All procedures were performed under protocol #1588223, approved by the Veterans Affairs Puget Sound Health Care System Institutional Animal Care and Use Committee and in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

1. Animal care

NOTE: Animal models of LLB are limited solely by their availability and the capacity of the shocktube to accommodate their size. The described shocktube herein was designed specifically for use with mice.

1. Use 3-4-month-old male or female C57BL/6J mice or other approved mouse strains/lines in accordance with experimental needs. Maintain the mice on a 12 h dark-light cycle in specific pathogen-free facilities with ad libitum access to food and water. Mice are typically socially housed with 4 or 5 in a cage. Maintain facility temperatures at 20-22 °C.
2. Bring cages containing blast and sham mice to a nearby holding area. Bring separate empty cages for transferring individual mice to and from the blast room.

2. Shocktube preparation

1. (Safety check) Confirm that the necessary safety checks have been completed for the specific system. Ensure that the gas supply (helium) and master power are off/disconnected.
2. Prepare membranes as needed for the specific number of low-intensity blasts to be conducted (Figure 1.1). Cut membrane dimensions as required for the specific shocktube used in this protocol:
   1. Cut one sheet of plastic cling wrap into a 5.5" x 5.5" square to seal the spool, allowing it to pressurize.
   2. Cut one sheet of standard 8.5" x 11" copier paper (75 g/m² weight) to 5.5" x 11"; fold the resulting sheet of paper in half to form a 5.5" x 5.5" square.
   3. Obtain one sheet of 500 G mylar membrane (125 µm thickness).
      NOTE: These sheets are not ruptured or significantly deformed by standard low-intensity blasting and can be reused for the duration of a day's procedure.
3. Take a square of cling wrap and a square of folded paper and set them out on a flat surface (Figure 1.2). Place the folded paper on top of the cling wrap and align the two with each other as best as possible (Figure 1.3). To expedite repetitive blasts, arrange all the membrane stacks now.
4. Insert the mylar membrane between the driver and the spool by rolling it up into a small tube (about as big around as one's index finger; Figure 1.4,1.5). Insert it completely into the mechanism, and let go to allow it to unroll against the rubber seal that separates the driver section from the spool. Push the spool toward the driver to secure the mylar sheet in place; this will unseal the spool from the driven section of the shocktube.
5. Place the fingers under the top half of the cling wrap and carefully roll both the cling wrap and paper toward you, ensuring they roll up together without becoming misaligned (Figure 1.6).
6. Insert the membrane stack between the spool and the driven sections of the shocktube (Figure 1.7).
7. Allow the membrane stack to unroll so that the plastic seal is facing toward the spool and the paper is facing toward the driven section of the tube (Figure 1.8).
   NOTE: This orientation will create an airtight seal so that the system can be pressurized.
8. Close the spool assembly (Figure 1.9,1.10). As appropriate, tighten the bolts by hand or hydraulically, securing the driver-spool-shocktube assembly so that the system can be pressurized. (Safety check; Figure 1.10)
   NOTE: For hydraulic systems, ensure that the closure assembly's target pressure is reached to prevent misfires, which can require membrane replacement and slow the LLB exposure process. We use hydraulics to close our assembly at 500 psi.

3. Animal preparation

1. Turn on the circulating water heating pad below the anesthesia chamber, with the temperature set to 37 °C (Figure 1.11). Place an absorbent medical pad on top of the heat pad.
2. In the holding room, remove one mouse from its home cage and place it into an empty transfer cage. Bring the caged mouse into the blast room.
3. Turn the oxygen flow rate to 1.0 L/min (lpm) and the vacuum scavenge system on (Figure 1.12).
4. Turn the isoflurane on to 5% (to induce rapid unconsciousness) and route the flow to the rodent anesthesia chamber (Figure 1.13).
5. Place the mouse in the chamber to induce anesthesia (Figure 1.14).
6. Once the mouse is fully anesthetized and displays stable breathing for an additional 30 s, reach into the chamber and ear punch the mouse for unambiguous long-term identification of the mouse throughout the remainder of the study. Doing this step now is necessary to avoid interfering with recovery times after the blast. Then, apply sterile ophthalmic lubricant to both eyes to prevent corneal drying.
7. Remove the mouse from the chamber and place its nose into the nosecone (Figure 1.15). Switch the flow of anesthesia (e.g., isoflurane) from the induction chamber to the nosecone.
8. Use small pieces of laboratory tape to lightly restrain the mouse's limbs against the gurney (Figure 1.16).
9. After restraining the mouse, place a wire twist tie around each limb and twist tightly, securing the mouse to the gurney at the wrists and ankles (Figure 1.17). Place a larger twist tie around the chest, tying it very loosely such that the mouse's breathing is not restricted. This will serve as a secondary restraint mechanism in case any of the limb restraints come loose.
10. Lift the mouse's tail and place it under the left foot to ensure it does not get pinched when the gurney is inserted into the shocktube (Figure 1.18).

4. LLB procedure

1. Open the animal exposure section of the shocktube and orient the mouse so that it faces the oncoming blast wave (Figure 1.19).
2. Secure/suspend the gurney in the animal exposure section (Figure 1.20).
3. Close the door tightly for the animal exposure section, ensuring that the anesthetic flow tube is not pinched by the door (Figure 1.21).
4. Reduce the anesthesia to 2.5-3% isoflurane, 1 lpm for the remainder of the session.
5. Power the system as appropriate (Figure 1.22).
6. Locate and connect the supply line for the compressed helium gas (Figure 1.23,1.24).
7. Leave the blast room to access the blast tube control console in an adjoining room, and ensure no personnel or animals are left in the blast room.
   NOTE: Hearing protection may be required by the institution or by operational conditions. Such conditions may include shocktube arrangements where the control console is located in the same open space as the shocktube.
8. From the console, turn on the acquisition software to record the blast event (see the green box in Figure 1.25).
   NOTE: For these procedures, we collect sensor data at a 20 kilo hertz (kHz) sampling rate, which is then processed using LabView software. We recommend acquiring sensor sampling at ≥10 kHz to achieve high-quality time versus pressure curves.
9. Disengage any safety lock (e.g., power control keys, which are depicted by a green arrow in Figure 1.26).
10. Close both gas vents and passively pressurize the spool (Figure 1.27). Do not use the driver side. Continue to fill until the membrane ruptures on its own at the target peak psi as determined by the number of membrane sheets used.
11. Record the peak pressure, positive phase duration, and impulse at the animal location. (Figure 1.28). Turn off the fill mechanism.
12. Return to the shocktube, disconnect the helium feed line, and turn off the power supply to the blast control circuit (Figure 1.29).
13. To conduct repeated LLB exposures on the same animal, open the spool, remove the spool membrane stack, and then roll and insert another spool membrane stack (Figure 1.30, 1.31, 1.32). Flatten the membrane stack and reclose the assembly.
    NOTE: To model the clinical experience of low-level blast exposures during empirically defined SOF training, we expose mice to 5-6 LLBs per day, capping daily exposures to a conservative ~20 cumulative total psi⁴⁵. Studies emphasizing mechanistic and dose-response relationships may alternatively choose to use a consistent number of LLB exposures with defined overpressure parameters per session.
14. After the final LLB for the current animal, remove it from the shocktube, leaving the anesthesia on (Figure 1.33).
15. Untie the animal while it is under anesthesia. Remove it from the anesthesia nosecone, placing it on its back onto the heated water pad (Figure 1.34).
16. Once the animal has been placed on the water pad, start a timer and record the amount of time until the mouse flips over onto its ventral side (i.e., its stomach) on its own (Figure 1.35). Record this time as the righting time. Once the mouse recovers, return it to the home cage and continue monitoring as needed.

5. Multiday procedures

1. To model routine LLB exposures from breaching charges used during SOF Close Quarter Battle training, perform repeated daily exposures on the mice 5 days a week (Monday through Friday) for a total of 15 days across 3 standard work weeks.

6. Altering peak LLB pressures

1. Increase peak pressure through the use of stronger membrane materials or by simply stacking additional membranes. For example, use Mylar Roll Clear 0.005 (500 G) membrane to produce ~20 psi peak pressure (when used as both driver and spool membranes) or Mylar Roll Clear 0.002 (200 G) membrane to produce ~10 psi peak pressure.
2. Adjust the parameters for the positive phase duration and impulse of the blast to meet experimental needs. To adjust positive phase durations and impulses, empirically determine target conditions by substituting compressed gas sources^47,49 or changing the driver length whenever possible. The above protocol uses helium to create a sharp peak pressure and waveform similar to an idealized Friedlander curve.

7. Tissue collection

NOTE: Tissue collection practices can be adjusted according to experimental needs.

1. Anesthetize the mouse via intraperitoneal injection with 210 mg/kg of pentobarbital.
2. Place the mouse into a mouse or rat cage with bars or a premade mesh; place the caged mouse into a fume hood.
3. Once the mouse is unresponsive, place it on its back on the bars on top of the cage and close its mouth around one of the bars to help it stay in place during perfusion.
4. Grab the skin of the stomach, pull it upwards, and use a pair of large scissors to cut a hole in the abdominal cavity, being careful not to cut any of the organs. Continue to cut farther down along the base of the ribs to allow for freer articulation of the ribcage.
5. Using a hemostat, approach the mouse from the side and grab the tissue directly on top of the ribcage, rolling the hemostat back to keep the base of the ribcage angled in an easily accessible position. Use a pair of forceps or a similar tool to hold the hemostat in place.
6. Using a small pair of surgical scissors, carefully cut the diaphragm to allow access to the heart. Use a pair of forceps to gently angle the heart such that the bottom is facing directly out of the open base of the ribcage. Work quickly so the heart will still be beating during perfusion.
7. If collecting blood, hold the heart with a pair of forceps and carefully pierce the right ventricle using a 3 mL syringe tipped with an 0.5" 25 G needle. Insert from the bottom of the ventricle and go in lengthwise, being careful not to pierce the opposite side of the ventricle. Gently pull on the syringe until 0.5-1.0 mL of blood has been collected or the flow stops, and then remove the syringe.
8. Use a pair of surgical scissors to cut a small incision in the right atrium to allow blood and perfusate to drain. Hold the heart with a pair of forceps, and carefully insert a 25 G butterfly needle into the left ventricle, inserting from the bottom. Hold the butterfly needle in place with a holding clamp or by hand.
9. Perfuse the animal.
   1. Connect a syringe containing 50 mL of phosphate-buffered saline (PBS) to a butterfly needle and perfuse at a rate of approximately 10 mL/min. Look for blanching of the liver as a sign of proper perfusion. After the syringe is emptied, disconnect it from the butterfly needle.
   2. For preparation of tissues for microscopy, replace the empty PBS syringe with a syringe containing 50 mL of 10% neutral buffered formalin (NBF) or 4% formaldehyde solution. Repeat the above steps to perfuse with formalin.
      NOTE: The perfused mouse should be observed to twitch during perfusion; this should result in full body rigor or rigidity after the procedure is completed.
10. Remove the butterfly needle from the heart and remove the mouse from the cage bars for tissue collection.
11. Remove and subdissect the target organs according to need; be careful to perform procedures on ice when fresh, unfixed materials are collected.
12. Flash freeze any unfixed tissues that were collected in liquid nitrogen and store them at -80 °C until used in protocols assaying protein or RNA targets.
13. For fixed tissues, remove to a labeled 50 mL conical tube filled with formalin (one tube per organ).