Protocol for performing traumatic spinal cord injury surgery on mice in germ-free isolators and assessing injury recovery and pathophysiology

Summary

An altered gut microbiome affects recovery from traumatic spinal cord injury (SCI). Here, we present a protocol for performing SCI survival surgery on mice in germ-free isolators. Specifically, we describe steps for colonization, preparing surgical tools, performing surgery, and post-surgical care. We then outline procedures for behavioral assessment, followed by collecting and processing tissue to assess injury recovery and pathophysiology. This protocol can be easily adapted to study the gut microbiome’s influence on other forms of central nervous system injury.

Graphical abstract

Highlights

• •Steps for performing spinal cord injury survival surgery in germ-free isolators
• •Instructions for colonizing germ-free mice
• •Guidance on analyzing neurological recovery in germ-free isolators

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

An altered gut microbiome affects recovery from traumatic spinal cord injury (SCI). Here, we present a protocol for performing SCI survival surgery on mice in germ-free isolators. Specifically, we describe steps for colonization, preparing surgical tools, performing surgery, and post-surgical care. We then detail procedures for behavioral assessment, followed by collecting and processing tissue to assess injury recovery and pathophysiology. This protocol can be easily adapted to study the gut microbiome’s influence on other forms of central nervous system injury.

Before you begin

Interactions between the gut microbiome and the central nervous system (CNS) are well established.^1,2,3 It is unsurprising then that conditions which disrupt CNS, like a spinal cord injury, also have profound impacts on the gut microbiome.^4,5,6,7 Use of a germ-free mouse model of SCI clarifies gut-CNS interactions in health and disease. The methods below outline how to perform SCI surgery in germ-free and colonized mice, allowing for the study of the CNS and gut microbiome changes that occur after injury. It is important to note that these instructions are intended for individuals who possess foundational experience in operating and maintaining sterile isolators. If you are not familiar with these methods, please refer to reference books such as Gnotobiotic Mouse Technology: An Illustrated Guide by Chriss J. Vowels.⁸

Innovation

To our knowledge, this is the first published step-by-step protocol outlining how to perform a standardized traumatic spinal cord injury in a germ-free isolator using the murine model. Specifically, we provide steps to assess neurological recovery over time in the germ-free isolator, how to colonize mice, and guidelines for histological assessment. We also demonstrate how to modify surgical tools to ensure successful surgery. The practices outlined in the protocol can be easily modified to model other central nervous system injuries. Thus, this protocol opens up a new area of research for the field of traumatic central nervous system injury by providing the tools necessary to study the influence of the microbiome on injury pathophysiology.

Institutional permissions

Obtain ethical approval from your institution before beginning these procedures. All described procedures and experiments complied with the guidelines and regulations of The Ohio State University and the National Institute of Health’s Guide for the Care and Use of Laboratory Animals. All animal-related procedures were approved by the Institutional Animal Care and Use Committee of the Office of Responsible Research Practices at The Ohio State University. Male and female germ-free C57BL/6J mice were acquired from Taconic Biosciences and bred in our germ-free facility for these experiments. Mice typically ranged in age from 3 to 6 months, but any age can be used for the protocol, depending on your experimental question. All mice received sterilized commercial food pellets (Teklad global 19% protein extruded rodent diet sterilizable; Cat # 2019S) and sterilized, germ-free chlorinated reverse osmosis water ad libitum. All animals were housed 5 mice/cage in cages containing Inotiv Teklad Diamond soft bedding (Inotiv Cat #7089). Housing facilities had a 12-h light-dark cycle at a constant temperature of 20°C ± 2°C and humidity of 50% ± 20%.

Colonization of germ-free mice before spinal cord injury

Timing: ∼1–2 h depending on group size; perform these steps 28 days prior to surgery

Note: For studies comparing germ-free controls to mice colonized with whole fecal transplants or specific microbial communities (such as Schlegal flora or cultured bacteria), be sure to carry out colonization at least 28 days before surgery. This timeframe was selected to ensure both microbial and immunological stability. While initial bacterial engraftment occurs within days, ecological stabilization of the gut microbiota generally requires 3 to 4 weeks to reach a state that closely resembles the donor profile.^9,10,11 Crucially, the host immune system requires a longer period for maturation. Specific adaptive immune responses, such as secretory IgA production and the expansion of intestinal T-cell populations, typically stabilize only after 4 weeks of continuous microbial exposure.^12,13 This 28-days period therefore establishes a robust homeostatic baseline, minimizing the potential for confounding results caused by transient post-colonization physiological states.¹¹ From this point onwards we will refer to colonization material as a fecal microbiota transplant (FMT) and the procedures outlined below are for whole fecal transplants but could be modified for select microbial communities. Germ-free mice housed in a dedicated germ-free isolator may serve as control subjects by administering an equal volume of sterile 0.9% saline via gavage, rather than the FMT as described below. We have also included a timeline of how to integrate this optional colonization step into the pre-surgery protocol (See Figure 1).

• 1.
  Prepare FMT for oral gavage using fecal pellets obtained from conventional mice.

  • a.
    Collect 1–2 freshly emitted fecal pellets from C57BL/6J mice by scruffing mice then holding a sterile Sarstedt 2ml screw cap tube (Fisher Scientific Cat # NC0418367) below the anus. Avoid collecting any urine.
  • b.
    Add cold, sterile 1xPBS (Fisher Scientific Cat # 10010049) to the pellets (1 mL 1xPBS per 100 mg of fecal material)
  • c.
    Homogenize the pellets by vortexing on a small bench top vortex for 2–5 min.
  • d.
    Centrifuge the sample (∼10–20s) using a benchtop centrifuge to pellet any large particulate matter.
  • e.
    Transfer the supernatant into new sterile 1.5 mL Eppendorf tubes (Eppendorf Cat # 022363204).

Note: Transferring the supernatant will result in a substantial loss of volume since much fecal material is insoluble. Collect and homogenize twice the needed number of fecal pellets to ensure enough volume for gavage.

• 2.
  Gavage germ-free mice with FMT slurry.

  • a.
    To sterilize the tubes containing the prepared suspension and disposable intragastric gavage needles (Fisher Scientific Cat # 0120887), soak them for a minimum of 20 min in cold sterilant (Spor-Klenz RTU Cat # 652501).
  • b.
    “Enter” the suspension tubes and gavage needles into the germ-free isolator via the port and pull through into the isolator.

    Note: You do not need to spray sterilize the port prior to entering the gavage needles, as the whole isolator will soon be colonized.

  • c.
    Administer 200 μl of the suspension by oral gavage to the germ-free mice designated for colonization.

    Note: Gavage mice within 2 h f fecal pellet collection. Although human studies show FMT is stable for up to 24 h (fresh) and longer with freeze-dried samples, we have not tested stability beyond 2 h in mice. Therefore, we do not recommend storing FMT samples for longer without confirming bacterial viability.^14,15

    CRITICAL: Once the mice are gavaged with fecal slurry, the isolator is no longer “germ-free”-- all mice in the isolator are now considered colonized. If you do not want all mice in the isolator to be colonized, be sure to move them to a second germ-free isolator before Step 2. Once the experiment is complete, you should decommission the colonized isolator(s), then re-sterilize before introducing any new germ-free mice into that isolator.

• 3.
  Collect fecal samples to verify success of colonization.

  Note: To ensure successful colonization or, in the case of germ-free mice, maintenance of a germ-free gut, one must test for the presence of gut microbiota. Therefore, in the weeks leading up to the surgery date, autoclave Sarstedt 2 mL screw cap tubes, then enter them into the isolator to collect the fecal pellets (see “surgical supply entry into germ-free isolators”, below).

  • a.
    Two days prior to your planned surgical date, collect 1–2 fecal pellets from each mouse as described in Step 1a.
  • b.
    Remove fecal pellets from the isolator and immediately freeze them in liquid nitrogen.
  • c.
    Store the collected fecal pellets at −80°C.

    Note: Analyze these samples using 16S sequencing or other analytic tools (e.g. shotgun sequencing) at the end of the experiment to verify that the colonized mice were successfully colonized and that the germ-free control mice remained germ-free. Alternatively, to quickly screen for colonization, you can plate homogenized fecal samples on blood agar plates (Fisher Scientific Cat # R01202), incubate for 24 h, and check for bacterial growth (see Figure 4A for example results).

Figure 1.

Example timeline of procedure

This example timeline depicts the minimum length of time needed to complete colonization of germ-free mice, entry of surgical supplies into the germ-free isolators, collection of pre-injury fecal samples, surgery, and post-surgery care and behavior. The timeline depicts the minimum 28d period for stable colonization, but a longer time period can be used if needed, or a shorter time period could be used if your experimental question involves exploring dynamic changes of colonization. Alternatively, if you do not need colonized mice for your design, this step can be excluded. The length of the surgical period will also depend on the number of surgical cohorts you need to conduct for your study. The post-surgical period can also be adjusted depending on the experimental questions and outcomes, hence why the final day post-injury (dpi) is depicted as the variable X. Cartoon timeline schematic created with Biorender.com.

Figure 4.

Schematic of Surgical Steps to Complete a Thoracic Level 9 Complete Crush Spinal Cord Injury

(A) A cartoon depiction of the mouse’s positioning for surgery, set up of surgical support pillow, and surgical skin prep.

(B) Depiction of the surgical incision location.

(C) Cartoon showing the rotated potion of the mouse and the fat pads exposed after the surgical incision is completed. The real surgical image inset depicts where the location-identifying blood vessel between vertebral T5 and T6 is located.

(D) Schematic again showing rotated body position of the mouse as well as placement of the retractor into the surgical site. The real surgical image inset outlines the incision path along the spinal column that should be followed to expose the spinal cord beneath the tissue.

(E) A cartoon showing forceps and retractor placement necessary to clear the tissue from the vertebrae. The realimage inset of a mouse vertebral column with all surrounding tissue cleared depicts the changes in vertebral direction (highlighted by arrows) that occurs at the T9 level to help clarify this anatomical landmark.

(F) A cartoon depiction of the different stages of a successful T9 laminectomy. The real surgical image insets demonstrate bone and tissue removal (i), identification of residual ligamentum flavum (LF) for removal, and a final image depicting a fully cleared spinal cord section and successful laminectomy (iii).

(G) A depiction of the expected line of bruising after a complete crush injury. All cartoon schematics created withBiorender.comwith original surgical inset images.

Surgical supply entry into germ-free isolators

Timing: ∼2 weeks, perform these steps in the 2 weeks leading up to your planned surgery date

Note: All supplies needed for surgery and post-surgery care must be sterilized with either cold sterilant, a dry sterilization autoclave cycle, or hydrogen peroxide before entering them into the isolators. This is true even for the colonized isolator to ensure that there is no introduction of additional environmental bacteria not already present in the FMT used to colonize mice. As mentioned, this protocol assumes that you are familiar with the basic steps of entering and exiting supplies through germ-free isolator ports. Please refer to the reference book Gnotobiotic Mouse Technology: An Illustrated Guide for any additional guidance needed.⁸

• 4.
  Enter recovery cages, bedding, food, and empty water bottles.

  • a.
    In an autoclavable metal cylinder wrapped with DW4 filter media (CBClean Cat # 3901875), add sterile standard mouse cage bottoms, wire tops, empty water bottles, bedding, and food.

    Note: Enter at least two empty cages. One cage will be designated for holding mice during anesthesia and surgical preparation, and another for use during post-surgery recovery. Include two cages per sex if you are using different sexes.

  • b.
    Sterilize supplies with an autoclave gravity cycle of dry heat at 125°C for 1 hour.

    CRITICAL: Include steam indicator strips (e.g. Steris Cat # 802210) in the autoclavable cylinder and a self-contained biological indicator (SCBI; Steris Cat # S3061) each time you autoclave.

    Note: After all supplies and steam indicators are entered into the autoclave cylinder, seal with mylar film (CBCClean Cat # 39105175) and tape (ULINE Cat # S-20217BLU) as per standard germ-free isolator procedures.⁸

    • i.
      Check that the steam indicator strips changed color, indicating successful autoclaving, before entering supplies into the isolator.
    • ii.
      Upon removing the SCBI from the autoclave, verify that the indicator has turned brown to confirm proper exposure to steam during the autoclave cycle.
    • iii.
      Break the spore disk inside the indicator with the vial activator (Steris Cat # S3075).
    • iv.
      Incubate the whole vial for 48 h at 55°C–59°C.

      Note: The autoclave cycle is successful if there is no color change or bacterial growth in the biological indicator and the supplies can be entered into the isolator.

  • c.
    Attach the autoclavable cylinders with sterile supplies to the isolator ports.
  • d.
    Sterilize the port with cold sterilant passed through an atomizer.
  • e.
    Allow the supplies to sit overnight (∼12 h).
  • f.
    Pull the supplies into the isolator by piercing the mylar film from the inside with crucible tongs (Fisher Scientific Cat #15–207).
• 5.
  Prepare and enter water to refill water bottles.

  • a.
    Wash at least one glass bottle (1L; DWK Life Sciences Cat # 219946305) per isolator with a standard laboratory dishwasher.
  • b.
    Fill clean bottles with reverse osmosis water.
  • c.
    Cap the bottles by soaking disposable rubber caps (DWK Life Sciences Cat # 292063002) in warm water for at least 10 min.

    • i.
      Invert the soaked cap on top of the bottle and snap down the sides of the cap around the spout.
    • ii.
      Make a half turn to ensure proper placement and seal.
  • d.
    Autoclave the filled and sealed bottles on a liquid setting that maintains 125°C for 60 min.

    CRITICAL: Include a liquid steam indicator (Steris Cat # PCC024). The indicator will change in color from red to green/deep blue when it is sterile.

  • e.
    Soak both the supplies and a replacement port cap in a tub of cold sterilant for at least 20 min prior to entering the bottles into the isolator port.

    • i.
      Place the soaked supplies into the port
    • ii.
      Cap the port with the soaked replacement port cap (CBCClean Cat #2212120) and rubber stoppers (Thomas Scientific Cat #1201G62).
    • iii.
      Spray cold sterilant into the port using an atomizer.
    • iv.
      Leave the bottles in the port overnight (∼12 h).
    • v.
      Open the port and pull supplies into the isolator.
• 6.
  Prepare autoclavable surgical tools and supplies (see Table 1).

  • a.
    Collect and autoclave surgical supplies by placing them into a metal autoclave cylinder.
  • b.
    Autoclave as described in Steps 4b-f above.

Note: Store surgical tools in a stainless-steel sterilization container (Fine Science Tools Cat # 20850–00) with a filter (Fine Science Tools Cat #20850–10) to protect them during autoclaving.

• 7.
  Prepare surgical supplies that need to be cold-sterilized (see Table 1).

  • a.
    Collect the supplies listed in Table 1 that require cold-sterilization.
  • b.
    Soak the individual sterile packages in cold sterilant for a minimum of 20 min.
  • c.
    Enter the supplies into the port.
  • d.
    Sterilize the port with cold sterilant and an atomizer.
  • e.
    Keep the cold-sterilized supplies in the port for ∼12 h overnight before opening.
  • f.
    Transfer supplies into the isolator.
• 8.
  Prepare surgical supplies for hydrogen peroxide sterilization.

  • a.
    Collect items requiring hydrogen peroxide sterilization listed in Table 1.
  • b.
    Place in Techni-Vent Header bags with a vaporized process indicator.

    Note: This indicator will change from pink to yellow after sterilization.

    CRITICAL: It is essential to use this particular header bag for hydrogen peroxide sterilization. After evaluating several brands, this was the only product that remained intact without leaking following submersion in cold sterilant, ensuring the sterile transfer of contents into germ-free isolators.

  • c.
    Sterilized via hydrogen peroxide.

    Note: Include a Vaporized VH202 Process Indicator (Steraffirm Cat # PCC063) and a S24 SCBI (Spordex Cat # NA340) SCBI) in each load.

  • d.
    Once sterilization is complete, activate the indicator vial to break the SCBI and release the media.

    • i.
      Incubate the vial for 24 h at 55°C–60°C.

      Note: If the media stays orange, this indicates successful sterilization. However, if it turns yellow and appears cloudy, the load has not been sterilized and should not be placed inside the isolator.

  • e.
    After confirming sterilization, soak the header bag in cold sterilant for 20 min.
  • f.
    Entering items into the isolator as described in 7a-f.

    Note: If you wish to test isolator sterility after entering the surgical supplies, standard methods such as testing of the isolator mold trap to assess environmental sterility, plating of fecal samples from mice (as described in Step 3), or sending sentinel mice fecal samples for 16S PCR bacterial testing can be done. More detailed methods for these standard germ-free maintenance procedures can be found in the reference book Gnotobiotic Mouse Technology: An Illustrated Guide.⁸

Table 1.

Surgical supplies to enter into germ-free isolator

Steam sterilization (autoclavable)	Cold-sterilization	Hydrogen-peroxide sterilization
Small surgical gauze	Sterile 1mL syringes	Electronic Razors
Sterile Cotton Tip Swabs	Sterile 30G needles	Battery power/Solar power scales
Glass Sharps Container	Sterile 18G needles	Spring Scales
2mm straight spring scissorsa	Sterile 10mL syringes	–
4mm straight spring scissorsa	Sterile insulin syringes	–
Friedman-Pearson Rongeursa	Eye lubricant	–
Extra Fine Bonn Scissorsa	Sterile 70% Ethanol	–
Scalpel Handlea	Sterile 0.9% saline	–
Serrated Micro-Adson Forceps (2)a	Cautery pen	–
Modified Micro-Adson Forcepsa	Sterile 50ml conical falcon tube	–
Wound Clip Applier	Ketamine	–
Wound Clips	Xylazine	–
Wound Clip remover	Atipamezole	–
Needle Holders with Suture Cutters	Surgical sutures	–
Retractora	Disposable heating packs	–
Dumont # 5 Forceps (1 regular, 1 modified)a	Reflective emergency recovery blankets	–
Dumont #4 Forcepsa	Scalpel Blades	–
Sterilization Container with filter	Small petri dishes	–
	GelFoam® Sterile Sponge	–

^aIndicates supplies specific for SCI surgeries. Supply lists can be customized for other protocols.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Chemicals, peptides, and recombinant proteins
Spor-Klenz Ready-to-Use Sterilant (cold sterilant)	Steris Life Sciences	Cat # 6525M2
0.9% Sterile Saline	Pfizer Hospital (or similar)	Cat # 00409-7101-04
Sterile 70% Denatured Ethanol (32 oz)	Texwipe	Cat # TX3265
Sterile Water for Injection, Preservative Free (10mL)	Pfizer Hospital (or similar)	Cat # 00409-4887-17
Systane® NIGHTIME eye ointment	Alcon	N/A
Sterile 1xPBS ph7.4	Fisher Scientific (or similar)	Cat # 10010049
Experimental models: Organisms/strains
C57BL/6J Mice (any age or sex depending on experimental question)	Jackson Laboratories	Strain #:000664
Germ-Free C57BL/6NTac Mice (any age or sex depending on experimental question)	Taconic Biosciences	B6-M (male) or B6-F (female)
Other
Husbandry Supplies
Surgical Germ-Free Isolator	Park Bio	custom
Teklad Global 19% Protein Extruded Rodent Diet (Sterilizable)	Inotiv	Cat # 2019S
Tekland Diamond Soft Bedding	Inotiv	Cat #7089
DURAN® 45 mm Push-on Natural Rubber Cap	DWK Life Sciences	Cat # 292063002
DURAN® borosilicate 3.3 glass Autoclave bottle	DWK Life Sciences	Cat # 219946305
Sterilization Supplies
Stoppers Rubber, Black #6.5 Solid	Thomas Scientific	Cat #1201G62
12″ x 12″ port cap	CBCClean	Cat # 2212120
DW4 Filter Media	CBCClean	Cat # 3901875
Autoclave cylinder (12″ x24″)	CBCClean	Cat # 28122401
Mylar film disc (17.5″ diamter) (pk of 25)	CBCClean	Cat # 39105175
Hi temp nylon tape (1″ x 36yards) pk 10	ULINE	Cat # S-20217BLU
Techni-Vent Header bag (Pouch), 10″x15″ Header	Technipac Inc.	Contact company
Crucible Tongs	Fisher Scientific	Cat #15-207
Thermo Scientific™ Blood Agar (TSA with Sheep Blood) Medium	Fisher Scientific	Cat # R01202
VERIFY™ Flash Integrator	Steris	Cat # 802210
VERIFY™ Dual Species Self Contained Biological Indicators (SCBI)	Steris	Cat # S3061
VERIFY™ Vial Activator Set	Steris	Cat # S3075
Vaporized VH202 Process Indicator	Steraffirm	Cat # PCC063
S24 Self-Contained Biological Indicator (SBCI)	Spordex™	Cat # NA340
Steris Sterafirm Control Tube for Steam (122°C for 15 min)	Steris	Cat # PCC024
Colonization Supplies
Intragastric Gavage Needles	Fisher Scientific	Cat # 0120887
Sarstedt Inc Screw Cap Micro tube 2mL, PP 1000/case	Fisher Scientific	Cat # NC0418367
Fisherbrand™ RNase-Free Disposable Pellet Pestles with tube	Fisher Scientific	Cat # 12-141-268
Eppendorf™ Safe-Lock 1.5mL Mircotube	Eppendorf (or similar)	Cat # 022363204
Surgical Supplies
Filters for Dual Clasping Stainless Steel Sterilization Container	Fine Science Tools	Cat # 20850-10
Dual Clasping Stainless Steel Sterilization Container	Fine Science Tools	Cat # 20850-00
Straight Vannas Spring Scissors- 2mm cutting edge	Fine Science Tools	Cat # 1500-03
Straight Vannas Spring Scissors- 4mm cutting edge	Fine Science Tools	Cat # 15018-10
Curved Micro Friedman-Pearson Rongeurs	Fine Science Tools	Cat # 16221-14
Extra Fine Bonn Scissors (Sharp/Sharp)	Fine Science Tools	Cat # 1408-08
Scalpel Handle # 3	Fine Science Tools	Cat # 1003-12
Scalpel Baldes #15	Fine Science Tools	Cat # 10015-00
Micro-Adson Forceps (Serrated)	Fine Science Tools	Cat # 11018-12
AutoClip ® System (applier, wound clips, & remover)	Fine Science Tools	Cat # 12020-00
Olsen-Hegar Needle Holders with Suture Cutters	Fine Science Tools	Cat # 12002-12
Moria Colibri Retractor	Fine Science Tools	Cat # 17300-02
Dumont #4 Forceps	Fine Science Tools	Cat # 11241-30
Dumont # 5 Forceps	Fine Science Tools	Cat # 11251-30
McKesson Argent™ Surgical Systems Cautery Tip	McKesson	Cat # 927942
Gauze Sponge McKesson 2 X 2 Inch 12-Ply NonSterile 200 per Pack	McKesson	Cat # 446029
Biosyn™ monofilament absorbable suture, taper point, violet (size 5)	Medtronic	Cat # CM-875
Swab Cotton 3″ Wood Pointed	Thomas Scientific	Cat # 1212W19
BravMini+™ Purple Clippers	Wahl Professionals	Cat # 41590-0438
Fisher Science Education™ Compact Scale	Fisher Scientific (or similar)	Cat # S72422
Weigh basket (Apipi-Pencil Basket-Round-12 Set)	Apipi (or similar)	Cat # 881395685191
GelFoam® Sterile Sponge	Pfizer Hospital (or similar)	Cat # 00009039605
BravMini+™ Purple Clippers	Wahl Professionals	Cat # 41590-0438
KIMBLE® GLS 80® Laboratory Bottle, Wide mouth, Amber, 250 mL autoclaveable glass bottle for sharps container	DWK Life Sciences (or similar)	Cat # 14394-0250
1mL BD Tuberculin Syringe Only	BD (or similar)	Cat #309659
30G X 1 BD PrecisionGlide™ Needle	BD (or similar)	Cat #305128
18G X 1 BD PrecisionGlide™ Needle	BD (or similar)	Cat # 305195
10mL BD Luer-Lok™ Syringe only	BD (or similar)	Cat #302995
Individually Wrapped Insulin Syringes	Allison Medical (or similar)	Cat # 90-8295
Spring Scale	United Scientific (or similar)	Cat # SB0100
Post-Surgery Recovery Supplies
Instant Hot Pack (5x7 in)	McKesson (or similar)	Cat # 520561
Emergency Rescue Blanket (Cut to appropriate size)	ULINE (or similar)	Cat # S-23360
C2Dx T/Pump® Circulating water heater	C2Dx	Cat # TP700
C2Dx T/Pump® Mult-T-Pads, Lightweight single-patient use with Clik-Tite connectors (water heating pads)	C2Dx	Cat # 800-0620912
Falcon™ Bacteriological Petri Dishes with Lid	Falcon (or similar)	Cat # 351008
Falcon™ 50 mL High Clarity Conical Centrifuge Tubes	Falcon	Cat # 352070

Materials and equipment

Custom germ-free isolators with a glass-viewing surgical area on top

To achieve optimal visibility during surgery, we developed germ-free isolators that Park Bio, LLC custom-built (Figure 2A). Our design included a surgical “shelf” topped with a glass roof (Figure 2Bi) to support microscopic visualization by allowing the surgical scope to be placed above the field (Figure 2Bii). While positioning the surgical scope outside a microisolator is an option, conventional bubble isolators typically use plastic covers that are not suitable for high-precision procedures, especially for surgeons with limited or moderate experience (Figure 2C). Our custom-designed surgical shelf provides a dedicated space for surgical procedures, distinct from the primary housing area, which minimizes clutter and facilitates more efficient access (Figure 2D). The inclusion of gloves on both sides of the surgical zone enables two surgeons to operate concurrently (with two scopes), or allows one surgeon to perform surgery while the other assists or closes the site, thereby enhancing procedural efficiency (Figures 2B–2D). Although the custom isolator offers many advantages, a traditional isolator can be used, as in Figure 2C.

Figure 2.

Schematics of custom germ-free isolators for SCI surgery

(A) Design plans and measurements for custom germ-free isolator with surgical area.

(B) Close-up image of surgical area with glass top without the dissection scope (i) and with the surgical scope in place (ii).

(C) A makeshift surgical table (1) and a surgeon performing laminectomy through one of two conventional microisolator access ports (2). Another surgeon (3) prepares a GF mouse for surgery.

(D) Image of full isolator to demonstrate adjacent surgical area and housing area.

(E) Image of water heater and pad attached to the bottom of the surgical area of the isolator. The heating pad is used to maintain body temperature during and after surgery.

Modification of surgical tools

In order to optimize surgical procedures within the microisolators, we have adapted a number of commercially available surgical instruments (Figure 3). This includes the modification of Dumont # 5 Forceps (Figure 3Ai,ii). The forceps’ outer edges are trimmed with a Dremel, and the tips are shortened and curved inward (Figure 3Aii) to ensure secure grip and spinal stabilization during the crush. We also built custom retractors using bendable wire filament to get the appropriate size to hold open the surgical area (Figure 3Bi,ii). If you prefer not to make your own, you can find similar retractors available commercially (for example, the Moria Colibri Retractor from Fine Science Tool, Cat # 17300-02). Additionally, we modified Micro-Adson Forceps (Serrated) by using a Dremel to trim the outer edges of the serrated tips (Figure 3Ci,ii). This allows for better grip of the vertebrae during the laminectomy procedure (Figure 3Cii). Finally, we modified rongeurs, by shaving the outer edges with a Dremel (Figure 3D). This modification makes laminectomy surgery, i.e., removal of the vertebral bone, easier. Although many of these changes simplify the surgical procedure and lower the chance of accidentally damaging the spinal cord, only the adjustment to the forceps shown in Figure 3A is crucial for achieving reliable and repeatable spinal cord injuries in microisolators.

Figure 3.

Images of modified surgical tools

(A) Image depicting modified and standard (commercially available) Dumont # 5 forceps from the front (i) and side (ii). Black arrows indicate areas of modification by thinning down with a small handheld Demel.

(B) Image of custom-build retractor from the side with ruler for size reference (i) and from a top-down view (ii).

(C) Image depicting modified and standard Serrated Micro-Adson Forceps from the front (i) and side (ii). Black arrows indicate areas of modification by thinning down with a small handheld Demel.

(D) Image depicting modified and standard Friedman-Pearson Rongeurs. Black arrows indicate areas of modification by thinning down with a small handheld Demel. Original technical images.

Step-by-step method details

Prepare the isolator for surgery

Timing: ∼1 h

This section explains how to prepare the germ-free isolator interior for surgery. Complete these steps the day before the procedure.

• 1.Turn on circulating water warmers (C2Dx Cat # TP700 and # 800-0620912) under the main isolator area and surgical area before starting first mouse.

Note: We attached the warming blankets of the circulating water warmers to the bottom of the isolator using either strong tape or Velcro (Figure 2E). Use distilled water in the pumps to avoid mineral buildup. Ensure O-rings on the heating pad connections are intact, or leaking may occur once the pump is turned on.

• 2.Prepare an anesthesia cage by placing a wire top on an empty cage to house animals after administering anesthesia.
• 3.
  Prepare a recovery cage.

  • a.
    Place an empty cage the main isolator area above the water heater.
  • b.
    Put a disposable heating pack inside the cage.
  • c.
    Cover it with a recovery blanket.

Note: You will need two recovery cages with you are using different sexes.

• 4.
  Prepare food mash for the recovery diet

  • a.
    Add a few pellets of autoclaved food (Teklad Global 19% Protein Extruded Rodent Diet (Sterilizable) Inotiv Cat # 2019S) into a 50mL Falcon tube.
  • b.
    Add sterile drinking water to the tube to submerge the pellets.
  • c.
    Allow the food to soak and become soft while performing surgeries.

Note: After the mash is made, it can be placed into small petri dish halves (Falcon Cat # 351008) and placed on the floor of the housing cages for mice post-surgery.

• 5.Check the electronic scale is functioning properly and equipped with a weigh basket.

Note: The spring scale can be used as a backup if the scale is non-functional.

• 6.Check that your electronic razors are working.
• 7.Dilute your ketamine, xylazine, and atipamezole to the appropriate concentrations for anesthesia.

Note: These concentrations will depend on your institutionally approved animal use protocols. For references, we prepare concentrations to be able to administer the following doses: ketamine (80 mg/kg), xylazine (10 mg/kg), and atipamezole (1 mg/kg).

• 8.
  Set up the microscope above the surgical area of the isolator.

  • a.
    Focus clearly over the surgical field.

Anesthesia and preparation of the surgical site

Timing: ∼10–30 min per mouse

This section details the appropriate procedures for anesthetizing mice and preparing the surgical site for spinal cord injury.

Note: It is helpful to have 3–4 people assist with performing surgeries to avoid having to continuously exit and re-enter the gloves on the isolator. As an example, one person records anesthesia and body weights, a second handles anesthesia, prepares the site, and manages recovery, a third performs surgery, and a fourth (optional) closes the surgical site.

• 9.
  Weigh the mouse using the scale and weight basket.

  • a.
    Record mouse weight.

Note: Keeping track of the mouse's weight after surgery will help you monitor their health and ensure they are eating and drinking well. If they are not, more mash can be provided, and additional supplemental saline can be given (2 mL subcutaneously with morning bladder expression and 1 mL subcutaneously with evening bladder expression).

• 10.Inject the mouse intraperitoneally using an insulin syringe with a weight-appropriate dose of ketamine (80 mg/kg) and xylazine (10 mg/kg).

Note: Mice usually take 10–15 min to become fully anesthetized. If, after 5–10 min, mice do not show signs of anesthesia—such as slower movements or absence of whisker twitching—supplemental anesthetic may be given. See troubleshooting problem 1 for additional information.

CRITICAL: When performing the intraperitoneal injection, it is essential to avoid injecting into the cecum. Germ-free mice have large cecums, usually located on the mouse’s left-hand side. The most effective approach is to administer the anesthetic on the lower right side of the mouse, roughly aligned with the middle of the knee while the mouse is scruffed. Cecal injections often cause incomplete anesthesia and higher mortality, as additional doses may increase the risk of overdose.

• 11.Shave the mouse just below the ears to just below the ribs with the electronic razor to clear the surgical site.
• 12.Wipe away excess fur and clean the skin with sterile gauze dampened with sterile 70% alcohol to further clean and clear the surgical site.
• 13.Apply eye gel to keep eyes moist during surgery.
• 14.
  Check that the mouse has reached an appropriate surgical plane of anesthesia before initiating surgery.

  • a.
    Gently pinch all four paw reflexes with fingers or forceps.

Note: There should be an absence of limb and whisker movement and steady breathing when the mouse is in the surgical plane.

Thoracic level 9 complete crush spinal cord injury

Timing: ∼15–20 min per mouse

These steps explain how to expose the thoracic spinal cord and perform a T9 laminectomy followed by a spinal cord crush injury.

• 15.
  Place the anesthetized mouse ventrally on a piece of sterile gauze (Figure 4A).

  • a.
    Face the mouse’s head towards you.
  • b.
    Place a prepared pillow (rolled sterile gauze) just below the mouse’s head.

Note: This will ensure easy maneuvering of the mouse and elevation of the spinal cord during surgery.

• 16.
  Use standard serrated micro-Adson forceps (Figure 3Ci-ii) to identify the area between the shoulder blades.

  • a.
    Use the forceps to pick up the loose skin.

    Note: Forceps should be held in your non-dominant hand.

  • b.
    Using your dominant hand, cut the skin in between from the shoulder blades with the extra fine Bonn scissors from the top of the shoulder blades to just below the bottom of the ribs.

    Note: The cut is long enough when you see exposed ligaments on each side of the spinal cord, they should appear silvery/white (Figure 4B).

• 17.Turn the mouse so the head is away from you.
• 18.Use the scissors and the serrated forceps to pick up the thin layer of brown fat on the now exposed muscle.

Note: It will appear slightly colored and shiny.

• 19.Cut the brown fat down the middle to allow access to two white fat pads underneath this brown fat layer.
• 20.Use two serrated forceps to gently pull apart the now visible white fat pads on either side of body’s midline until you see the large vessel between the T5 and T6 vertebrae.
• 21.Lay the fat on either side of the incision to reveal the muscle and tissue on top of the spinal column (Figure 4C).
• 22.Turn mouse to face left.

Note: If you are left-handed, you can turn the mouse to the right to make the incision. Rotate the mouse as needed to complete the incisions on both sides of the vertebral column.

• 23.
  Use a #11 scalpel blade and handle to make an incision in the muscle/on each side of the spinal column (Figure 4D).

  • a.
    Position the blade at an angle toward the vertebral column, rather than aligned parallel to it.
  • b.
    Make the incision as close to the vertebral column as possible.

Note: Each incision should require only 2–3 strokes of the blade until you reach the vertebral bone. Very little bleeding should occur with these incisions. If excessive bleeding does occur, see management techniques in troubleshooting problem 2.

• 24.
  Place the small retractor in the incisions to hold open tissue and muscle.

  • a.
    Orient the joining tip of the retractor towards the tail (Figure 4D).
• 25.
  Clear the tissue surrounding the vertebral column using small spring scissors and clean cotton swabs to reveal the T8, T9 and T10 vertebrae.

  • a.
    With your non-dominant hand, use small modified serrated forceps to stabilize the spinal column around the T8 vertebrae by gently grasping the sides of the vertebral column while clearing tissue with your dominant hand.

Note: The T9 and T10 vertebrae can be distinguished by noting the change in direction of the vertebral arches at T9 (Figure 4E). Alternatively, the blood vessel between T5 and T6 can be identified, and the spinal processes counted to T9; however, this approach may be less precise, as the large blood vessel is not consistently located at this specific site.

• 26.
  Once you have cleared the tissue and identified T9, use rongeurs to perform a T9 laminectomy by removing the T9 spinous process and the arcus vertebrae (the top of the vertebrae on either side of the spinous process).

  • a.
    Hold the T8 vertebral body with the modified serrated forceps held in your non-dominant hand to stabilize the spinal cord as you remove the bone, (Figure 4Fi).

Note: Rather than trying to remove large pieces of bone, slowly remove small pieces of bone; otherwise, bleeding and accidental damage to the spinal cord may occur. It is often also easier to remove one side of the arcus vertebrae at a time. Be careful not to damage the cord while clearing the surrounding tissue. It is critical to remove the bone completely to the sides of the column to ensure that there is enough space to insert the forceps for the crush later (Figure 4Fiii). However, avoid damaging/removing too much muscle on the sides, as there are blood vessels there and damaging them will cause bleeding. If bleeding occurs, see troubleshooting problem 2.

• 27.Clear any ligamentum flavum (LF) remaining on top of the dura with #5 forceps after you have removed the bone (Figure 4Fii,iii).

Note: This will increase visibility and prevent bleeding during the crush injury.

• 28.
  Using microscissors, make small parallel incisions in the muscle on both sides of the spinal column at the T10 level.

  Note: This allows for the placement of modified #5 forceps, which will assist in stabilizing the cord during the crush procedure.

  • a.
    Once these “slots” are formed, turn the mouse so the tail is on the left.
• 29.Confirm under the microscope that the #4 forceps fully close before using them to crush the spinal cord.

Note: Reserve these #4 forceps only for this procedure. This will prevent damage to the tips.

• 30.
  Insert the modified #5 forceps into the “slots” you created in the muscle at the T10 level during Step 26.

  • a.
    Hold with your non-dominant hand to stabilize the mouse.
• 31.
  Insert the #4 forceps so that the tips are along each side of the spinal cord, perpendicular to the long axis of the mouse and in the middle of the vertebral T9 space.

  • a.
    Ensure that the bottom of the spinal column can be felt by the tips.
  • b.
    Very gently move the forceps side to side to verify that you did not puncture the spinal cord when inserting the forceps prior to the crush and that you are deep enough to crush the entire cord.
• 32.
  Tightly close the forceps in one swift motion, “crushing” the spinal cord.

  • a.
    Hold for 2 s (count: “one thousand, two thousand”).
  • b.
    Release and remove the forceps.

    CRITICAL: Crushing for shorter or longer periods of time will yield spinal cord injuries that are less or more severe, respectively, as determined using morphometric measures of lesion pathology and behavioral recovery. Thus, the time can be adjusted to fit your experimental aims.

  • c.
    Verify that a continuous line of bruising appears across the cord once the forceps are removed (Figure 4G).

    Note: If there is not a continuous line of bruising, the spinal cord was not adequately crushed and this mouse should be excluded from the study as the injury is incomplete. If you are unsure if the crush was incomplete, this can be verified in the histological analysis (see expected outcomes section).

  • d.
    If bleeding occurs during crushing, apply saline to the crush site as described in troubleshooting step 2.
• 33.Remove the retractor.
• 34.Close the muscle layers located above the spinal cord with one or two sutures tied 4 times.
• 35.Use saline-moistened cotton swabs to position the pushed-aside fat pads back on top of the surgical site.
• 36.Remove any blood from the skin using saline-moistened cotton swabs, then close the surgical incision with 2–3 surgical wound clips.

Note: If blood is not properly cleaned from the skin at the surgical site, either the other mice in the cage or the operated mouse itself may try to remove the clips after waking up from anesthesia.

Post-surgical anesthetic reversal and care

Timing: ∼1 h per mouse

The following steps provide detailed instructions on how to administer an anesthesia reversal agent (e.g., Atipamezole hydrochloride) and provide post-surgical care as the mouse recovers from anesthesia.

• 37.After closing the surgical site, give an anesthesia reversal agent like Atipamezole hydrochloride (1mg/kg) subcutaneously (30G needle on 1mL syringe) on the mouse’s back to reverse sedation and anesthesia.
• 38.Inject 2mL of 0.9% sterile saline subcutaneously (30G needle on 1mL syringe) into the mouse’s back to restore fluids and replace blood loss.

Note: Use an 18G needle to easily draw-up saline.

• 39.Activate a disposable heating pack and add it to the already prepped recovery cages (Figure 2E).
• 40.
  Place the mouse on top of the activated heating pad to maintain body temperature during recovery.

  • a.
    Occasionally shake the heating pad to keep it heated and/or replace it once the heating reaction is completed.
• 41.Cover the cage with an emergency blanket to keep the heat in.
• 42.
  Monitor mice while they recover from anesthesia.

  • a.
    Once fully awake, transfer mouse to a home cage that has been prepared with food on the bottom and a small petri dish full of prepared mash.
• 43.Keep injured mice above the water heater for the remainder of the experiment.

Note: Post-surgery, mice require manual bladder expression twice daily (morning and evening) and subcutaneous saline administration: 2 mL daily for days 1–3 dpi, and 1 mL daily for days 3–7 dpi. After 7dpi supplemental saline can be administered as needed.

Assessment of sterility post-surgery

Timing: ∼15 min to 1 h to collect samples

The following steps provide detailed instructions on how to determine if germ-free isolators and mice maintained their sterility during the surgery and post-surgery period.

• 44.
  Collect post-surgical fecal samples (1–2 pellets per mouse) as described in Step 3 of the colonization of germ-free mice before spinal cord injury section.

  • a.
    Perform 16S RNS sequencing, 16S PCR, or plate and culture the fecal samples to verify the maintenance of the sterility of germ-free mice and/or colonization of colonized mice after injury.

Note: Fecal samples can be difficult to collect fresh from SCI mice due to reduced intestinal motility after injury. If you are unable to obtain fresh fecal samples from mice after injury, fecal samples can be directly collected from the colon of mice during dissections. Just ensure the use of either autoclaved or flame-sterilize dissection tools for the collection.

• 45.
  Collect a sample from the isolator mold trap to plate.

  • a.
    Culture as per standard procedures to check for environmental sterility.
• 46.Record cecum weight from mice during tissue dissection.

Note: Colonized mice should have reduced cecum weights and germ-mice should have enlarged cecums (see Figure 5 for example data).

Figure 5.

Expected outcomes for complete crush SCI in germ-free and colonized mice

(A) Blood agar plates showing plated fecal samples from colonized (14d post colonization) and germ-free mice show the results of colonization via oral gavage.

(B) Successful colonization can also be determined by seeing a decrease in total cecum weight, as depicted in the graph demonstrating larger cecum weights in germ-free (GF) Naïve and SCI mice compared to colonized Naïve and SCI mice. One-way ANOVA with Tukey’s Post-Hoc Test. ∗∗∗∗ P < 0.0001. Graphs represent mean ± SEM.

(C) Line graphs comparing historical BMS score data from T9 crush and control mice for surgeries performed inside without (Germ-Free SCI) or with colonization (Colonized SCI) and outside an isolator (Conventional SCI). No difference between scores for germ-free and conventional injured mice. Two-way ANOVA. Graphs represent mean ± SEM.

(D) Line graph of historical BMS score data from T9 crush germ-free mice from the acute to chronic recovery period (3d to 28dpi). BMS was performed weekly starting at 7dpi. As expected, mice with complete crush injuries do not recovery a BMS score greater than 2. Graphs represent mean ± SEM.

(E) Horizontal section images of the spinal cord with an Eriochrome Cyanine strain showing typical myelin loss after a complete crush T9 injury in germ-free (i, Scale bars = 200 μm) and an example of an incomplete T4 Crush injury (ii, Scale bars = 200 μm).

(F) Horizontal section images of the typical SCI lesion morphology of astrogliotic scarring (i, visualized with GFAP) and microglia/macrophages accumulation (ii, visualized with CD11b). Scale bars = 200 μm.

Expected outcomes

This protocol outlines how to successfully perform a complete crush SCI in germ-free and colonized mice. It is expected that germ-free mice show no detectable bacteria from fecal homogenate plated on blood agar plates, while colonized mice should have detectable bacteria (Figure 5A). Additionally, colonized mice, both naïve and SCI, typically have cecum weights of less than 2g, whereas large cecum weights are a distinctive feature of germ-free mice (Figure 5B).

A successful SCI should produce consistent outcomes as defined by post-SCI locomotor function and spinal cord pathology. Given the severity of crush SCI used in this protocol, it is expected that mice will not achieve Basso Mouse Scale (BMS)¹⁶ locomotor scores of 1 when assessed at 1dpi or 7dpi (Figures 5C and 5D) or greater than 2 when assessed past 7dpi (Figure 5D). If mice exceed these thresholds, they should be excluded from analyses if a complete crush injury is desired. As seen in Figure 5C, it is possible to achieve consistent BMS scores, whether SCI surgeries are performed in the germ-free isolator or in conventional surgical spaces. Consistent with this level of neurological impairment, one should expect a visibly clear and complete lesion as revealed by staining with an Eriochrome Cyanine (EC) stain for myelin (Figure 5Ei). Specifically, a horizonal spinal cord section of the lesion epicenter (defined as the horizontal section at the level of the central canal) should be devoid of spared myelin from the caudal to ventral area. Animals that do not show this complete lack of spared myelin, as depicted in Figure 5Eii, have an incomplete crush injury and should be excluded from analysis if a complete crush model is desired. Immunohistochemistry should also reveal a well-defined glial border (GFAP staining; Figure 5Dii) and a central zone of inflammation marked by accumulation of activated myeloid cells (microglia/macrophages) (Figure 5Fiii). While the histological images here are representative, a complete description of how to quantify crush injury histology and expected outcomes in control animals can be found in our previously published work.¹⁷

Limitations

Performing experimental models of SCI in a germ-free isolator is primarily limited by the types of injuries that can be performed. As the isolator is a closed sterile system, SCIs that require the use of large mechanical injury devices, like the Infinite Horizon Impactor, which is used to administer graded levels of spinal contusion injury, are not possible. However, the custom-designed surgical shelf described herein, makes it possible to introduce accurate surgical lesions, including graded crush injuries, dorsal hemisections, or funiculotomies. Importantly, these surgical lesions can be created at different spinal levels, enabling a more complete understanding of how the microbiome (or absence thereof) influences different aspects of motor, sensory, and autonomic functions controlled by the spinal cord. A second limitation of performing SCI in microisolators is “throughput”. Because of the numerous logistical difficulties involved with working in microisolators—such as delivering and monitoring anesthesia, overseeing recovery, and performing precise surgery using isolator gloves—each procedure takes about twice as long as it would outside of an isolator. While the custom-build isolators can comfortably house up to 8 cages (40 mice) while still maintaining room to perform surgery, we would not recommend performing more than 10-12 surgeries per day, as each surgery takes ∼30min – 1 hour, excluding the full recovery time. Thus, only smaller group sizes can be completed each day. To address this limitation, groups should be randomly allocated to daily cohorts, with consecutive surgery days scheduled to increase sample sizes. Although this approach could introduce batch effects into the dataset, our findings indicate that when surgeries are conducted on successive days using mice from identical age cohorts, any batch effects are minimal. Another challenge is the difficulty in evaluating behavioral outcomes for mice kept in microisolators. Because mice cannot be removed from these isolators, most behavioral tests—such as kinematic assessments, ladder tasks, and beam walking—are not feasible. To address this, we assess open-field locomotion using the Basso Mouse Scale. Other tests, like the cylinder task for forelimb function, could also be adapted for use within the microisolators.

Troubleshooting

Problem 1

Mouse is unresponsive to anesthesia (related to Step 10).

Germ-free mice frequently do not respond to anesthesia, making it difficult to achieve the depth needed for surgery. This complication is often reduced by avoiding injecting anesthetic into the enlarged cecum of germ-free mice (as outlined in the critical section of step 10); however, challenges may still arise even with proper technique. If the mouse does not respond to anesthesia within 10 min post-administration, proceed with the following steps.

Potential solution

• •Massage the abdomen: If the mouse still shows reflex responses and whisker twitching 10 min after anesthesia administration, gently massage the abdomen with your fingers in a circular motion for 1–2 min. This will help distribute the anesthetic throughout the peritoneal cavity.
• •Administer supplemental anesthesia: If after 10–15 min of anesthesia administration, the mouse is still demonstrating strong reflex responses and/or whisker movements, a supplemental dose of anesthesia can be administered. If the mouse is primarily immobile with weak response, we recommend giving a quarter dose of ketamine/xylazine. It is also possible to administer half or a third of the dose depending on the magnitude of those mouse’s reflex responses. However, if the mouse is freely moving and showing no signs of anesthesia, a second full dose can be administered.
• •Assign mouse to naïve/control group: If two full doses of anesthesia fail to achieve the surgical plane, move the mouse to the control group. More than two doses risk overdose and may prevent recovery after surgery.

Problem 2

Uncontrollable bleeding during surgical procedures (related to Steps 21, 24, and 30).

Occasionally during surgical procedures, unexpected or uncontrollable bleeding can occur. This often happens if blood vessels adjacent to the spinal cord are disrupted as vertebrae are removed during laminectomy. Such bleeding can generally be managed using the methods described below and does not usually affect experimental results. It is essential to manage bleeding without delay, as substantial blood loss may compromise the animal’s chances of surviving the procedure.

Potential solution

• •Apply gentle pressure: In the event of minor bleeding during the procedure, use sterile cotton swabs to apply gentle pressure on the affected area. Once the bleeding stops, new sterile cotton swabs can be used to clear the area and continue with the procedure.
• •Apply 0.9% sterile saline: If the bleeding continues after applying gentle pressure with a cotton swab, flood the surgical site with 0.9% sterile saline. This can be accomplished using a pre-filled syringe and needle. Allow the saline to rest and apply pressure for approximately 1 min, then remove the saline with sterile gauze or small cotton swabs. The added pressure from the saline is usually enough to control bleeding.
• •Apply gel foam: If the application of 0.9% sterile saline fails, you can apply a small piece of GelFoam® sterile sponge to the site of bleeding. This will absorb the blood and promote clotting. You can also apply the foam and then apply 0.9% sterile saline as above if needed. The foam can remain in place while you finish the procedure and then should be removed prior to closure of the surgical site.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Kristina A. Kigerl (kristina.kigerl@osumc.edu).

Technical contact

Technical questions on executing this protocol should be directed to and will be answered by the technical contact, Lori C. Hudson (lori.hudson@osumc.edu).

Materials availability

This protocol did not generate any unique reagents.

Data and code availability

This protocol did not generate any code or large datasets.

Acknowledgments

The laboratory of P.G.P. is supported by the National Institutes of Neurological Disorders and Stroke R35 (1R35NS111582 to P.G.P.) and the Ray W. Poppleton Endowment (P.G.P.). The Belford Center for Spinal Cord Injury also provides support to the laboratory of P.G.P. K.A.K. is supported by a Craig H. Neilsen Foundation Senior Research Award (890085) and an Ohio Department of Higher Education Third Frontier grant. The Center for Brain and Spinal Cord Repair core was supported by NINDS core facility grant P30NS104177.

Author contributions

Conceptualization, K.A.K., L.C.H., K.A.M., and P.P.P.; data acquisition and technical expertise/assistance, K.A.K., K.A.M., L.C.H., and Z.G.; data analysis, K.A.K. and K.A.M.; writing and manuscript revision, K.A.K., L.C.H., K.A.M., Z.G., and P.P.P.; funding acquisition, K.A.K. and P.P.P.

Declaration of interests

The authors declare no competing interests.