphoid process at the upper border of the incision, incise the skin at the center of the abdomen, revealing the linea alba of the abdominal muscles.
    2. Center incision of the abdominal muscle on the linea alba to avoid bleeding from the abdominal muscles.
  3. Intra-abdominal exposition
    1. Maintain the abdominal wall on both sides with self-retaining retractors.
    2. Ensure the absence of anomalies (tumors), mobilize the viscera, stomach, spleen, and liver to the right side of the abdomen, and then expose the supra-celiac aorta.
    3. Kept all viscera inside the abdomen as much as possible by a small compress folded lengthwise and soaked in warm (37 °C) saline solution.
    4. Cover all viscera with a compress soaked in saline solution warmed to 37 °C, to prevent hypothermia and dehydration.
  4. Major intra-abdominal vessels identification
    1. Identify the supra-celiac aorta by its pulsations and its location between the diaphragmatic pillars.
    2. Control of the aorta between the diaphragmatic pillars with gentle and careful dissection using cotton-tipped applicators and dissecting forceps to release the aorta as atraumatically as possible.
    3. Encircle the supra-celiac aorta using a 5-0 suture with a dissecting and ligature forceps.
    4. Verify the origin of the celiac trunk and the underlying mesenteric artery. The duration of this surgical procedure is 10 to 15 minutes per rat.
    5. The underlying identification of the subrenal inferior vena cava is easy due to the transparency of the retroperitoneum.
      NOTE: The supraceliac aorta is identified for cross-clamping, the downstream aorta is exposed to confirm the absence of pulsatility, and the inferior vena cava is used for injection (pre- or postconditioning) and for blood sampling at euthanasia.
    6. Release the subrenal inferior vena cava only on its anterior surface, which allows for direct intravenous administration of any pharmacological agent.
  5. Intravenous injection
    1. For this purpose, use a 30-G needle for injection, and place a few millimeters of hemostatic compress to achieve hemostasis.
    2. If performing pharmacological preconditioning, administer the injection before aortic clamping. For post-conditioning treatment, administer it during or after aortic clamping.
    3. This protocol studied pharmacological preconditioning, so administer 5 min before aortic clamping.
  6. Ischemic period
    1. Clamp the celiac aorta with an atraumatic microvascular clamp for the entire 40-min ischemic period. Verify the effectiveness of the clamp by the disappearance of underlying aortic pulsations and the discoloration of the viscera and limbs multiple times during clamping.
    2. Close partially the abdomen with three simple 5-0 braided absorbable sutures to limit water and heat loss. Cover the surgical incision with a sterile compress soaked in warm saline solution for the entire duration of ischemia.
  7. Reperfusion period
    1. At the end of the ischemic period, reopen the abdomen and expose the aorta in the same manner as described earlier.
    2. Remove the vascular clamp, and verify the resumption of underlying aortic pulsatility and the recoloration of the viscera and limbs. The reperfusion period begins immediately upon removal of the microclamp.
  8. Abdomen closing
    1. Return the viscera to their original position, and close the abdomen with a 5-0 braided absorbable suture on the linea alba and the skin with a 5-0 monofilament absorbable suture.
  9. Animal awakening
    1. Place the rat in a left lateral decubitus position to avoid compressing the inferior vena cava by the visceral mass and the context of hypovolemia. Place the rat in an individual cage with a heat lamp to maintain temperature until awakening, approximately 10 min later.
  10. Second general anesthesia
    1. At the end of the 180-min reperfusion period, anesthetize the rat using the same gas anesthesia through a ventilated chamber with isoflurane (5%) at an airflow rate of 2 L/min for 2 min for the induction phase.
    2. For maintenance of anesthesia, switch the rat to a custom-fitted mask for spontaneous ventilation with isoflurane (2-2.5%) at an airflow rate of 2 L/min throughout the tissue collection period.
    3. Confirm the depth of anesthesia via a toe-pinch before making the incision.

5. Sampling

  1. Blood samples
    1. Collect blood samples using a 23-G needle and a 5 mL syringe by direct puncture into the inferior vena cava below the renal veins.
    2. Use a small portion (100 µL) of the total blood for gas and standard blood analysis on a single room temperature test card.
    3. Collect the remaining whole blood in a 4 mL EDTA tube, and centrifuge it at 1,000 × g for 10 min at +4 °C.
    4. Froze the plasma in cryotubes in liquid nitrogen and store at -80 °C until analysis.
  2. Tissue samples
    1. Tissue sampling
      1. Collect tissue samples (lung, intestine, kidney, liver, and heart) during euthanasia of the animal. Perform euthanasia by blood sampling leading to exsanguination after general anesthesia with isoflurane according to the same protocol as previously mentioned.
      2. Through the open abdomen, harvest 50 mg of each tissue: the upper pole of the left kidney, the left hepatic lobe, and the distal small intestine 1 cm before the cecum.
      3. Then, perform a median sternotomy quickly to identify the heart and lungs.
      4. Free the left lung from its ligament and remove it entirely for histological analysis. Free the right multilobe lung from its ligaments. Separate the anterior lobe for nucleic acid and protein analyses, store the rest in a cup with physiological serum at 37 °C for a few minutes to subsequently perform gravimetry.
      5. Extract the heart as a whole block with sections of the aorta, pulmonary artery, and both vena cavae. Sample a piece of the left ventricle for nucleic acid and protein analyses, store the remainder in a cup with physiological serum at 37 °C for a few minutes to subsequently perform gravimetry.
    2. Nucleic acid analysis
      1. For nucleic acid analysis, collect immediately a 50 g portion of tissue. Freeze it in liquid nitrogen and then store it at -80 °C until analysis.
      2. Extract total RNA with ready-to-use reagent for the isolation of high-quality total RNA according to the manufacturer's protocol.
      3. Determine RNA concentration and purity with a spectrophotometer. Measure RNA concentration using 2 µL of sample, with 2 µL of RNase-free water as a blank reference. Use the absorbance at 260 nm to determine RNA concentration.
        NOTE: Acceptable RNA purity was defined as a 260/280 and 260/230 absorbance ratio ranging between 1.7 and 2.0.
      4. Use 1 µg of each RNA sample for reverse transcription.
      5. Measure the mRNA levels of pro-inflammatory cytokines (IL-1beta, TNF-alpha, IL-6) and an internal reference (beta2-microglobulin), by real-time polymerase chain reaction (RT-PCR) in a rapid, plate-based, thermal block cycler with integrated real-time, online detection capabilities using ready-to-use hot start reaction mix (Primers are listed in Table 1).
      6. Calculate the relative mRNA level using the 2-ΔΔCT method.
    3. Protein analysis
      1. For protein analysis, place 50 mg of tissue in a 2 mL lysing tube containing fifty 2.8 mm zirconium oxide ceramic beads. This lysing tube contains 1 mL of radioimmunoprecipitation assay (RIPA) lysis buffer with a protease inhibitor tablet and a phosphatase inhibitor tablet.
      2. Extract protein from the samples by homogenization on the laboratory tube mill with 3 repetitions of 5-s checking, followed by a 10-s resting period.
      3. Centrifuge the samples at 10,000 g at +4 °C for 10 min, and store the supernatants in cryotubes at -20 °C until analysis.
      4. Determine protein concentrations with a Bradford-based method protein assay and keep samples at −80 °C.
      5. Separate an equal amount of proteins from each sample using SDS-PAGE.
      6. Transfer separated proteins on 0.2 µm nitrocellulose membranes.
      7. Incubate the membrane after blocking with 5% TBS-Tween non-fat dry milk for 1 h, with a primary antibody against the specific protein of choice. For example, to explore heparanase, use a rabbit polyclonal primary antibody against human heparanase (dilution 1:2000) and a control protein as GAPDH, use a goat monoclonal primary antibody against GAPDH (dilution 1:5000) overnight at 4 °C.
      8. Wash the blots three times with TBST-Tween 0.1% buffer.
      9. Incubate with an HRP-conjugated secondary antibody directed against the primary antibody at room temperature (RT) for 1 h in blocking solution. For example, to explore heparanase, use the goat against rabbit secondary antibody (dilution 1:5000) and to explore GAPDH, use the rabbit against goat secondary antibody (dilution 1:10000).
      10. Detect the immune complexes using enhanced chemiluminescence (ECL) detection system.
      11. Analyze the membranes using dedicated software to determine the densitometric values of the target bands, using GAPDH as the internal control. Perform densitometric analysis of the band corresponding to heparanase (50 kDa), and normalize it to the GAPDH protein band (35 kDa) to determine the relative signal intensity.
    4. Histological analysis
      1. For histological analysis, take the left unilobar lung at the hilum, then infuse through the left main bronchus with a cryopreservation solution using a 16 G catheter. Freeze it in liquid nitrogen and store it at -80 °C.
    5. Gravimetry analysis
      1. After the samples for nucleic acid and protein analysis, analyze the remaining portion of the heart and right multilobular lung by gravimetry, with desiccation in ovens at 70°C, allowing the wet/dry weight ratio to provide an estimate of tissue edema.

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Results

This protocol was found to be safe, with minimal animal losses (less than 5%) during anesthesia and surgery, and effective for the induction of acute lung injury. Anesthesia with isoflurane was safer than ketamine combined with xylazine anesthesia (loss approximately 15%), with rapid awakening and better rat recovery (data not shown). All anesthetized animals were able to undergo safe and effective aortic clamping with this method. All animals (100%) with effective aortic clamping survived the periods of ischemia and rep...

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Discussion

This surgical model of aortic cross-clamping offers the advantage of allowing a comprehensive analysis, both tissue-based and plasma-based, of the acute lung injury following visceral ischemia-reperfusion. It also provides the opportunity to test pharmacological molecules with intraperitoneal, enteral, or intravenous administration in pre- or post-conditioning.

One of the critical steps remains supraceliac aortic clamping during surgery. When mastered, this step can be performed quickly. The l...

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Disclosures

The authors have nothing to disclose.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
0.2 µm nitrocellulose membranes Trans-Blot Turbo Transfert PackBio-Rad16201500.2 µm nitrocellulose membranes precut
1cc Luer Slip syringe w/o NeedleTerumoMDSS01SE1cc Luer Slip syringe w/o Needle
2cc Luer Slip syringe w/o NeedleBecton Dinckinson2cc Luer Slip syringe w/o Needle
4 mL EDTA tube BD K3 13 mm x75 mmBD VacutainerBD3688605 mL EDTA tube BD K3 13 mm x 75 mm
5-0 braided suture / Vicryl 5-0 (Ethicon) or Novosyn (B. Braun)Ethicon - Johnson and JohnsonV7970EVicryl 5-0 Needle Taper Point 1/2 circle 13 mm Lenght 45 cm
5-0 monofilament absorbable suture / Monocryil PoliglecaproneEthicon - Johnson and JohnsonY493GNeedle 3/8 circle Lenght 18in
5cc Luer Slip syringe w/o NeedleTerumoSS-05LE15cc Luer Slip syringe w/o Needle
Atraumatic microvascular clamp/Disposable vessel clamps for veinAROSurgicalTKF-1-20gAtraumatic microvascular clamp for vein use in order to clamp the rat aorta
BD Microlance 3 23 G x 1 0.6 x 25 mmBD Vacutainer300800BD Microlance 3 23 G x 1 0.6 x 25 mm
BD Microlance 3 25 G x 5/8 0.5 x 16 mmBD Vacutainer300600BD Microlance 3 25 G x 5/8 0.5 x 16 mm
BD Microlance 3 30 G x 1/2 0.3 x 13 mmBD Vacutainer304000BD Microlance 3 30 G x 1/2 0.3 x 13 mm
beta-actine InvitrogenCarlsbad, USA15G5A11/E2
Bradford-based method protein assayBiorad, Marnes-la-Coquette, France5000201
BuprenorphineBuprenorphine
Centrifugator Sigma 3-30Sigma, Germany10375, 10376, 10377, 10378, 10379, 91276, 109014Centrifugator for eppendorf
cOmplete MiniRoche11836153001protease inhibitor cocktail 25 tablets
Cotton swab
Dacryogel /  Ocular gel Lubrithal 10 gDechra Veterinary productsCode ACL7376464Ocular gel Lubrithal 10 g
Disposable scalpelParamountPDSS11Disposable scalpel blade n°11
ElectrocauteryDentRMedEMB/87400Hemostatic pen with hight temperature
Enhanced ECL Detection SystemBiorad, Hercules, California, USA1705060S
Fisherbrand Bead Mill 24Fisher scientific15515799Used for  grinding, lysis and homogenization of biological samples prior to molecular extraction
Gauze swabs Raffin.010106Gauze swabs 
ImageLabTM softwareBioradon USB drive or license
Isoflurane 5%Isoflurane 5%
Lidocaine Prilocaine gel 5% Aguettant, Lyon, FranceLidocaine Prilocaine gel 5% 
LightCycler 480 InstrumentRoche, Bâle, Swiss5015243001thermal cycler
Lubricating Jelly Sterile K-Y 82 gRECKITT BENCKISERLubricating Jelly Sterile K-Y 82 g
NanoDrop 2000TermoScientific, Waltham, Massachusetts, USAND-2000spectrophotometer
Peha Pince anatomicHartmann991063Anatomic pliers
PhosSTOPRoche4906837001phosphatase inhibitor 20 tablet 
Plaster/Adhesive TapeAnapore135320Surgical adhesive tape/plaster
Povione-iodine solution / Vétédine SolutionVétoquinolAntiseptic solution for external use
Precellys Lysing Kit tube Hard tissue homogenizing CK28, BertinCK28Precellys Lysing Kit tube 
Prolène Prolypropylene sutureEthicon - Johnson and Johnson7472HProlène 5-0 Needle Taper cut 3/8 circle 13 mm Lenght 75 cm
Safe-Lock tubes 1.5 mLEppendorf.0030120086Safe-Lock tubes 1.5cc no color 1000 tubes for centrifugation
Safe-Lock tubes 2 mLEppendorf.0030120094Safe-Lock tubes 2.0cc no color 1000 tubes for centrifugation
Safe-Lock Tubes 5 mLEppendorf.0030119401Safe-Lock tubes 5.0cc no color 200 tubes for centrifugation
Saline solution NaCl 0,9%Aguettant, Lyon, France600502Saline solution 0.9% 500 mL
SDS-PAGESigma-Alrich
Surflash IV Catheter 16 GTerumoSR*FF1651
SurgicelEthicon - Johnson and JohnsonHemostatic resorbable swabs
Suture set n°12Hartmann4701762Suture sterile set with adhesine champ, cisors, Adson pliers, needle-holder
SYBR Green KIT Light Cycler 480 SYBR Green I MasternRoche4707516001SYBR Green
TBST-Tween 0.1% buffer Sigma-Alrich
Test-Cards Epoc 24 cards 13 parametersSiemens-Healthineers
Tissue TEKSakura FinetecCryopreservation solution 
TRIzol reagent InvitrogenThermo Fisher Scientific, Carlsbad, California, USA15596026TRIzol reagent

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