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Gastric Aspiration Models
胃内容物的气管吸入模型   

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Abstract

The procedures described below are for producing gastric aspiration pneumonitis in mice with alterations for rats and rabbits described parenthetically. We use 4 different injury vehicles delivered intratracheally to investigate the inflammatory responses to gastric aspiration:
1) Normal saline (NS) as the injury vehicle control
2) NS + HCl, pH = 1.25 (acid)
3) NS + gastric particles, pH ≈ 5.3 (part.)
4) NS + gastric particles + HCl, pH = 1.25 (acid + part.)
The volume, pH, and gastric particle concentration all affect the resulting lung injury. In mice, we generally use an injury volume of 3.6 ml/kg (rat: 1.2 ml/kg, rabbit: 2.4 ml/kg), an injury pH (for the acid-containing vehicles) of 1.25, and a gastric particulate concentration (in the particulate-containing vehicles) of 10 mg/ml (rat: 40 mg/ml). In our hands this results in a maximal, non-lethal lung injury with ≤ 10% mortality for the most injurious vehicle (i.e., acid + part.) The maximum tolerable particulate concentration needs to be determined empirically for any new strains to be used, especially in genetically-altered mice, because an altered inflammatory response may have detrimental affects on mortality.
We have extensive experience utilizing these procedures in the outbred strain, CD-1, as well as many genetically-altered inbred stains on the C57BL/6 background. Choice of strain should be carefully considered, especially in terms of strain-specific immune bias, to assure proper data interpretation. The size of the mouse should be ≥ 20 g at the time of injury. Smaller mice can be attempted, if necessary, but the surgical manipulation becomes increasingly more difficult and the surgery survival rate decreases substantially. There are no size or strain constraints for rat and rabbit models, but we generally use Long-Evans rats at 250-300 g and New Zealand White rats at ≈ 2 kg at the time of initial injury.

Keywords: Pneumonitis(肺炎), Acute lung injury (ALI)(急性肺损伤(阿里)), Acute respiratory distress syndrome (ARDS)(急性呼吸窘迫综合征(ARDS)), Inflammation(炎症), Rodent models(啮齿类动物模型)

Materials and Reagents

  1. Isoflurane
  2. Topical antiseptic microbicide prep solution (e.g. Medline Industries, catalog number: MDS093906 )
  3. 0.5% bupivicaine
  4. Hank’s Balanced Salt Solution (HBSS) with Ca2+, Mg2+ (e.g. Life Technologies, catalog number: 14025 )
  5. HBSS without Ca2+, Mg2+ (e.g. Life Technologies, catalog number: 14175 )
  6. Liquid nitrogen
  7. Bovine serum albumin (BSA)
  8. Cytospin filter cards (e.g. Thermo Fisher Scientific, catalog number: 5991022 )
  9. Diff Quik solutions kit (Fisher Scientific, catalog number: NC9943455 )
  10. Cytoseal 60 (VWR International, catalog number: 48212-154 )
  11. 100x protease inhibitor cocktail (e.g. Calbiochem®, catalog number: 80053-844 )
  12. Bupivacaine
  13. 100 mg/ml mouse (rat) gastric particles (see Recipes)
  14. Acid injury solution (acid) (see Recipes)
  15. Gastric particles injury solution (part.) (see Recipes)
  16. Acid + particles injury solution (acid + part.) (see Recipes)
  17. Phosphate buffered saline (PBS), pH 7.2 (see Recipes)
  18. Ammonium chloride lysis buffer (see Recipes)
  19. Lung homogenate buffer (see Recipes)
  20. 50 mM potassium phosphate buffer, pH = 6.0 (see Recipes)
  21. MPO homogenate buffer (see Recipes)

Note: All salts and other chemicals are from Sigma-Aldrich unless otherwise noted (however, any source for such chemicals is probably okay to use).

Equipment

  1. 1-0 braided silk material, bulk spool (e.g. Look catalog number: MBJF210)
  2. 12 mm x 75 mm x 1 mm microscope slides
  3. 2 x 2 gauze sponges, 8-ply (VWR International, catalog number: 82004-740 )
  4. Sterile 4 x 4 gauze sponges, 8-ply (VWR International, catalog number: 82004-742 )
  5. 6-0 monofilament polypropylene suture with P-13 cutting needle (e.g. Syneture, catalog number: SP-5695 )
  6. 60° incline dissection board (homemade out of plexiglass)
  7. Syringes (0.5, 1, 3, 5, 20 cc (cubic centimeter))
  8. Needles (14, 20, 22, 26, 29 gauge)
  9. Tracheal cannula (23 gauge x 1/2” stainless steel tubing adapter) (Becton, Dickinson and Company, catalog number: 408213 )
  10. 3” curved serrated forceps (2)
  11. 3” curved tissue (“toothed”) forceps
  12. 1.8 ml microfuge tubes
  13. 12 x 75 mm polystyrene tubes
  14. 22 x 22 mm #1.5 coverslips
  15. 4” curved micro dissecting scissors
  16. 37 °C water bath
  17. Disposable skin stapler (e.g. 3M, model: DS-25 )
  18. Hemocytometer or Coulter counter (e.g. Beckman Coulter, model: MultiSizer III )
  19. Cytocentrifuge with cytospin funnels (e.g. Shandon CytoSpin®)
  20. Tissue homogenizer (e.g. BrinkmannTMPolytronTM, model: PT2000 )
  21. Probe sonicator (e.g. Branson Sonifier®, model: 450 )

Procedure

  1. "Injury" procedure
    1. Fill a 0.5 cc syringe with 22 gauge needle with 0.2 ml air “chaser” then “injury” solution (rat: 1 cc syringe with 14 gauge needle with 0.5 ml air, rabbit: 5 cc syringe with 14 gauge needle with 2 ml air).
    2. Induce anesthesia in a chamber with 3-4% isoflurane in oxygen delivered at 1-2 L/min (rabbit: 30 mg/kg ketamine intramuscularly prior to 2% isoflurane) (Figures 1 and 2). Suspend mouse by its front teeth with a 1-0 suture strand on a 60° incline dissection board and maintain anesthesia with 2-2.5% isoflurane administration with nose cone.


      Figure 1. Lab bench set-up for surgical procedures. Procedures are performed in a fume hood with charcoal filtering for anesthetic gas scavenging that contains: small animal anesthetic gas exposure chamber (front right), lamps, and the plexiglass 60° incline dissection board (middle of hood). On the left of the picture, on a floor stand outside the hood, is the anesthetic vaporizer for delivering the isoflurane vapor.


      Figure 2. Instruments used for aspiration injury procedure. Plexiglass 60° incline dissection board is shown with nose cone setup to deliver isoflurane during the procedure. The green hose delivers the isoflurane in 100% oxygen to an inner nose cone, whereas the larger bore blue hose provides vacuum for scavenging the anesthetic through the outer nose cone. Also shown are (left to right): a 5 cc syringe with 26 gauge needle containing 1.5 ml normal saline for the subcutaneous fluid injection, a 0.5 cc syringe with 22 gauge needle containing the injury solution and 0.2 ml air “chaser”, 3 curved dissecting forceps (2 serrated and 1 “toothed”), curved dissecting scissors, a vial of antiseptic antimicrobial solution with cotton-tipped applicator, and a 6” strand of 1-0 braided silk suture material. Missing from picture: disposable skin stapler.

    3. Shave ventral neck, paint with topical antiseptic prep solution and remove excess with gauze.
    4. Infiltrate future incision site with 100 μl 0.5% bupivacaine (provides local post-operative analgesia).
    5. Cut a 2 cm longitudinal incision in the skin with scissors (Figure 3).


      Figure 3. Neck incision. After suspending the mouse by its front incisors with a suture such that its nose is within the inner isoflurane nose cone (step A2), prepping the surgical site (steps A3 & A4), a 2 cm longitudinal incision is made with scissors and tissue forceps (step A5).

    6. Expose trachea by blunt dissection with 2 curved toothless forceps (Figures 4, 5 and 6).


      Figure 4. Tracheal exposure by blunt dissection. The fascia membrane is teased away with curved serrated forceps and the salivary glands are pulled to the side in order to expose the trachea (faint white vertical between the forceps) that is surrounded by paratracheal musculature (step A6).


      Figure 5. Tracheal exposure by blunt dissection, continued. Grasp one side of the paratracheal musculature and pull it to the side while rubbing along the muscle longitudinally. This will result in a separation of the muscle fiber bundles and allow the muscle to be reflected to the side (step A6). Notice the inner and outer nose cone configuration for delivering the anesthetic and scavenging. Mouse’s nose has been positioned lower than usual for clarity.


      Figure 6. Tracheal exposure by blunt dissection, continued. Use the two curved serrated forceps to reflect the paratracheal musculature, fully exposing the trachea (step A6).

    7. Work curved forceps under trachea (Figure 7). and use to pull a 6" strand of 1-0 silk suture material through (Figure 8).


      Figure 7. Work curved serrated forceps under trachea (step A7)


      Figure 8. Use forceps to place suture under trachea (step A7)

    8. Discontinue isoflurane administration by removing nose cone. Before instilling injury vehicle by the steps below, allow mouse’s plane of anesthesia to rise until it just starts reacting to a forceps toe pinch then instigate instillation steps quickly. Especially for the acid-containing injury vehicles, if the plane of anesthesia is too deep the animal will not begin to breathe spontaneously after the injury vehicle is instilled.
    9. Lift trachea up with suture to facilitate inserting injury syringe needle into the trachea (2-3 cartilage rings below the larynx) with bevel facing surgeon (advance until bevel of needle is just past the trachea insertion point) (Figure 9). It is important that the needle be as parallel with the trachea as possible, otherwise, the needle can easily pierce the other side of the trachea. This will result in the injury vehicle not being injected into the lungs and a high probability that the animal will not survive.


      Figure 9. Use suture to facilitate injecting injury solution into trachea. Remove nose cone apparatus from incline board to provide unrestricted access to upper section of trachea. Lift trachea up with suture to facilitate inserting injury syringe needle into the trachea (2-3 cartilage rings below the larynx). Notice bevel of needle facing surgeon. Advance needle until bevel of needle is just past the trachea insertion point (step A9). It is important that the needle be as parallel with the trachea as possible, otherwise, the needle can easily pierce the other side of the trachea. This will result in the injury vehicle not being injected into the lungs and a high probability that the animal will not survive. While assistant squeezes rib cage (to expel most of vital capacity), quickly instill syringe contents. Release chest just before the injection begins. This maneuver, along with the air “chaser” assures deposition of the injury vehicle into the distal lung (step A10).

    10. While assistant squeezes rib cage (to expel most of vital capacity), quickly instill syringe contents. Release chest just before the injection begins. This maneuver, along with the air “chaser” assures deposition of the injury vehicle into the distal lung.
    11. Leave animal on incline board until breathing commences and close incision with 2 staples (for rats and rabbits: trachea needle wound needs to be repaired with one 6-0 suture) (Figure 10).


      Figure 10. Close neck skin incision with surgical staples (step A11)

    12. Inject 1 ml (rat: 10 ml, rabbit: 20 ml) sterile NS subcutaneously into the scruff of the neck for fluid resuscitation. There is virtually no fluid loss during the procedure, but the animal will not drink for a while after the procedure and can dehydrate (Figure 11).


      Figure 11. Fluid resuscitation. Inject 1 ml sterile NS subcutaneously into the scruff of the neck using a syringe with 26 gauge needle for fluid resuscitation (step A12).

    13. Put into heated chamber (37 °C) perfused with 100% O2 until ambulatory (Figure 12).


      Figure 12. Recovery chamber. Place mouse in recovery chamber that is continually perfused with 100% O2 (supplied by green hose) and maintained at 37 °C with heat lamp and temperature controller (notice thermistor probe in left chamber) until the mouse is ambulatory (step A13).

  2. Harvest procedure (Figure 13)


    Figure 13. Instruments and materials used for harvest. From left to right: two serrated dissecting forceps, tissue (“toothed”) forceps, dissecting scissors, 23 gauge stainless steel blue-hubbed cannula, 1 cc blood collection syringe with 26 gauge needle, bronchoalveolar lavage apparatus (top syringe filled with 5 ml HBSS without Ca2+, Mg2+, left syringe is for fluid collection), two 2 x 2 gauze sponges (bottom left) for retracting abdominal organs, and 4” strand of 1-0 braided silk suture (bottom, middle). Missing from picture: 5 cc syringe with 26 gauge needle containing 5 ml HBSS with Ca2+, Mg2+ for flushing pulmonary vasculature.

    1. Anesthetize mouse with isoflurane in 100% O2.
    2. When mouse is unresponsive, place on a dissecting board in supine position and continue isoflurane administration with a nose cone.
    3. Using forceps and scissors, make a longitudinal incision up the abdomen to the xyphoid.
    4. Make a latitudinal incision across the belly (Figure 14) and expose vena cava and abdominal aorta using gauze sponges to reflect abdominal organs (Figure 15).


      Figure 14. Abdominal incision for collecting blood. Using forceps and scissors, make a longitudinal incision up the abdomen to the xyphoid and a latitudinal incision across the belly (steps B3 & B4).


      Figure 15. Collect blood. Expose vena cava and abdominal aorta using gauze sponges to reflect abdominal organs and collect blood using a 1 cc syringe with 26 gauge needle from either vessel (steps B4 & B5).

    5. Harvest blood by (used to assess systemic inflammatory responses (e.g. serum or plasma cytokine levels)):
      1. Collect blood from abdominal aorta using 27 gauge needle on a 1 cc syringe (if plasma is to be isolated syringe must contain appropriate anticoagulant) (Figure 15).
      2. The vena cava can be used to collect blood, but generally, not as much blood can be collected as from the aorta (Figure 15).
      3. Dispense blood into 1.8 ml μ-fuge tube. Until blood can be processed, keep on ice if plasma will be prepared, or keep at room temperature if serum will be prepared. Process blood, as described below.
      4. Transect vena cava/abdominal aorta to euthanize by exsanquination.
    6. Flush pulmonary vasculature by (performed to remove the blood from the pulmonary circulation so measurements of compounds in the processed lung tissue reflect only values from the pulmonary compartment that do not include a systemic contribution):
      1. Grasp xyphoid process with forceps, puncture diaphragm with tips of scissors to deflate lungs and carefully cut away ventral aspect of diaphragm (Figure 16).


        Figure 16. Cut-away diaphragm. Grasp xyphoid process with forceps, puncture diaphragm with tips of scissors to deflate lungs and carefully cut away ventral aspect of diaphragm (step B6a).

      2. Continue the longitudinal incision up the sternum and through the neck.
      3. Cut away sternum to expose lungs (be careful not to puncture lungs) (Figure 17).


        Figure 17. Perform sternotomy. Continue the longitudinal incision up the sternum and through the neck. Cut away sternum to expose lungs (be careful not to puncture lungs) (steps B6b-c).

      4. Grasp apex of heart with toothless forceps and inject 5 ml (rat: 20 ml) 1x HBSS with Ca2+, Mg2+ (at 37 °C) into right ventricle (it will be on the left side of the heart since the animal is on its back) with 5 cc (rat: 20 cc) syringe + 26 gauge needle (Figure 18). The rate of injection should be fast enough to keep the heart “inflated”, but not so fast as to force the fluid out through the injection site. The calcium in the buffer, and its being at 37 °C, aids the heart to keep beating to facilitate flushing the pulmonary vasculature.


        Figure 18. Flush pulmonary vasculature. Grasp apex of heart with serrated forceps (tissue forceps have a tendency to tear the heart tissue) and inject 5 ml 1x HBSS with Ca2+, Mg2+ (at 37 °C) into right ventricle (it is on the left side of the heart in this picture since the animal is on its back) with 5 cc syringe + 26 gauge needle. The rate of injection should be fast enough to keep the heart “inflated”, but not so fast as to force the fluid out through the injection site (step B6d).

    7. Perform bronchoalveolar lavage (BAL) by (performed to collect cells and compounds secreted into the airspaces (i.e. alveoli and bronchial lumen) that can be assessed to determine the degree of pulmonary inflammation (i.e. neutrophil influx, various inflammatory cytokine levels (i.e. tumor necrosis factor-alpha (TNFα), interleukin-1beta (IL-1β), IL-6, macrophage inflammatory protein-2 (MIP-2), monocytic chemotactic protein-1 (MCP-1)), and lung injury (i.e. total protein or albumin concentration):
      1. Expose trachea and work tip of curved forceps under it to pull a 4" strand of 1-0 silk suture through.
      2. Insert a 23 gauge x 1/2” (rat: 14 gauge, rabbit: 3 mm ID, 4.5 mm OD) stainless steel cannula (rabbit: polyethylene catheter) into the trachea and secure with suture (Figures 19 & 20).


        Figure 19. Insert tracheal cannula. Expose trachea and work tip of curved forceps under it to pull a 4" strand of 1-0 braided silk through, as described in the injury procedure (steps A6, A7 & A9). Use a 20 gauge needle to make a hole in the upper trachea and insert a 23 gauge 1/2” stainless steel cannula into the hole.


        Figure 20. Secure tracheal cannula. Secure cannula in the trachea with the suture (steps B7a-b). Be sure the suture is tight and 2-3 mm below the insertion site so a good seal is made. Tip of cannula should be at least 3 mm before the carina (bifurcation of the trachea). If the 1/2” cannula is inserted just below the larynx, as depicted in the figure, the tip of the cannula will be in the proper position.

      3. Connect lavage apparatus (2, 5 cc syringes connected to a 4-way stopcock) to catheter and slowly, lavage lungs with 5 x 1 ml (rat: 10 ml, rabbit: 50 ml) 1x HBSS without Ca2+, Mg2+ (at 37 °C) (Figures 21 & 22). The lack of calcium in the buffer facilitates harvesting of the airway cells by diminishing adherence.


        Figure 21. Perform bronchoalveolar lavage (BAL)-injection. Connect lavage apparatus to cannula and slowly inject 1 ml of 1x HBSS with Ca2+, Mg2+ to insulflate lungs (step B7c).


        Figure 22. Perform BAL-collection. Switch stopcock valve and slowly collect the injected lavage fluid. Repeat the injection and collection process for a total of 5 lung washings (step B7c).

      4. Dispense collected BAL fluid (BALF) into appropriately sized tube, record recovered volume (by weighing), and put on ice until BALF processing, described below.
    8. Harvest lungs by (performed to assess the pulmonary inflammatory state distinct from the airspaces, i.e., parenchymal and interstitial tissue):
      1. Excise lungs, heart, and thymus, en bloc, by the trachea (Figure 23).


        Figure 23. Harvest lungs, heart, and thymus, en bloc. Be sure clavicle has been cut away. Grasp cannula, trachea, and suture with fingers and cut trachea by larynx. While pulling up and out, work scissors dorsally under lungs and spread to disengage tissue connecting lungs and heart to thoracic cavity. Some cutting of connective tissue will be needed, as well as the esophagus and vessels to successfully remove the lungs, heart, and thymus, en bloc (step B8a).

      2. Remove lung lobes by cutting lobe’s main stem bronchus at hilum (Figures 24 & 25), place in 2 ml cryovial, and flash freeze in liquid N2.


        Figure 24. Harvest lung lobes. Collect individual lobes by using scissors and forceps to cut main stem bronchi at the lobe’s hilum (step B8b).


        Figure 25. Mouse lung lobes. Clockwise from left: left lung lobe, right lung-cranial lobe, right lung-middle lobe, right lung caudal lobe, right lung-post-caval lobe.

      3. Store at -80 °C until appropriate processing can be performed, described below.

  3. Blood processing
    1. Plasma
      1. Centrifuge at 2,000 x g for 4 min at 4 °C.
      2. Remove plasma with a pipet and dispense into 1.8 ml microfuge tube.
      3. Store at -80 °C.
    2. Serum
      1. Incubate at RT for 30 min then overnight at 4 °C to allow clot to contract.
      2. Centrifuge at 5,000 x g for 15 min at 4 °C.
      3. Remove serum with a pipet and dispense into 1.8 ml microfuge tube.
      4. Store at -80 °C.

  4. BALF processing (keep samples on ice)
    1. Spin BALF at 1,500 x g for 5 min at 4 °C.
    2. Without disturbing pellet, collect supernatant with pipet (leave 50-100 μl residual volume of uncollected supernatant to prevent sucking-up cells) and dispense into equal volume aliquots in 1.8 ml microfuge tubes (the number and volume of the aliquots will depend on the assays that will be performed and should be designed to limit the amount of freeze/thaw cycles). Store at -80 °C.
    3. Resuspend pellet in the residual volume.
    4. Add 1 ml ammonium chloride lysis buffer (37 °C) and incubate for 2 min to lyse red blood cells.
    5. Layer entire volume on 4 ml ice cold 2% BSA in PBS in a 12 x 75 mm PS tube.
    6. Spin at 150 x g for 15 min at 4 °C.
    7. Aspirate supernatant (containing cell debris and lysed red blood cells), resuspend pellet in residual volume, and add 1 ml PBS. Removing the cell debris and lysed red blood cells by centrifuging the cells through 2% BSA makes counting the cells much easier and more accurate.
    8. Count cells on hemocytometer or Coulter counter.
    9. Prepare cytospin slide for microscopic viewing by:
      1. Add 5 x 104 white blood cells to 2% BSA in PBS in a cytospin funnel (300 μl total volume) attached to a microscope slide and filter card and spin at 28 x g (500 rpm) for 5 min.
      2. Remove slides and allow to air dry (IMPORTANT: stain immediately when dry).
      3. Stain with Diff Quik by:
        1. Dip slides 5 x 1 sec in fixative, Solution I, Solution II (Diff-Quik kit), rinse with H2OMQ & air dry.
        2. Mount slide with small drop of "Cytoseal 60" on cell spot and putting a 22 x 22 mm #1.5 coverslip on it (be sure no bubbles) and allow to air dry.
        3. Determine percentage of macrophages, neutrophils, lymphocytes, and eosinophils by light microscopy.

  5.  Lung processing (Lungs can be processed in a number of different ways depending on what cellular compartment (e.g. whole lung homogenate, nuclear fraction, cytosolic fraction, etc.) and molecular species (e.g. protein, mRNA, etc.) are to be assayed and what assay techniques will be utilized (e.g. ELISA, Western blot, PCR, etc.). This procedure produces a lung homogenate supernatant that can be assayed for various cytokine protein levels by ELISA, as well as an extraction of myeloperoxidase from the lung homogenate pellet that can be assayed for enzymatic activity as a surrogate marker of neutrophil infiltration)
    1. Thaw frozen lungs and transfer to a round bottom centrifuge capable of withstanding 40,000 x g. Weigh tube before adding lungs then weigh the tube + lungs.
    2. Add enough cytokine homogenate buffer to bring the weight of the lungs + buffer to 3 g (rat: 10 g).
    3. Homogenize tissue with Polytron homogenizer with tube on ice to prevent heating of sample. Pick out any tissue caught in homogenizer blades with forceps. Rinse homogenizer with 70% EtOH and then sterile H2O between samples.
    4. Centrifuge homogenate at 40,000 x g for 10 min at 4 °C.
    5. Dispense supernatant into equal volume aliquots in 1.8 ml microfuge tubes and store at -80 °C.
    6. Add 2 ml MPO buffer to pellet and resuspend by vortexing.
    7. Store at -80 °C (in same tube) until MPO extraction can be performed.

  6. MPO extraction procedure
    1. Quick thaw sample in 37 °C water bath.
    2. Sonicate for 1 min, on ice to prevent sample heating, on maximum output at 50% duty cycle.
    3. Incubate at 55 °C for 2 h.
    4. Centrifuge at 40,000 x g for 15 min at 4 °C.
    5. Dispense into 0.5 ml aliquots in 1.8 ml microfuge tubes and store at -80 °C until MPO activity can be assessed

Recipes

  1. 100 mg/ml mouse (rat) gastric particles
    1. Harvest stomach from freshly necropsied mouse (rat) first thing in the morning and put in 50 ml tube on ice. Harvesting early in the morning assures a full stomach.
    2. In laminar flow hood
      1. Put stomach in Petri dish, cut longitudinally with scissors.
      2. Place stomach contents in a clean, 50 ml centrifuge tube.
      3. Add 10 ml sterile normal saline (NS) and vortex vigorously for 15 sec.
      4. Pour particle suspension through 8 layers of sterile gauze into a beaker.
      5. Use an additional 10 ml sterile NS to transfer residual particles in tube to gauze.
      6. Wring gauze to collect absorbed fluid.
      7. Transfer filtrate to a clean, 50 ml centrifuge tube.
    3. Spin filtrate at 5,000 x g for 5 min at 4 °C.
    4. Discard supernatant, and resuspend in 25 ml, sterile NS.
    5. Repeat steps 1c & 1d.
    6. Transfer particle suspension to an Erlenmeyer flask and autoclave 121 °C, 121 psi for 30 min (use an additional 25 ml, sterile NS to transfer residual particles).
    7. Transfer cooled, sterile particle suspension to a pre-weighed 50 ml centrifuge tube.
    8. Spin filtrate at 5,000 x g for 5 min at 4 °C.
    9. Decant supernatant and stand tube inverted on KIMWipe to remove all liquid.
    10. Re-weigh tube+particle pellet to determine particle wet weight.
    11. Add sterile NS to 100 mg/ml (particle wet weight/total volume).
    12. Store at 4 °C for up to 3 weeks until used for injury.
  2. Acid injury solution (acid)
    Normal saline (NS), sterile
    9.44 ml
    1 N HCl, sterile  
    562 μl
    Adjust pH = 1.25 with sterile 1 N HCl
    Make fresh
  3. Gastric particles injury solution (part.)
    Normal saline (NS), sterile
    9 ml
    100 mg/ml gastric particles in NS
    1 ml (10 mg/ml)
    Make fresh.
  4. Acid + particles injury solution (acid + part.)
    Normal saline (NS), sterile
    8.44 ml
    1 N HCl, sterile
    562 μl
    100 mg/ml gastric particles in NS
    1 ml (10 mg/ml)
    Adjust pH = 1.25 with sterile 1 N HCl
    Make fresh
  5. Phosphate buffered saline (PBS), pH 7.2
    NaCl (58.44)
    8 g (137 mM)
     
    Na2HPO4 (141.96)
    1.15 g (8.1 mM)

    KCl (74.56)
    200 mg (2.7 mM)

    KH2PO4 (136.09) 
    200 mg (1.5 mM)

    H2OMQ to 1 L
    Adjust pH = 7.2, filter sterilize
    Stored at room temperature
  6. Ammonium chloride lysis buffer
    NH4Cl (53.49)
    4.13 g (154 mM)

    KHCO3 (100.1)
    500 mg (10 mM)

    EDTA, tetrasodium salt (380.2)
    18.5 mg (0.1 mM)

    Mix powders together, thoroughly
    Dispense equal amounts into 10, 50 ml centrifuge tubes (0.46 g/tube)
    On day of use, add 50 ml H2OMQ and mix until dissolved
    Discard any unused solution
  7. Lung homogenate buffer
    NaCl (58.44)
    8.77 g (150 mM)
    Tris base (121.14)
    1.82 g (15 mM)
    CaCl2.2H2O (147.02)
    147 mg (1 mM)
    MgCl2.6H2O (203.3)
    203 mg (1 mM)
    H2OMQ to 1 L
    Adjust pH = 7.4, autoclave at 121 °C, 15 psi, for 15 min
    Stored at 4 °C
    At time of use, add 1/100th volume of 100x protease inhibitor cocktail to volume of lung homogenate buffer needed for the day’s processing
  8. 50 mM potassium phosphate buffer, pH = 6.0
    KH2PO4 (136.01)
      3.4 g (50 mM) 

    H2OMQ to 500 ml
    Adjust pH = 6.0 with 2 M NaOH
    Stored at room temperature
  9. MPO homogenate buffer
    Hexadecyltrimethylammonium bromide (364.5) 
     2 g (0.5%)

    EDTA (372.24)  
     744 mg (5 mM)

    50 mM potassium phosphate buffer, pH = 6.0
     400 ml

     Stored at room temperature (Do not refrigerate)

Acknowledgments

The work presented here was supported by NIH grant HL048889, “Pathogenesis of Aspiration Pneumonitis” to Paul R Knight, M.D., Ph.D. and Bruce A. Davidson, Ph.D. To cite this protocol please also use the following reference: Davidson et al. (2013).

References

  Mouse models

  1. Davidson, B. A., Vethanayagam, R. R., Grimm, M. J., Mullan, B. A., Raghavendran, K., Blackwell, T. S., Freeman, M. L., Ayyasamy, V., Singh, K. K., Sporn, M. B., Itagaki, K., Hauser, C. J., Knight, P. R. and Segal, B. H. (2013). NADPH oxidase and Nrf2 regulate gastric aspiration-induced inflammation and acute lung injury. J Immunol 190(4): 1714-1724. 
  2. Guo, W. A., Davidson, B. A., Ottosen, J., Ohtake, P. J., Raghavendran, K., Mullan, B. A., Dayton, M. T. and Knight, P. R., 3rd (2012). Effect of high advanced glycation end-product diet on pulmonary inflammatory response and pulmonary function following gastric aspiration. Shock 38(6): 677-684.
  3. Hutson, A. D., Davidson, B. A., Raghavendran, K., Chess, P. R., Tait, A. R., Holm, B. A., Notter, R. H. and Knight, P. R. (2006). Statistical prediction of the type of gastric aspiration lung injury based on early cytokine/chemokine profiles. Anesthesiology 104(1): 73-79. 
  4. Raghavendran, K., Davidson, B. A., Mullan, B. A., Hutson, A. D., Russo, T. A., Manderscheid, P. A., Woytash, J. A., Holm, B. A., Notter, R. H. and Knight, P. R. (2005). Acid and particulate-induced aspiration lung injury in mice: importance of MCP-1. Am J Physiol Lung Cell Mol Physiol 289(1): L134-143.
  5. Segal, B. H., Davidson, B. A., Hutson, A. D., Russo, T. A., Holm, B. A., Mullan, B., Habitzruther, M., Holland, S. M. and Knight, P. R., 3rd (2007). Acid aspiration-induced lung inflammation and injury are exacerbated in NADPH oxidase-deficient mice. Am J Physiol Lung Cell Mol Physiol 292(3): L760-768. 

  Rat models

  1. Davidson, B. A., Knight, P. R., Helinski, J. D., Nader, N. D., Shanley, T. P. and Johnson, K. J. (1999). The role of tumor necrosis factor-alpha in the pathogenesis of aspiration pneumonitis in rats. Anesthesiology 91(2): 486-499.
  2. Davidson, B. A., Knight, P. R., Wang, Z., Chess, P. R., Holm, B. A., Russo, T. A., Hutson, A. and Notter, R. H. (2005). Surfactant alterations in acute inflammatory lung injury from aspiration of acid and gastric particulates. Am J Physiol Lung Cell Mol Physiol 288(4): L699-708. 
  3. Kennedy, T. P., Johnson, K. J., Kunkel, R. G., Ward, P. A., Knight, P. R. and Finch, J. S. (1989). Acute acid aspiration lung injury in the rat: biphasic pathogenesis. Anesth Analg 69(1): 87-92.
  4. Knight, P. R., Druskovich, G., Tait, A. R. and Johnson, K. J. (1992). The role of neutrophils, oxidants, and proteases in the pathogenesis of acid pulmonary injury. Anesthesiology 77(4): 772-778.
  5. Knight, P. R., Rutter, T., Tait, A. R., Coleman, E. and Johnson, K. (1993). Pathogenesis of gastric particulate lung injury: a comparison and interaction with acidic pneumonitis. Anesth Analg 77(4): 754-760.
  6. Knight, P. R., Davidson, B. A., Nader, N. D., Helinski, J. D., Marschke, C. J., Russo, T. A., Hutson, A. D., Notter, R. H. and Holm, B. A. (2004). Progressive, severe lung injury secondary to the interaction of insults in gastric aspiration. Exp Lung Res 30(7): 535-557. 
  7. Nader-Djalal, N., Knight, P. R., Davidson, B. A. and Johnson, K. (1997). Hyperoxia exacerbates microvascular lung injury following acid aspiration. Chest 112(6): 1607-1614.
  8. Nader-Djalal, N., Knight, P. R., 3rd, Thusu, K., Davidson, B. A., Holm, B. A., Johnson, K. J. and Dandona, P. (1998). Reactive oxygen species contribute to oxygen-related lung injury after acid aspiration. Anesth Analg 87(1): 127-133.
  9. Nader, N. D., Knight, P. R., Bobela, I., Davidson, B. A., Johnson, K. J. and Morin, F. (1999). High-dose nitric oxide inhalation increases lung injury after gastric aspiration. Anesthesiology 91(3): 741-749.
  10. Nader, N. D., Knight, P. R., Davidson, B. A., Safaee, S. S. and Steinhorn, D. M. (2000). Systemic perfluorocarbons suppress the acute lung inflammation after gastric acid aspiration in rats. Anesth Analg 90(2): 356-361.
  11. Nader, N. D., Davidson, B. A., Tait, A. R., Holm, B. A. and Knight, P. R. (2005). Serine antiproteinase administration preserves innate superoxide dismutase levels after acid aspiration and hyperoxia but does not decrease lung injury. Anesth Analg 101(1): 213-219, table of contents.
  12. Rotta, A. T., Shiley, K. T., Davidson, B. A., Helinski, J. D., Russo, T. A. and Knight, P. R. (2004). Gastric acid and particulate aspiration injury inhibits pulmonary bacterial clearance. Crit Care Med 32(3): 747-754.
  13. Raghavendran, K., Davidson, B. A., Knight, P. R., Wang, Z., Helinski, J., Chess, P. R. and Notter, R. H. (2008). Surfactant dysfunction in lung contusion with and without superimposed gastric aspiration in a rat model. Shock 30(5): 508-517.
  14. Raghavendran, K., Davidson, B. A., Huebschmann, J. C., Helinski, J. D., Hutson, A. D., Dayton, M. T., Notter, R. H. and Knight, P. R. (2009). Superimposed gastric aspiration increases the severity of inflammation and permeability injury in a rat model of lung contusion. J Surg Res 155(2): 273-282.
  15. Raghavendran, K., Davidson, B. A., Hutson, A. D., Helinski, J. D., Nodzo, S. R., Notter, R. H. and Knight, P. R. (2009). Predictive modeling and inflammatory biomarkers in rats with lung contusion and gastric aspiration. J Trauma 67(6): 1182-1190.
  16. Shanley, T. P., Davidson, B. A., Nader, N. D., Bless, N., Vasi, N., Ward, P. A., Johnson, K. J. and Knight, P. R. (2000). Role of macrophage inflammatory protein-2 in aspiration-induced lung injury. Crit Care Med 28(7): 2437-2444.

  Rabbit model

  1. Knight, P. R., Kurek, C., Davidson, B. A., Nader, N. D., Patel, A., Sokolowski, J., Notter, R. H. and Holm, B. A. (2000). Acid aspiration increases sensitivity to increased ambient oxygen concentrations. Am J Physiol Lung Cell Mol Physiol 278(6): L1240-1247.

简介

下面描述的方法用于在小鼠中产生胃吸入性肺炎,其中大鼠和家兔进行了改变。我们使用4种不同的伤口车内气道来调查胃抽吸的炎症反应:
1)生理盐水(NS)作为损伤车辆对照
2)NS + HCl,pH = 1.25(酸)
3)NS +胃颗粒,pH≈5.3(部分)
4)NS +胃颗粒+ HCl,pH = 1.25(酸+部分)
体积,pH值和胃液浓度都会影响所产生的肺损伤。在小鼠中,我们通常使用的损伤体积为3.6ml / kg(大鼠:1.2ml / kg,兔子:2.4ml / kg),伤害pH(含酸载体)为1.25,胃颗粒浓度在含颗粒的载体中)10mg / ml(大鼠:40mg / ml)。在我们手中,这导致了最大,非致命的肺损伤,对于最有害的车辆(即酸+部分),死亡率低于10%。最大可耐受的颗粒浓度需要根据经验确定任何新的菌株的使用,特别是在遗传改变的小鼠中,因为改变的炎症反应可能对死亡率有不利影响。
我们在C57BL / 6背景下,在近交系CD-1以及许多遗传修饰的近交染色体上使用了这些方法。应该仔细考虑应变的选择,特别是在应变特异性免疫偏倚方面,以确保适当的数据解释。受伤时,小鼠的大小应≥20 g。如果需要,可以尝试较小的小鼠,但外科手术变得越来越困难,并且手术存活率显着降低。大鼠和兔子模型没有尺寸或应变约束,但是我们通常在初始损伤时使用250-300克的长伊文思大鼠和约2公斤的新西兰白老鼠。

关键字:肺炎, 急性肺损伤(阿里), 急性呼吸窘迫综合征(ARDS), 炎症, 啮齿类动物模型

材料和试剂

  1. 异氟烷
  2. 局部消毒杀微生物剂制备溶液(例如Medline Industries,目录号:MDS093906)
  3. 0.5%布比卡因
  4. Hank's平衡盐溶液(HBSS)与Ca 2+ 2+,Mg 2+ 2+ (例如Life Technologies,目录号:14025)。
  5. HBSS,不含Ca 2+ 2+ ,Mg 2 + (例如Life Technologies,目录号:14175)
  6. 液氮
  7. 牛血清白蛋白(BSA)
  8. Cytospin过滤卡(例如 Thermo Fisher Scientific,目录号:5991022)
  9. Diff Quik溶液试剂盒(Fisher Scientific,目录号:NC9943455)
  10. Cytoseal 60(VWR International,目录号:48212-154)
  11. 100x蛋白酶抑制剂混合物(例如 Calbiochem ,目录号:80053-844)
  12. 布比卡因
  13. 100mg/ml小鼠(大鼠)胃颗粒(见Recipes)
  14. 酸伤害溶液(酸)(参见配方)
  15. 胃颗粒损伤溶液(部分)(参见配方)
  16. 酸+颗粒损伤溶液(酸+部分)(参见配方)
  17. 磷酸盐缓冲盐水(PBS),pH 7.2(见Recipes)
  18. 氯化铵溶解缓冲液(见配方)
  19. 肺匀浆缓冲液(见配方)
  20. 50mM磷酸钾缓冲液,pH = 6.0(参见配方)
  21. MPO匀浆缓冲液(见配方)

注意: 除非另有说明,所有盐和其他化学物质均来自Sigma-Aldrich(但是,此类化学物质的任何来源可能都可以使用)。

设备

  1. 1-0编织丝材料,散装卷轴(例如查看目录号:MBJF210)
  2. 12 mm x 75 mm x 1 mm显微镜载玻片
  3. 2×2纱布海绵,8层(VWR International,目录号:82004-740)
  4. 无菌4×4纱布海绵,8层(VWR International,目录号:82004-742)
  5. 6-0单丝聚丙烯缝合线与P-13切割针(例如Syneture,目录号:SP-5695)
  6. 60°倾斜剥离板(自制有机玻璃)
  7. 注射器(0.5,1,3,5,20cc(立方厘米))
  8. 针(14,20,22,26,29规格)
  9. 气管套管(23号×1/2"不锈钢管适配器)(Becton,Dickinson and Company,目录号:408213)
  10. 3"弯曲锯齿镊子(2)
  11. 3"弯曲组织("齿")镊子
  12. 1.8 ml微量离心管
  13. 12 x 75毫米聚苯乙烯管
  14. 22 x 22毫米#1.5盖玻片
  15. 4"弯曲显微解剖剪刀
  16. 37°C水浴
  17. 一次性皮肤吻合器(如 3M,型号:DS-25)
  18. 血细胞计数器或库尔特计数器(例如 Beckman Coulter,型号:MultiSizer III)
  19. 带有细胞离心管漏斗的细胞离心机(例如 Shandon CytoSpin ®
  20. 组织匀浆器(例如,Brinkmann TM Polytron TM ,型号:PT2000)
  21. 探头超声仪(例如 Branson Sonifier ®,型号:450)

程序

  1. "损伤"程序
    1. 用0.2ml空气"追加剂"然后"损伤"溶液(大鼠:1cc注射器,具有14号针头,具有0.5ml空气,兔子:5cc注射器,具有14号针头,2ml空气) 。
    2. 在以1-2L/min递送的3-4%异氟烷(兔:30mg/kg氯胺酮,在2%异氟烷之前肌内)递送的室中诱导麻醉(图1和图2)。 暂停鼠标的前牙 在60°斜面解剖板上用1-0缝合线保持麻醉,用鼻锥用2-2.5%异氟烷维持麻醉。


      图1.用于外科手术的实验室工作台设置。在通风橱中进行用于麻醉气体清除的木炭过滤的过程,其包括:小动物麻醉气体暴露室(右前),灯和有机玻璃60°倾斜解剖板(中间的罩)。在图片的左边,在发动机罩外的地板上,是用于输送异氟烷蒸气的麻醉蒸发器。


      图2.用于抽吸损伤程序的仪器。 有机玻璃60°倾斜解剖板显示与鼻锥设置,以在手术期间提供异氟烷。绿色软管将100%氧气中的异氟烷递送到内部鼻锥,而较大孔蓝色软管提供真空用于通过外部鼻锥清除麻醉剂。还示出了(从左到右):具有26号针的5cc注射器,其含有用于皮下注射的1.5ml生理盐水,具有22号针的0.5cc注射器,其含有损伤溶液和0.2ml空气"追加器",3个弯曲解剖钳(2个锯齿状和1"齿状"),弯曲解剖剪刀,一个具有棉头敷药器的防腐抗微生物溶液瓶和6"股1-0编织丝线材料。图片缺失:一次性皮肤吻合器。

    3. 剃刮腹侧颈,涂抹局部防腐制剂溶液,并用纱布除去多余的
    4. 用100μl0.5%布比卡因(提供局部术后镇痛)浸润未来切口部位。
    5. 用剪刀剪切皮肤上2厘米的纵向切口(图3)

      图3.颈切口用缝合线悬挂小鼠的前切牙,使其鼻子在内异氟烷鼻锥内(步骤A2),准备手术部位(步骤A3和A4 ),用剪刀和组织钳制作2cm纵向切口(步骤A5)
    6. 通过钝性解剖用2个弯曲的无齿钳暴露气管(图4,5和6)

      图4.通过钝性解剖的气管暴露。用弯曲的锯齿镊子梳理筋膜,并且唾液腺被拉到侧面以暴露气管(在镊子之间的微白的垂直)被气管肌肉组织包围(步骤A6)

      图5.钝性解剖的气管暴露 持续。握住气管肌肉组织的一侧,将其拉向侧面,同时沿着肌肉纵向摩擦。这将导致肌肉纤维束的分离并允许肌肉被反射到侧面(步骤A6)。注意内部和外部鼻锥配置,用于输送麻醉和清除。为了清楚起见,鼠标的鼻子位置低于通常的位置。


      图6.钝性解剖的气管暴露 持续。使用两个弯曲的锯齿镊子反映气管的肌肉系统,完全暴露气管(步骤A6)。

    7. 在气管下工作弯曲钳(图7)。并且使用通过(图8)拉出6"股1-0丝缝合材料

      图7.气管下工作弯曲锯齿钳(步骤A7)


      图8.使用镊子在气管下放置缝合线(步骤A7)

    8. 通过去除鼻锥停止异氟烷给药。在通过以下步骤灌注伤害车辆之前,允许小鼠的麻醉平面上升,直到它刚刚开始对镊子脚趾捏合反应,然后迅速启动滴注步骤。特别是对于含酸的伤害车辆,如果麻醉平面太深,在伤害车辆被滴注之后动物将不会自发地呼吸。
    9. 用缝合线提起气管,以便于用斜面外科医生(直到针的斜面刚好通过气管插入点)将伤口注射器针插入气管(在喉下方2-3个软骨环)(图9)。重要的是,针与气管尽可能平行,否则针可以容易地刺穿气管的另一侧。这将导致受伤车辆不被注射到肺中,并且很可能动物将不能存活

      图9.使用缝合线便于将损伤溶液注射到气管中。 从倾斜板上取下鼻锥装置,以便不受限制地进入气管上段。用缝合线提起气管,以方便将损伤注射器针插入气管(喉下方2-3个软骨环)。注意针尖外科医生的斜面。提前针直到针的斜面刚好通过气管插入点(步骤A9)。重要的是,针与气管尽可能平行,否则针可以容易地刺穿气管的另一侧。这将导致损伤车辆不被注射到肺中,并且很可能动物将不能存活。虽然辅助挤压肋骨笼(以排出大部分重要的能力),快速灌注注射器内容。在注射开始之前释放胸部。这种操纵以及空气"追赶"确保伤害车辆沉积到远端肺中(步骤A10)。

    10. 虽然辅助挤压肋骨笼(以排出大部分重要的能力),快速灌注注射器内容。在注射开始之前释放胸部。这种机动,以及空气"追赶"确保伤害车辆沉积到远端肺
    11. 将动物留在倾斜板上,直到呼吸开始并用2个吻合钉闭合切口(对于大鼠和兔子:气管针伤口需要用一个6-0缝合线修复)(图10)。

      图10.用手术吻合钉闭合颈部皮肤切口(步骤A11)

    12. 注射1ml(大鼠:10ml,兔子:20ml)无菌NS皮下注射到颈部的颈部用于液体复苏。在手术过程中实际上没有液体损失,但动物在手术后不会喝一会儿,并且可能脱水(图11)。


      图11.液体复苏使用带26号针的注射器将1 ml无菌NS皮下注射到颈部的颈部,用于液体复苏(步骤A12)。

    13. 放入用100%O 2灌注的加热室(37℃)直到可走动(图12)。


      图12.恢复室。将鼠标放在连续灌注100%O 2(由绿色软管提供)并用加热灯保持在37℃的恢复室中,温度控制器(通知左室中的热敏电阻探针),直到鼠标走动(步骤A13)
  2. 收获程序(图13)


    图13.用于收获的仪器和材料从左到右 右:两个锯齿状解剖钳,组织("齿")镊子, 解剖剪刀,23规格不锈钢蓝色hubbed套管,1 cc 血液采集注射器与26规针,支气管肺泡灌洗 装置(顶部注射器,其填充有不含Ca 2+的5ml HBSS,Mg 2+),左 注射器用于流体收集),两个2×2纱布海绵(左下) 用于收缩腹部器官,和4"股1-0编织丝 缝合(下,中)。 图片缺失:5 cc注射器与26 包含具有Ca 2+ 2 H +的HBSS的5ml针,用于冲洗肺的   脉管系统
    1. 用100%O 2中的异氟烷麻醉小鼠。
    2. 当小鼠无反应时,以仰卧位置放置在解剖板上,并用鼻锥继续异氟烷给药。
    3. 使用镊子和剪刀,做一个纵向切口腹部到xyphid
    4. 在腹部进行横向切口(图14),并使用纱布海绵反映腹部器官,暴露腔静脉和腹主动脉(图15)。


      图14.收集血液的腹部切口。 使用镊子和剪刀,沿着腹部纵向切开至腹部,并穿过腹部进行横向切口(步骤B3和B4)。


      图15.收集血液。使用纱布海绵来暴露腔静脉和腹主动脉以反映腹部器官,并使用1cc注射器用26号针从两个容器中收集血液(步骤B4和B5)。< br />
    5. 收集血液(用于评估全身性炎症反应(例如血清或血浆细胞因子水平)):
      1. 使用27号针头在1cc注射器(如果要分离血浆,注射器必须包含适当的抗凝剂)从腹主动脉收集血液(图15)。
      2. 腔静脉可用于收集血液,但通常不能从主动脉收集血液(图15)。
      3. 将血液分配到1.8 mlμ-fuge管中。 直到血液可以被处理,如果血浆将准备在冰上,或保持在室温,如果血清将被准备。 处理血液,如下所述。
      4. 横断腔静脉/腹主动脉通过exsanquination安乐死
    6. 冲洗肺血管系统(执行从肺循环中去除血液,因此测量处理的肺组织中的化合物仅反映来自不包括全身贡献的肺区室的值):
      1. 使用镊子抓住xyphoid过程,穿刺隔膜用剪刀尖放气肺,并仔细切除隔膜的腹侧面(图16)。


        图16.切开的隔膜。用镊子抓住xyphoid过程,用剪刀尖穿刺隔膜,使肺缩小,并仔细切除隔膜的腹侧面(步骤B6a)。

      2. 继续沿胸骨和颈部纵向切开。
      3. 切开胸骨以暴露肺部(注意不要刺伤肺部)(图17)。


        图17.进行胸骨切开术。沿着胸骨和颈部继续纵向切口。切开胸骨以暴露肺(小心不要刺伤肺)(步骤B6b-c)。

      4. 用无齿镊握住心尖,并注射5mL(大鼠:20ml)1×具有Ca 2+ 2 +,Mg 2+(在37℃)的HBSS进入右心室(大鼠:20cc)注射器+ 26号针(图18),将它放置在心脏的左侧(因为动物在其背部)。注射速率应当足够快以保持心脏"充气",但不能快到迫使流体通过注射部位流出。缓冲液中的钙及其在37℃,有助于心脏保持跳动,以便于冲洗肺血管系统。


        图18.冲洗肺脉管系统。用锯齿钳(组织钳具有撕裂心脏组织的倾向)抓紧心尖,并注射5ml 1x HBSS与Ca 2+ 2 + ,Mg 2+(在37℃)进入右心室(在该图中在心脏的左侧,因为动物在其背部)用5cc注射器+ 26号针头。注射速率应该足够快以保持心脏"充气",但不能快到迫使流体通过注射部位流出(步骤B6d)。

    7. 通过进行支气管肺泡灌洗(BAL)(进行以收集分泌到空间(即肺泡和支气管腔)中的细胞和化合物,可以评估它们以确定肺部炎症的程度(即, (TNF-α),白细胞介素-1β(IL-1β),IL-6,巨噬细胞炎症蛋白-2(MIP-2),嗜中性粒细胞, ,单核细胞趋化蛋白-1(MCP-1))和肺损伤(即总蛋白或白蛋白浓度):
      1. 暴露气管和弯曲镊子的工作尖在它下面拉一条4"股1-0丝绸缝合线。
      2. 将23号x 1/2"(大鼠:14号,兔:3mm ID,4.5mm OD)不锈钢插管(兔:聚乙烯导管)插入气管并用缝合线固定(图19和20) br />

        图19.插入气管套管。 按照损伤程序(步骤A6,A7和A9)中所述,将气管和弯曲镊子的工作尖暴露于其下,拉出1-0编织丝的4"股,使用20号针在上气管中制作一个洞,并在孔中插入一个23号1/2"不锈钢插管

        图20.固定气管套管。用缝线将套管固定在气管中(步骤B7a-b)。确保缝线紧紧,并在插入位点下面2-3 mm,以便进行良好的密封。插管尖端应在隆突前至少3mm(气管分叉)。如果1/2"套管插入喉部正下方,如图所示,套管尖端将处于正确的位置。

      3. 将连接灌洗装置(2个,5cc注射器连接到4路活塞)导管和缓慢,灌洗肺与5×1毫升(大鼠:10毫升,兔:50毫升)1 HBSS无Ca 2+ 2+/sup>, Mg 2+(在37℃)(图21和22)。缓冲液中缺乏钙有助于通过降低依从性来收获气道细胞

        图21.进行支气管肺泡灌洗(BAL)注射。将灌洗装置连接到套管,并缓慢注射1ml 1×HBSS与Ca 2+,Mg 2+, 以硫化肺(步骤B7c)。


        图22.执行BAL收集。 切换旋塞阀,并慢慢收集注射的灌洗液。重复注射和收集过程,共进行5次肺洗涤(步骤B7c)。

      4. 将收集的BAL液(BALF)分配到适当大小的管中,记录回收体积(通过称重),并置于冰上直到BALF处理,如下所述。
    8. 收获肺(进行以评估不同于空气空间的肺炎症状态,即实质和间质组织):
      1. 通过气管将肺,心脏和胸腺整合在一起(图23)。


        图23.收集肺,心脏和胸腺, 。确定锁骨已切除。用手指抓住套管,气管和缝线,并用喉切开气管。当拉起和拉出时,工作剪刀背在肺下并扩展以脱离将肺和心脏连接到胸腔的组织。将需要切除结缔组织,以及食道和血管,以成功地集中肺,心脏和胸腺(步骤B8a)。

      2. 通过在肺门处切除肺叶主干支气管(图24和25),置于2ml冷冻管中并在液氮中快速冷冻除去肺叶。


        图24.收获肺叶。使用剪刀和镊子收集单个肺叶,以切除肺叶的主干支气管(步骤B8b)。


        图25.小鼠肺叶。从左侧顺时针:左肺叶,右肺 - 颅盖,右肺中叶,右肺尾叶,右肺 - 后腔静脉。 >
      3. 存储在-80°C,直到可以进行适当的处理,如下所述
  3. 血液处理
    1. 等离子体
      1. 在4℃下以2,000xg离心4分钟。
      2. 用移液管取出血浆,分装到1.8ml微量离心管中
      3. 储存于-80℃。
    2. 血清
      1. 在室温下孵育30分钟,然后在4℃过夜,以使凝块收缩
      2. 在4℃下以5,000xg离心15分钟。
      3. 用移液管取出血清,分装到1.8ml微量离心管中
      4. 储存于-80℃。

  4. BALF处理(将样品保存在冰上)
    1. 在4℃下,在1,500×g下旋转BALF 5分钟。
    2. 在不干扰沉淀的情况下,用移液管收集上清液(留下50-100μl剩余体积的未收集的上清液以防止吸出细胞),并分配到等体积的等分试样在1.8ml微量离心管中(等分试样的数量和体积将取决于测定 将被执行并且应该被设计为限制冻融循环的量)。 储存于-80℃。
    3. 在剩余体积中重悬沉淀。
    4. 加入1ml氯化铵裂解缓冲液(37℃),孵育2分钟以裂解红细胞
    5. 在12ml 75mm PS管中的4ml冰冷的2%BSA的PBS中的整个体积层
    6. 在4℃下以150×g离心15分钟。
    7. 吸出上清液(含细胞碎片和裂解红细胞),重悬沉淀的残余体积,并加入1ml PBS。 通过将细胞离心通过2%BSA除去细胞碎片和裂解的红细胞使得细胞计数更容易和更准确。
    8. 在血细胞计数器或库尔特计数器上计数细胞
    9. 准备细胞离心片幻灯片用于微观观察:
      1. 将5×10 4个白细胞加入附于显微镜载玻片和滤纸卡的细胞离心漏斗(300μl总体积)中的PBS中的2%BSA中,并以28×g (500rpm)5分钟
      2. 取出载玻片并风干(重要:干燥时立即染色)。
      3. 用Diff Quik染色:
        1. 在固定剂,溶液I,溶液II(Diff-Quik试剂盒)中浸渍载玻片5×1秒,用H 2 O 风干。
        2. 安装幻灯片与小滴"Cytoseal 60"在细胞点上,放一个22 x 22毫米#1.5盖玻片(确保没有气泡),并允许风干。
        3. 通过光学显微镜确定巨噬细胞,嗜中性粒细胞,淋巴细胞和嗜酸性粒细胞的百分比
  5.  肺处理(肺可以以多种不同的方式进行,取决于什么细胞区室(例如全肺匀浆,核部分,胞质部分,等)分子种类(例如蛋白质,mRNA,等),以及将利用什么测定技术(例如ELISA,Western印迹, PCR等)。该程序产生肺匀浆上清液,其可以通过ELISA测定各种细胞因子蛋白水平,以及从肺匀浆沉淀中提取髓过氧化物酶,可以测定酶促活性作为嗜中性粒细胞浸润的替代标记)
    1. 解冻冷冻的肺并转移到能够耐受40,000×g的圆底离心机中。在加肺之前称重管,然后称重管+肺
    2. 加入足够的细胞因子匀浆缓冲液,使肺+缓冲液的重量达到3g(大鼠:10g)。
    3. 使用Polytron匀浆器在冰上均质化组织以防止样品的加热。用镊子取出均质器刀片中抓住的任何组织。用70%EtOH冲洗匀浆器,然后无菌 H 2 O
    4. 在4℃下以40,000×g离心匀浆10分钟。
    5. 将上清液分配到等体积的等分试样在1.8ml微量离心管中并储存在-80℃
    6. 加入2 ml MPO缓冲液沉淀,并通过涡旋重悬
    7. 存储在-80°C(在同一管),直到MPO提取可以执行
  6. MPO提取程序
    1. 在37℃水浴中快速解冻样品。
    2. 超声处理1分钟,在冰上防止样品加热,在50%占空比的最大输出
    3. 在55℃孵育2小时
    4. 在4℃下以40,000×g离心15分钟。
    5. 分配到0.5ml等分试样在1.8ml微量离心管中,并存储在-80℃下,直到MPO活动可以评估

食谱

  1. 100mg/ml小鼠(大鼠)胃颗粒
    1. 早上从新鲜尸检鼠(大鼠)第一件事收获胃,并置于冰管50毫升。 早晨收获保证胃饱满。
    2. 在层流罩
      1. 把胃放在培养皿中,用剪刀纵向切开
      2. 将胃内容物放在干净的50ml离心管中
      3. 加入10ml无菌生理盐水(NS),并剧烈涡旋15秒
      4. 通过8层无菌纱布将颗粒悬浮液倒入烧杯中
      5. 使用额外的10ml无菌NS将管中的残留颗粒转移到纱布上
      6. 拧紧纱布收集吸收的液体。
      7. 将滤液转移到干净的50ml离心管中
    3. 在4℃下以5,000xg离心5分钟旋转滤液
    4. 弃去上清液,并重悬于25ml无菌NS中
    5. 重复步骤1c& 1d。
    6. 将颗粒悬浮液转移到锥形瓶中,121℃,121psi压力下30分钟(使用另外25ml,无菌NS转移残余颗粒)。
    7. 将冷却的无菌颗粒悬浮液转移到预称重的50ml离心管中
    8. 在4℃下以5,000xg离心5分钟旋转滤液
    9. 在KIMWipe上倒置上清液和倒置的管以除去所有液体
    10. 重新称量管+颗粒丸以确定颗粒湿重
    11. 将无菌NS添加至100mg/ml(颗粒湿重/总体积)
    12. 在4°C存储长达3周,直到用于受伤
  2. 酸伤溶液(酸)
    生理盐水(NS),无菌
    9.44 ml
    1 N HCl,无菌
    562微升
    用无菌1N HCl调节pH = 1.25 新鲜
  3. 胃颗粒损伤溶液(部分)
    生理盐水(NS),无菌
    9 ml
    100 mg/ml NS中的胃粒子
    1ml(10mg/ml)
    让新鲜。
  4. 酸+颗粒损伤溶液(酸+部分)
    生理盐水(NS),无菌
    8.44 ml
    1N HCl,无菌
    562微升
    100 mg/ml NS中的胃粒子
    1ml(10mg/ml)
    用无菌1N HCl调节pH = 1.25 新鲜
  5. 磷酸盐缓冲盐水(PBS),pH 7.2
    NaCl(58.44)
    8g(137mM)  

    1.15g(8.1mM)
    KCl(74.56)
    200mg(2.7mM)
    KH 2 PO 4 (136.09) 
    200mg(1.5mM)
    H <2> MQ 到1 L
    调节pH = 7.2,过滤灭菌
    在室温下贮存
  6. 氯化铵裂解缓冲液
    NH 4 Cl(53.49)
    4.13g(154mM)
    KHCO 3 (100.1)
    500毫克(10毫摩尔)
    EDTA,四钠盐(380.2)
    18.5mg(0.1mM)
    彻底将粉末混合在一起
    将等量分配到10,50ml离心管(0.46g /管)中 使用当天,加入50ml H 2 O O MQ,并混合直至溶解
    丢弃任何未使用的解决方案
  7. 肺匀浆缓冲液
    NaCl(58.44)
    8.77克(150毫摩尔)
    Tris碱(121.14)
    1.82g(15mM)

    CHCl 2 2H O(147.02)
    147毫克(1毫摩尔)

    203mg(1mM)
    H <2> MQ 到1 L
    调节pH = 7.4,在121℃,15psi下高压灭菌15分钟
    储存在4°C
    在使用时,将100倍蛋白酶抑制剂混合物的1/100体积加入到一天处理所需的肺匀浆缓冲液体积中
  8. 50mM磷酸钾缓冲液,pH = 6.0
    KH 2 PO 4 (136.01)
      3.4克(50mM)

    H O MQ 到500ml
    用2M NaOH调节pH = 6.0 在室温下贮存
  9. MPO匀浆缓冲液
    十六烷基三甲基溴化铵(364.5)
      2克(0.5%)

    EDTA(372.24)  
      744 mg(5 mM)

    50mM磷酸钾缓冲液,pH = 6.0   400 ml

     储存在室温下(不要冷藏)

致谢

这里提出的工作是由NIH授予HL048889,"Pathogenesis of Aspiration Pneumonitis"给Paul R Knight,M.D.,Ph.D. 和Bruce A. Davidson,Ph.D。 要引用此协议,请同时使用以下参考文献:Davidson (2013)。

参考文献

 鼠标模型

  1. Davieson,BA,Vethanayagam,RR,Grimm,MJ,Mullan,BA,Raghavendran,K.,Blackwell,TS,Freeman,ML,Ayyasamy,V.,Singh,KK,Sporn,MB,Itagaki,K.,Hauser, ,Knight,PR和Segal,BH(2013)。 NADPH氧化酶和Nrf2调节胃抽吸诱导的炎症和急性肺损伤 J Immunol 190(4):1714-1724。 
  2. Guo,W.A.,Davidson,B.A.,Ottosen,J.,Ohtake,P.J.,Raghavendran,K.,Mullan,B.A.,Dayton,M.T.and Knight,P.R.,3rd(2012)。 高度晚期糖基化终末产品饮食对胃部吸入后对肺部炎症反应和肺功能的影响。/a> Shock 38(6):677-684。
  3. Hutson,A.D.,Davidson,B.A.,Raghavendran,K.,Chess,P.R.,Tait,A.R.,Holm,B.A.,Notter,R.H.and Knight,P.R。(2006)。 基于早期细胞因子/趋化因子谱的胃抽吸肺损伤类型的统计预测。 104(1):73-79。
  4. Raghavendran,K.,Davidson,B.A.,Mullan,B.A.,Hutson,A.D.,Russo,T.A.,Manderscheid,P.A.,Woytash,J.A.,Holm,B.A.,Notter,R.H.and Knight, 小鼠中酸和颗粒物诱发的肺损伤:MCP-1的重要性。 Am J Physiol Lung Cell Mol Physiol 289(1):L134-143。
  5. Segal,B.H.,Davidson,B.A.,Hutson,A.D.,Russo,T.A.,Holm,B.A.,Mullan,B.,Habitzruther,M.,Holland,S.M.and Knight,P.R。,3rd(2007)。 酸性吸入诱导的肺部炎症和损伤在NADPH氧化酶缺陷小鼠中加剧。 Am J Physiol Lung Cell Mol Physiol 292(3):L760-768。 

 大鼠模型

  1. Davidson,B.A.,Knight,P.R.,Helinski,J.D.,Nader,N.D.,Shanley,T.P.and Johnson,K.J。(1999)。 肿瘤坏死因子-α在大鼠吸气性肺炎的发病机制中的作用。 91(2):486-499
  2. Davidson,B.A.,Knight,P.R.,Wang,Z.,Chess,P.R.,Holm,B.A.,Russo,T.A.,Hutson,A.and Notter,R.H。(2005)。 急性的表面活性剂改变 炎症性肺损伤,其来自酸和胃颗粒的吸入。美国生理学杂志(J Physiol Lung Cell Mol Physiol)288(4):L699-708。
  3. Kennedy,T.P.,Johnson,K.J.,Kunkel,R.G.,Ward,P.A.,Knight,P.R.and Finch,J.S。(1989)。 大鼠急性酸性吸入性肺损伤:双相发病机制 Anesth Analg 69(1):87-92。
  4. Knight,P.R.,Druskovich,G.,Tait,A.R.and Johnson,K.J。(1992)。 中性粒细胞,氧化剂和蛋白酶在酸性肺损伤发病机制中的作用。 麻醉学 77(4):772-778
  5. Knight,P.R.,Rutter,T.,Tait,A.R.,Coleman,E。和Johnson,K。(1993)。 胃颗粒肺损伤的发病机制:与酸性肺炎的比较和相互作用 Anesth Analg 77(4):754-760。
  6. Knight,PR,Davidson,BA,Nader,ND,Helinski,JD,Marschke,CJ,Russo,TA,Hutson,AD,Notter,RHand Holm,BA(2004)。进行性的严重肺损伤继发于胃抽吸中的侮辱的相互作用。 Exp Lung Res 30 ):535-557。 
  7. Nader-Djalal,N.,Knight,P.R.,Davidson,B.A。和Johnson,K。(1997)。 高氧会加重酸性吸入后的微血管性肺损伤。 胸部 112(6):1607-1614。
  8. Nader-Djalal,N.,Knight,P.R.,3rd,Thusu,K.,Davidson,B.A.,Holm,B.A.,Johnson,K.J.and Dandona,P。(1998)。 反应性氧物质会在酸吸入后造成与氧相关的肺损伤。 Anesth Analg 87(1):127-133。
  9. Nader,N.D.,Knight,P.R.,Bobela,I.,Davidson,B.A.,Johnson,K.J.and Morin,F。(1999)。 高剂量一氧化氮吸入增加胃抽吸后的肺损伤麻醉 91(3):741-749。
  10. Nader,N.D.,Knight,P.R.,Davidson,B.A.,Safaee,S.S。和Steinhorn,D.M。(2000)。 系统性全氟化碳抑制大鼠胃酸吸入后的急性肺部炎症。 Anesth Analg 90(2):356-361。
  11. Nader,N.D.,Davidson,B.A.,Tait,A.R.,Holm,B.A。和Knight,P.R。(2005)。 丝氨酸抗蛋白酶施用在酸性吸入和高氧后保留先天超氧化物歧化酶水平,但不减少肺损伤。/a> Anesth Analg 101(1):213-219,目录。
  12. Rotta,A.T.,Shiley,K.T.,Davidson,B.A.,Helinski,J.D.,Russo,T.A。和Knight,P.R。(2004)。 胃酸和颗粒性吸入性损伤会抑制肺部细菌清除。 Crit Care Med 32(3):747-754。
  13. Raghavendran,K.,Davidson,B.A.,Knight,P.R.,Wang,Z.,Helinski,J.,Chess,P.R。和Notter,R.H。(2008)。 在大鼠模型中,在有和没有叠加胃抽吸的肺挫伤中的表面活性剂功能障碍。 em> Shock 30(5):508-517。
  14. Raghavendran,K.,Davidson,B.A.,Huebschmann,J.C.,Helinski,J.D.,Hutson,A.D.,Dayton,M.T.,Notter,R.H.and Knight,P.R。 叠加的胃吸入增加了肺挫伤的大鼠模型中的炎症和通透性损伤的严重性。 a> J Surg Res 155(2):273-282
  15. Raghavendran,K.,Davidson,B.A.,Hutson,A.D.,Helinski,J.D.,Nodzo,S.R.,Notter,R.H.and Knight,P.R。(2009)。 具有肺挫伤和胃抽吸的大鼠的预测建模和炎症生物标志物。 J Trauma 67(6):1182-1190。
  16. Shanley,T.P.,Davidson,B.A.,Nader,N.D.,Bless,N.,Vasi,N.,Ward,P.A.,Johnson,K.J.and Knight,P.R。(2000)。 巨噬细胞炎症蛋白-2在吸气诱导的肺损伤中的作用。 Crit Care Med 28(7):2437-2444。

  兔模型

  1. Knight,P.R.,Kurek,C.,Davidson,B.A.,Nader,N.D.,Patel,A.,Sokolowski,J.,Notter,R.H.and Holm,B.A。(2000)。 酸性抽吸会增加环境氧气浓度的增加。美国生理学肺 Cell Mol Physiol 278(6):L1240-1247。
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Copyright: © 2013 The Authors; exclusive licensee Bio-protocol LLC.
引用:Davidson, B. A. and Alluri, R. (2013). Gastric Aspiration Models. Bio-protocol 3(22): e968. DOI: 10.21769/BioProtoc.968.
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