Mouse Injury Model of Polytrauma and Shock

Abstract

Severe injury and shock remain major sources of morbidity and mortality worldwide. Immunologic dysregulation following trauma contributes to these poor outcomes. Few, if any, therapeutic interventions have benefited these patients, and this is due to our limited understanding of the host response to injury and shock. The Food and Drug Administration requires preclinical animal studies prior to any interventional trials in humans; thus, animal models of injury and shock will remain the mainstay for trauma research. However, adequate animal models that reflect the severe response to trauma in both the acute and subacute phases have been limited. Here we describe a novel murine model of polytrauma and shock that combines hemorrhagic shock, cecectomy, long bone fracture, and soft-tissue damage. This model produces an equivalent Injury Severity Score associated with adverse outcomes in humans, and may better recapitulate the human leukocyte, cytokine, transcriptomic, and overall inflammatory response following injury and hemorrhagic shock.

1. Introduction

Trauma remains one of the leading causes of death in all age groups [1, 2]. Implementation of timely and standardized resuscitative interventions, as well as advances in critical care medicine, have improved early survival in trauma. However, the morbidity and mortality from late complications after severe injury remains high [2–5]. In an effort to identify the etiology and immunologic basis for late multiple organ failure (MOF) that contributes to death after severe trauma, a number of paradigms have been established and revised over the past three decades [5–8]. Chronic critical illness, or CCI, and the persistent inflammation, immunosuppression, and protein catabolism syndrome or PICS, now define a population of ICU patients that survive their initial insult, but remain in the ICU for prolonged periods of time [5, 9] (see Fig. 1). These patients have manageable organ dysfunction, are malnourished despite nutritional intervention, suffer from recurrent infections, and rarely rehabilitate to a functional life [5, 10].

Fig. 1.

Persistent inflammation-immunosuppression catabolism syndrome (PICS). Improved methods of trauma resuscitation have created a population of intensive care unit (ICU) patients who survive their initial insult and early multiple organ failure (MOF) to remain in the ICU for prolonged periods of time with manageable organ dysfunction, malnourishment despite nutritional intervention, suffer from recurrent infections, and rarely rehabilitate to a functional life. Adapted from [5]

There have been numerous attempts at therapeutic interventions meant to improve early survival and prevent the development of PICS after trauma [11, 12]. Unfortunately, efforts at pharmacologic modulation of the immunologic and pathophysiologic response to injury (or infection) in humans have failed, despite promising preclinical results in murine models [11, 12]. This failure to repeat the success of preclinical murine studies in humans has raised questions about the suitability of current murine models of severe inflammation [13–16]. In particular, critics have highlighted the genetic homogeneity of the inbred mouse, the genome-wide differences between man and mice, the heterogeneity of the human condition, as well as the specific variance in cellular composition between murine and human tissues [13, 15]. Thus, researchers are still attempting to improve our methodology for investigating the consequences of severe shock and injury.

Numerous animal models of trauma and shock have been employed over the past century [17–20]. One of the earliest described murine models of trauma was the Noble–Collip drum, described in 1942. In this model, a mouse without anesthesia is tumbled repeatedly in a metal drum, leading to intra-abdominal organ injury, diffuse bruising with muscle injury, and death [20]. Subsequently, researchers developed individual models of hemorrhagic shock, thoracic trauma, brain injury, long bone fracture with soft tissue injury, and intra-abdominal trauma [18, 19, 21–27]. Some of these models have been combined in an attempt to better imitate the human condition (see Table 1). These previous rodent models of trauma and shock have made significant contributions to our understanding of the biological response to injury, yet critical analysis and revision of these models are necessary with technological advances and our expanding knowledge of the biology of inflammation, particularly at the genomic level [15, 16, 18, 28–30].

Table 1.

Examples of murine models of hemorrhagic shock, trauma, burn, or combination

Model name	Modifications to make more similar to human condition
Trauma-hemorrhage (T-H) model followed by cecal ligation and puncture	Two hit model of hemorrhagic shock followed by polymicrobial sepsis, usually 24 h later [46, 47]
T-H model with chronic restraint	Hemorrhagic shock combined with pulmonary contusion and subsequent chronic restraint and stress of the rodent [48]
Pseudofracture with or without T-H	Recreates the features of long bones without breaking the native bone. A bilateral muscle crush injury to the hind limbs is performed, followed by injection of a bone solution into these injured muscles [49]
T-H with fracture and associated soft tissue injury	Hemorrhagic shock combined with a closed mid-shaft fracture of the femur and fracture related soft tissue injury [21]
T-H with fracture and midline laparotomy	Hemorrhagic shock combined with a closed mid-shaft fracture of the femur and midline laparotomy to reflect soft tissue injury [23, 24]
Traumatic brain injury (TBI)	Clinically relevant murine model using an impacting rod directly on the skull [50]
Burn injury followed by infection	Clinically relevant severe burn model that subsequently subjects the mouse to a secondary infection such as pneumonia. This attempts to recreate secondary infections and morbidity and mortality that are common in human burn patients [51]

The criticisms surrounding current murine trauma and shock models are longstanding [15–19, 28, 29], but recent concerns stem from studies that have further highlighted the inherent differences in the murine and human genomic response to inflammatory diseases [13, 14, 16, 31]. The Mouse ENCODE Consortium [32] and the “Inflammation and Host Response to Injury” (Glue Grant) [30, 33] have catalogued the transcriptomic response in both health and disease in mice and humans. These human and murine datasets have been extensively evaluated by different groups, often leading to contradictory conclusions on whether the genomic response to inflammatory disease in mice recapitulates the human response to severe injury of infection [13, 14]. Although there is considerable controversy over the value of these murine models, there is general consensus that these animals models have some value but must be used with caution and an appropriate understanding of their limitations—this is the only way translatable progress in immunotherapy will occur in the future [34–36].

One fundamental criticism of existing murine models is that they fail to recapitulate the severity and multicompartmental nature of human trauma [16, 30, 31, 37]. Also, animal welfare issues appropriately limit our ability to completely imitate human trauma [38]. However, our laboratory has developed a murine model that better reflects the early inflammatory response of the severely injured patient while remaining within the guidelines of humane treatment of laboratory animals [31, 39, 40]. In an effort to better replicate human trauma in a murine model, we focused on combining insults that produce an Injury Severity Score greater than 15 [26]. The Injury Severity Score (ISS) is an anatomical scoring system that provides an overall score for patients with multiple injuries [41]. This value is intended to accurately represent the patient’s degree of critical illness. It has been considered the gold standard of classifying trauma patients early after their injury: Minor (ISS 1–3), Moderate (ISS 4–8), Serious (ISS 9–15), Severe (ISS 16–24), and Critical (ISS 25–75). An ISS greater than 15 is generally used as the minimum score for trauma studies of severely injured patients, as these individuals are more likely to have poor outcomes [1, 39, 41].

The polytrauma model described in this text utilizes the well-established trauma hemorrhage murine model of 90 min of hemorrhagic shock [42, 43]. In addition, a cecectomy, a long bone fracture, and muscle tissue damage are subsequently carried out in the mouse under anesthesia. The combination of these elements creates an insult which equates to an ISS of 18 in a human [39]. Although a further increased ISS would likely lead to a superior model that better represents human trauma, Institutional Animal Care and Use Committee’s (IACUC) limitations that rightly guard the humane treatment of animals typically will not allow for further insult to the rodents. Notwithstanding, in our experience, we have found that this model better reproduces the human immune response to injury and shock [31, 39].

Mice subjected to the polytrauma model below have an inflammatory response that better reflects the cytokine, chemokine, and leukocyte reaction seen after human trauma [39]. For example, we demonstrated that the plasma concentrations of interleukin-10 (IL-10), interferon-inducible protein-10 (IP-10), macrophage inflammatory protein-1α (MIP-1α), keratinocyte chemoattractant (KC), interleukin-6 (IL-6), and monocyte chemoattractant portein-1 (MCP-1) were significantly greater when compared to a traditional trauma hemorrhage model and more comparable to what is displayed in humans after trauma [39] (see Fig. 2). Furthermore, these animals had a leukocytosis associated with neutrophilia (see Fig. 3), had decreased major histocompatibility complex class II expression in bone marrow myeloid cells, and both of these effects were sustained beyond 1 day after the operation in this model; these phenomena were not displayed in the historical murine models of hemorrhagic shock and minor trauma [39]. In addition, this new model demonstrated improved correlations between the leukocyte gene expression patterns of severe human trauma patients, although species differences were still very evident [31]. However, when we evaluated the top responsive genes to trauma in humans, 88–99% of murine orthologues were identified to respond similarly to human transcriptomic alterations [31]. Many of these genes were involved in early inflammation, innate and adaptive immunity, indicating the polytrauma model’s usefulness as a method of studying inflammation after severe shock and injury [31]. We believe this model of hemorrhagic shock and polytrauma better recapitulates the human inflammatory response to severe injury and provides new insights in the pathophysiology of trauma while also offering a new approach to evaluating therapeutic interventions aimed at alleviating the short and long-term sequela of traumatic injury.

Fig. 2.

Plasma cytokine concentration in sham, trauma-hemorrhage (TH), and polytrauma (PT) mice. *p < 0.05, ^**p < 0.01 vs sham. Adapted from [39]

Fig. 3.

Blood neutrophils in sham, trauma-hemorrhage (TH), and polytrauma (PT) mice. ^**p < 0.01, ^***p < 0.001 vs sham. Adapted from [39]

2. Materials

2.1. Materials for Anesthesia and Preparation

• - Sterile gloves.

• - Weighing scale.

• - One-liter induction Plexiglas^® chamber (VetEquip, Livermore, CA).

• - Desiccant anesthetic scavenger canister (VetEquip, Livermore, CA) and tubing connectors.

• - Inhalant anesthesia system for veterinary surgery (VetEquip, Livermore, CA).

• - Universal rodent nosecone.

• - Isoflurane.

• - Oxygen tank.

• - Electric razor.

• - Petrolatum ophthalmic solution.

• - Platform for animal surgery (6 × 6 in. Plexiglas^® plates).

• - Fisher™ paper tape, 0.5 in.

• - Vis-U-All Self Seal sterilization pouch (Weck, A Squibb Company; Princeton, NJ).

2.2. Materials for Surgery

• - Mice (see Note 1).

• - Blood pressure analyzer (Micro-Med, Louisville, KY).

• - Dissecting microscope (Scienscope, Matthews, NC).

• - Bead sterilizer.

• - Infusion pump.

• - Polyethylene (PE)-10 catheter tubing (see Note 2).

• - Betadine scrubs.

• - Sterile water.

• - Lidocaine.

• - Heparin.

• - Normal saline.

• - Lactated Ringer’s (LR) solution.

• - Microforceps, angled, 3.5 in, 0.3 × 5 mm.

• - Nontoothed forceps.

• - Microdissection scissors, 4.5 in.

• - Microvessel clips, 0.75 × 4 mm.

• - Bulldog vessel clips.

• - Small scissors.

• - Small needle driver.

• - Small bone cutter.

• - 30 g, ½ in., hypodermic needles.

• - 1 mL syringe—slip tip.

• - 5–0 silk suture.

• - 2–0 silk suture.

• - 5–0 Ethilon suture.

• - 3–0 Vicryl suture.

• - BD Autoclip Closing System™.

• - Auto-clips, 7 mm.

• - Buprenorphine for injection (see Note 3).

• - Circulating water warming pad.

3. Methods

3.1. Induction and preparation

1. All instruments must be sterilized either by heat, ethylene oxide or other approved method. Merely cleaning, sanitizing or using antimicrobial washes are not adequate. In addition, all surgical materials such as sutures or sponges must be sterile and still within their expiration date. Solutions used for animals such as normal saline, Lactated Ringers (LR) solution or other drugs must be sterile and either the US Pharmacopeial Convention (USP) compliance standard or veterinary-grade (unless otherwise approved by the local IACUC).

2. Weigh the mouse (see Note 4).

3. Place the mouse in the induction chamber. Adjust the isoflurane concentration to 3.5–4.5% with O₂ flow at 1 L/min until an adequate plane of anesthesia is reached (see Note 5).

4. Shave the abdomen and inguinal area.

5. Apply USP grade petrolatum ophthalmic ointment to both eyes to prevent drying during the procedure.

6. Place the mouse on the Plexiglas^® plate or other nonabsorbent board in the supine position.

7. Place the anesthesia cone to the snout of mouse and maintain isoflurane concentration at 1.5–2.0% with 100% O₂ flow at0.5–1 L/min.

8. Restrain the mouse by taping all four extremities and its tail (see Note 6).

3.2. Femoral Artery Catheterization

1. Wipe the inguinal area with Betadine™ scrub and sterile water, alternating three times.

2. Place the mouse on a water-circulating, nonabsorbent warming pad under a dissecting microscope.

3. Visualize the femoral vessels through the skin, lift the overlying skin with forceps and excise a small section of skin to expose the femoral vessels using the microdissection scissors.

4. Carefully separate the femoral nerve and vein from the artery by blunt dissection using the microforceps.

5. Pass two 5–0 silk ties under the femoral artery spaced approximately 0.5 cm apart.

6. Ligate the femoral artery with the distal tie (5–0 silk) and attach a bulldog clip to the loose ends of the distal tie (see Note 7).

7. Tie the proximal tie (5–0 silk) loosely by creating a knot but not tightening it in order to keep the femoral artery lumen patent.

8. Place a vessel clip proximal to the proximal tie.

9. Apply a drop of 1% lidocaine to area (see Notes 8 and 9).

10. Puncture the artery just above the distal tie using a 30-gauge needle. The needle should have its bevel up such that the whole in the needle is facing the ventral or anterior portion of the mouse.

11. Gently insert the PE-10 catheter attached to a 1 mL syringe filled with LR solution, again with the bevel facing up (see Notes 10 and 11).

12. Remove the vessel clip from the femoral artery.

13. Gently advance the catheter using microforceps (see Image 1).

14. Secure the catheter by the proximal tie (5–0 silk) by tightening the knot made in step 7.

15. Another 5–0 silk tie may be added and tied to secure the catheter.

16. Tape the catheter to the Plexiglas^® plate to further secure it in place.

17. Repeat steps 3 through 16 on the contralateral side.

3.3. Hemorrhagic Shock and Resuscitation

1. Remove the mouse from the inhalational anesthesia and remove the 1 mL syringe from one of the catheters and attach this catheter to the pressure transducer line (see Notes 12 and 13).

2. Allow the mouse to emerge from anesthesia (at this point its mean arterial pressure (MAP) should be ~95 mmHg).

3. Reattach a new 1 mL syringe flushed with 1000 USP units/mL heparin to the contralateral catheter and slowly aspirate blood until the animal’s MAP is 30–40 mmHg (see Image 2).

4. Maintain the mouse in hemorrhagic shock for 90 min (see Notes 9 and 14).

5. Attach the catheter used for hemorrhagic shock to the infusion pump.

6. Resuscitate the mouse by administrating of LR solution. The volume of LR will be four times the volume of blood removed, and the infusion rate of the LR solution should be at a rate of 10 mL/h.

3.4. Decannulation and Bone Fracture

1. Disconnect the catheter connected to the blood pressure transducer and attach a 1 mL syringe filled with LR. Also disconnect the catheter connected to the infusion pump and attach a new 1 mL syringe filled with LR.

2. Reanesthetize the mouse. Place the anesthesia cone to the snout of mouse and maintain isoflurane concentration at1.5–2.0% with O₂ flow at 0.5 L/min. Again, ensure an adequate plane of anesthesia is reached (see Note 5).

3. Gently pull back the catheter until just past the proximal tie.

4. Ligate the artery by further tightening the proximal tie.

5. Completely remove the catheter and discard it in appropriate trash bag/container.

6. On one side, proceed to bluntly dissect the adjacent muscles to expose the long bone (tibia).

7. Fracture the bone using a bone cutter or small scissors (see Image 3).

8. Realign the bone (see Image 4).

9. Grasp the superior muscle tissue with a clamp for 30 s

10. Close the inguinal incisions with a running stitch or with a figure of eight stitch using a 5–0 Ethilon suture.

3.5. Cecectomy

1. Wipe the abdomen with Betadine™ scrub and sterile water, alternating the solutions three times.

2. Apply the sterile drape.

3. Make a 1 cm midline skin incision.

   1. Using either straight-edge or iris scissors, make a 1 cm midline cut into the skin only, approximately 0.5–1 cm away from xiphoid process.
   2. Entry into the peritoneum should be avoided.

4. Identify the abdominal wall and cut through the abdominal musculature and into the peritoneum (see Notes 15 and 16).

5. Identify and exteriorize the cecum using nontoothed forceps.

6. Doubly clamp the cecum about 1 cm from the tip (see Note 17) (see Image 5).

7. Ligate the stump with silk 2–0 using at least one square knot.

8. Excise the cecum (see Image 6).

9. Return the cecal stump to the abdominal cavity.

10. Close the abdominal wall using an absorbable suture (3–0 Vicryl) using a running suture.

11. Close the skin using surgical clips.

12. Inject the mouse subcutaneously with 0.05–1 mg/kg buprenorphine in 1 mL saline (see Note 18).

13. Return the animals to their cages after they awaken from anesthesia. This is usually indicated by their capacity to independently turn over onto their bellies after being placed on their sides or back. Keep the mouse on the warming pad until fully recovered.

14. Continue to monitor the mouse until it has fully recovered from anesthesia (see Note 19).

15. If additional mice are to be used, the instrument tips are to be washed in distilled water taking care to remove any blood or tissue. The instruments are then placed into a hot bead sterilizer for at least 10 min while the next animal is prepared. The sterile field should not be broken and the instruments are not to be handled without sterile gloves.