Improved Methodology for Studying Postnatal Osteogenesis via Intramembranous Ossification in a Murine Bone Marrow Injury Model

Summary

The murine bone marrow injury model is a useful tool for studying postnatal osteogenesis through intramembranous ossification. This simplified surgical protocol describes the bone marrow ablation procedure for downstream assessment of new bone formation and cellular responses following injury.

Abstract

Long bone injuries heal through either endochondral or intramembranous bone formation pathways. Unlike the endochondral pathway that requires a cartilage template, the process of intramembranous ossification involves the direct conversion of skeletal stem and progenitor cells (SSPCs) into bone-forming osteoblasts. There are limited surgical methods to model this process in experimental mice. Here, we have improved upon a bone marrow injury model in mice to facilitate the study of bone repair via intramembranous ossification and to assess postnatal regulators of osteogenesis. This method is highly reproducible and user-friendly, and it allows temporal assessment of new bone formation in a short period (3-7 days post-injury) using micro-computed tomography (µCT) and frozen section histology. Furthermore, the contributions of SSPCs and mature osteoblasts can be readily assessed using a combination of fluorescent reporter mice and this intramembranous bone marrow injury model. In clinical contexts, intramembranous bone formation is relevant for healing critical size defects, stabilized fractures, cortical defects, trauma from tumor resections, and joint replacements.

Introduction

Long bone injuries heal through the endochondral or intramembranous bone formation pathway. Unlike the endochondral pathway, which requires a cartilage precursor step, bone repair via intramembranous ossification involves direct conversion of skeletal stem and progenitor cells (SSPCS) into bone-forming osteoblasts¹. Clinically, this process is relevant for bone healing in a variety of contexts, including critical size defects, stabilized fractures, cortical defects, distraction osteogenesis, trauma from tumor resections, and osseointegration of joint replacement implants^2,3. Interestingly, in studies of patients with long bone fractures, 66%-82% had displaced fractures, which required fixation devices to stabilize the bone^4,5. These rigidly stabilized bones primarily heal through intramembranous ossification as the broken bone ends come in direct contact with each other^6,7. Yet, the mechanistic regulation of intramembranous bone regeneration remains relatively understudied compared to bone formation via the endochondral pathways. Food and Drug Administration (FDA)-approved therapies to augment bone healing are hindered by limited clinical efficacy and high costs and are associated with significant adverse effects⁸.

Mechanical bone marrow ablation provides a valuable model for studying intramembranous bone formation. This simple injury model allows for studying bone regeneration in the bone marrow without disrupting the cortical bone. Following BM ablation, the healing process involves distinct yet overlapping phases. The initial phase occurs in the first 1 to 5 days and involves clot formation and inflammation, where inflammatory cells and cytokines initiate healing. From days 3 to 14, the second phase is characterized by regeneration, including neovascularization, mesenchymal stem cell (MSC) migration and proliferation, osteoblastic differentiation, and woven bone formation. The final remodeling phase begins approximately 10 days after surgery, with bone tissue undergoing maturation and restructuring until the marrow is fully restored by day 56⁹. This rapid healing timeline makes the BM ablation model ideal for studying early bone repair responses, particularly during the critical periods of inflammation, progenitor cell recruitment, and osteoblast differentiation. Using techniques like histology, flow cytometry, and quantitative µCT analysis, cellular responses and bone formation in early repair stages can be evaluated. Using the aforementioned techniques, the BM ablation model can provide insight into mechanisms of intramembranous bone regeneration and aid in identifying key therapeutic targets for enhancing bone healing.

Protocol

The following procedures were performed with approval from the University of Connecticut Health Center Institutional Animal Care and Use Committee (IACUC). All surgeries were performed under sterile conditions as outlined by the National Institutes of Health (NIH) guidelines. Pain and risk of infections were managed with proper analgesics and antiseptics to ensure a successful outcome.

1. Preparation of surgical area and instruments

1. Prepare the surgical procedure room by disinfecting working surfaces with a clinical-grade disinfectant cleaner.
   NOTE: The surgeon should wear appropriate personal protective equipment (PPE), including sterile disposable gloves, hair bonnet, gown, and mask.
2. Prepare a heating pad covered with a sterile surgical pad. Gather and arrange all sterile instruments and reagents (Table of Materials) with convenient access.

2. Pre-operative anesthesia

1. Weigh all mice before surgery. Calculate the volumes of analgesic to be used based on the available formulation of buprenorphine.
2. Assemble an animal anesthesia unit within the sterile working space, which includes an induction chamber and a nose-cone assembly. Set oxygen output to 2 L min^-1 and isoflurane to 1.5-3% (v/v) depending on mouse age and weight.
3. Once the mouse is sedated, remove the animal from the induction chamber and place it on a sterile surgical pad in the supine position. Apply ophthalmic lubricant to both eyes. Place the nose cone over the head. Check that the mouse is adequately anesthetized by gently pinching the toe. Ensure there is no hindlimb reflex response before proceeding.
4. Prior to surgery, briefly remove the mouse from nose cone to inject the total dose of extended-release buprenorphine via the subcutaneous route; the total dose is 3.25 mg/kg.

3. Surgical procedure description

NOTE: This surgical procedure utilizes C57BL/6 male and female mice aged 10 weeks to 24 weeks. The procedure can be performed on mice as young as 6 weeks up to 33 weeks old and on various inbred strains, as shown by others^10,11. Younger mice may require shorter needle lengths, while older or larger mice may require a longer needle.

1. Fill a 3 mL syringe equipped with a 26 G needle with sterile saline. Place this to the side.
2. Remove the fur from the hindlimb on which procedure will be performed with electric or battery-operated clippers (#40 blade) and remove any loose fur from the surgical field. Using sterile cotton-tipped applicators, treat the surgical site with an iodine-based scrub (10% Povidone-iodine), followed by 70% isopropyl alcohol. Scrub the surgical site starting at the center of the knee and making a circular sweep outward. Allow the site to dry. Repeat the scrub steps 3 times. 
3. Place the mouse on its back and flex the knee of the operative leg. Hold the leg of the mouse in the non-dominant hand such that the knee is bent over the middle finger, the second finger rests on the femur, and the thumb rests on the tibia.
4. Make a small, shallow horizontal incision just below the patella using a scalpel.
   NOTE: This step improves visualization of the insertion site. With experience, this step can be skipped.
5. Locate the joint space by placing the needle horizontally across the bent knee and feeling for the space between the femur and tibia. Using this anatomical site to guide, take a 25 G needle and manually drill (by twisting the needle and simultaneously applying moderate pressure onto the needle) into the distal femur from the knee toward the proximal side. Insert the needle through or around the patella.
6. X-ray the mouse to ensure the needle has been appropriately placed in the medullary cavity, as close to the center as possible (Figure 1).
7. Remove the 25 G needle and immediately insert a 26 G needle into the same hole within the marrow cavity. Ream the sides of the medullary cavity in a circular motion at least 5 times until it feels smooth. Keep the number of reaming motions consistent between each mouse.
   NOTE: Steps 3.5 and 3.7 may cause needlestick injury to the surgeon if not handled carefully.
8. Remove the 26 G needle and replace the 26 G needle connected to the saline syringe. Flush the cavity until the backflow runs clear. This typically requires 1.5-2 mL of saline. Remove the syringe and needle and safely dispose of them in a sharp container.
   NOTE: There should not be resistance when pushing the fluid. If so, re-insert the needle to ensure it is within the cavity. It may be necessary to pull the needle out halfway.
9. Close the skin wound using a topical adhesive suture.

4. Postoperative care

1. Immediately following surgery, administer the remaining half dose of extended-release buprenorphine using dosing from step 2.4.
   NOTE: Extended-release Buprenorphine provides pain relief for up to 72 h.
2. Monitor the animals on the heating pad for signs of normal, unobstructed breathing until they awaken from surgery. Once ambulatory, return mice to their cage.
3. Continue to monitor the mice once per day for 3 days after the surgery to ensure they are healing properly and have full mobility, watching for signs of pain.

5. Processing bones for µCT scanning and analysis

1. Euthanize the mice according to the humane euthanization method regulated and approved by the IACUC or other applicable institutional policy. Here, the primary euthanasia method of CO₂ inhalation was used, followed by a secondary method of cervical dislocation.
2. Harvest the femurs by peeling away the skin around the hindlimbs and removing the entire limb at the hip joint.
3. Sandwich each limb between two sponges in an embedding cassette. Transfer the bones in their cassettes to a 10% formalin solution and fix them at 4 °C for 5-7 days, considering the age and size of the mouse.
   NOTE: Both over-fixation and under-fixation result in poor marrow resolution.
4. After fixation, wash the cassettes in phosphate buffered saline (PBS) for 5 min 3 times.
5. Remove the limbs from the cassettes and trim away muscle using a scalpel as needed.
   NOTE: Depending on downstream applications, rough handling can disrupt the periosteal layer of bone.
6. Separate the femur at the knee by carefully cutting through the ligaments using small surgical scissors or a scalpel.
   NOTE: The tibias can be used for other applications or discarded.
7. Transfer the femurs to 1.7 mL or 15 mL conical tubes into a 70% ethanol solution. Bones can remain in 70% ethanol at 4 °C until ready for µCT scanning.
8. Scan and assess the femurs for trabecular parameters, producing qualitative and quantitative data.

6. Processing bones for frozen sectioning and histology

NOTE: Bones can be processed for frozen sectioning and histology after undergoing µCT scanning. The scanning does not affect fluorescent labeling or subsequent antibody staining.

1. Transfer the bones from 70% ethanol to PBS and wash 3 times, 5 min per wash.
2. Incubate in a series of sucrose solutions from 10%, 20%, to 30% in PBS at 4 °C overnight at each concentration. Store the bones in 30% sucrose for up to 5 days at 4 °C or in this solution at -20 °C for 6-12 months until ready for embedding.
   NOTE: Using a sucrose gradient results in better tissue penetration. After step 6.1, bones may be placed directly into a 30% sucrose solution overnight.
3. Embed bones in optimal cutting temperature (OCT) compound using cryoembedding molds. Store at -80 °C until ready to section.
4. Section embedded bones using the cryofilm tape method with cryostat temperature set between -24 °C and -26 °C.
5. Adhere the sections on the cryofilm to an adhesive slide using optical adhesive and cure it until hardened using an ultraviolet (UV) crosslinker.

7. Processing for flow cytometry

NOTE: This protocol is compatible with a variety of antibodies. For characterizing skeletal stem and progenitor cells, pooling 3-4 bones together is recommended as these are rare cell populations in the bone marrow.

1. Prepare fluorescence-activated cell sorting (FACS) staining media (FSM). To make 1000 mL solution of FSM, combine: 100 mL of Hanks' balanced salt solution (HBSS), 10 mL of 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 20 mL of Fetal bovine serum (FBS), 870 mL of dd H2O. Pass the FSM solution through a 0.2 µm filter.
2. Before harvesting tissue, fill Petri dishes with 4-5 mL of sterile PBS without calcium and magnesium.
3. Prepare the digestion media
   1. For every 4-6 femurs, prepare a digestion mixture in 10 mL of PBS containing 0.005 g of collagenase P and 0.02 g of hyaluronidase. Mix by gentle vortexing and pass the mixture through a 0.22 µm filter.
4. Harvest bones on post-surgery Day 3 as described in step 5.2. Remove all muscle and connective tissue with a scalpel blade.
5. Fill a Petri dish with 5 mL of digestion media. Place 4-6 bones in the digestion media. Chop each bone finely, making the first cut longitudinally and laterally.
6. Transfer tissue to a 14 mL round-bottom tube and seal with parafilm. Cover tubes in aluminum foil.
7. Shake at 37 °C for 20 min by keeping the tube horizontally on the shaker.
8. Remove the tube from the shaker. Cells will be clumped at the bottom. Gently collect the digested material without disturbing the cells at the bottom, and pass the digest through a 70 µm filter. Collect the filtrate in a new 50 mL tube containing 10 mL of cold FSM and keep it on ice.
9. To the cells, add the remaining 5 mL of the digestion mix. Seal again with parafilm and cover with aluminum foil. Shake the tube as described above. Repeat steps 7.7-7.8.
10. After the 20 min incubation at 37 °C, transfer the digest to the same tube containing the digest from step 7.8, passing through a 70 µm filter with another 10 mL of FSM.
11. Centrifuge the suspension at 274 x g at 4 °C. Aspirate the supernatant and collect the cell pellet. Dislodge the pellet by gently tapping the tube.
    NOTE: The centrifugal force (g) is based on a rotor radius of 170 mm. Please check the rotor radius and adjust the speed accordingly.
12. To perform the red blood cell lysis, add 1 mL of ammonium-chloride-potassium (ACK) lysing buffer. Keep all reagents and cells on ice.
13. Incubate the cells in ACK for about 5 min at RT. Do not exceed 5 min to avoid overly lysing the cells.
14. Add 9 mL of FSM. Centrifuge at 274 x g for 5 min at 4 °C.
15. Aspirate the supernatant. Break up the cell pellet by gently raking the tube on a tube rack.
16. Add 10 mL of FSM (volume for 4 bones, can be adjusted based on how many bones are combined) and resuspend.
17. Take an aliquot of each sample and add propidium iodide (PI) as per the manufacturer's instructions. Count live and total cells with a cell counter.
18. Proceed with antibody staining of samples using desired markers for flow cytometry analysis^12,13,14.

Results

The success of the mechanical bone marrow (BM) ablation procedure was validated by X-ray imaging, while the 26G needle was inserted into the medullary cavity (Figure 2A). When X-ray imaging indicated that the needle was inserted at an improper angle or penetrated through the bone, the needle was re-inserted at the proper angle and re-imaged to confirm accurate placement. When a successful position could not be attained, the animal was excluded from analysis. While radiographic two-dimensiona...

Discussion

The goal was to establish a reliable and reproducible method to study the molecular and cellular processes governing bone repair, focusing on the direct conversion of skeletal stem and progenitor cells (SSPCs) into bone-depositing osteoblasts in the bone marrow space/cavity. The mechanical bone marrow ablation method described here is an optimized and validated protocol for bone marrow ablation by mechanical injury, which allows for the investigation of intramembranous bone formation. Several steps in this protocol are c...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
10% Formalin Solution	Sigma-Aldrich	HT501128
25 G 0.625 in Air-tite EZ Flo Hypodermic Needle	Ait Tite/ Fisher Scientific	14-817-239; ref #NEZ25058
26 G 1/2" Disposable Hypodermic Needles	Exel International/ Fisher Scientific	14-840-83; ref #26402	Any 26 G needles may be substituted, must be detachable from syringe
3 mL Sterile Syringe	Fisherbrand/ Fisher Scientific	14-955-457
AB Sterile Pad	McKesson/ Fisher Scientific	NC0593755	Alternatively can use autoclavable absorbant pads
Acetic Acid, Glacial	Millipore Sigma	AX0073	Needs to be diluted to 1%
Ammonium-Chloride-Potassium (ACK) Lysing Buffer	Gibco/ Thermo Fisher Scientific	A1049201	Used for red blood cell lysis in preparation for flow cytometry
Anesthesia Induction Chamber 2 L	VetEquip	941444
Betadine (Providone-Iodine) Solution	Penn Veterinary Supply Inc/ Fisher Scientific	NC0158124
Buprenorphine HCL Injection 0.3 mg/mL	Generic/ Covetrus	59122	Must obtain state and federal DEA license for purchasing controlled substance
Calcein-AM	Sigma Aldrich	206700-1MG
Collagenase P	Roche	11249002001	Used for digesting bone
Cotton Swabs and Applicators	Fisherbrand/ Fisher Scientific	22-363-172
Cryofilm Type II C(10), 3.5 cm	Section Lab, Japan	CFS 105
DAPI Staining Solution	Abcam	ab228549
Digital Gram Scale	Toprime/ Amazon	B06X6LW4V9	Can use any brand scale to weigh mice
Dissecting Forceps	Fisherbrand/ Fisher Scientific	08-953E; 08-953F
Dissecting Scissors	Fisherbrand/ Fisher Scientific	08-940
Endure Low Profile Cryostat/Microtome Blades	Tanner Scientific	TNR315LP
Ethiqa XR Buprenorphine Extended-Release 1.3 mg/mL	Ethiqa XR/ Cevetrus	72117	Must obtain state and federal DEA license for purchasing controlled substance
Fast Green, 0.2%	Electron Miscroscopy Sciences EMS	50-319-57
Fetal Bovine Serum heat inactivated	Gibco/ Thermo Fisher Scientific	A5256801	Used to make FACS Staining Media (FSM)
Fluoromount-G Mounting Solution	SouthernBiotech	0100-01
GLUSEAL Topical Skin Adhesive	AD Surgical	GLU-550
Goat anti-Rabbit Alexa 647	Thermo Fisher	A21244	1:300 dilution
Hanks' Balanced Salt Solution (HBSS)	Gibco/ Thermo Fisher Scientific	14065056	Used to make FACS Staining Media (FSM)
Heating Pad (large)	Boncare/ Amazon	B0CLHSWXSX	Can use any brand heating pad, remove the cloth cover
Hematoxylin	MilliporeSigma/ Fisher Scientific	M1051740500
Hyaluronidase	MilliporeSigma	HX0514	Used for digesting bone
Isoflurane Animal Anesthesia System	VetEquip	901810
KUBTEC Parameter 2D Cabinet X-ray System	OncoMed Solutions	N/A	Any X-ray Cabinet imaging system can be used
Microcentrifuge Tubes	Corning/ Thomas Scientific	8600A52
Microscope Cover Glass, 18 mm x 18 mm, # 1 Thickness	Globe Scientific	1414-10
N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid HEPES (1M)	Gibco/ Thermo Fisher Scientific	156300-80	Used to make FACS Staining Media (FSM)
Norland optical Adhesive NO61	Edmund Optical	37-322	Used to adhere cryofilm sections to a slide
Osterix Rabbit Monoclonal Antibody	Abcam	ab209484	1:500 dilution
Peroxigard Ready to Use One Step Disinfectant	Virox Technologies	29101	Can be substituted with any another clinical/lab grade disinfectant
Propidium Iodide (PI)	Invitrogen/ Thermo Fisher Scientific	P1304MP	Nuclear fluorescent stain used to detect dead cells for vioability counts
Rechargeable Electric Trimmer	Wahl/ Amazon	B001FWXKUE	Any clippers can be used, a smaller sized trimmer will make it easier to shave smaller animals
Rodent Nose Cone	VetEquip	921609
Safranin O, 85% pure, certified	Fisher Scientific	AC419211000
Seal'N Freeze Cryotray Square Molds	Electron Miscroscopy Sciences EMS	62642-01	Used for embedding, other embedding molds can be used. This specific model has a deeper mold, as well as a lid to seal the block
Silver Nitrate	Sigma Aldrich	S6506	Silver nitrate for von Kossa stain
Sterile Phosphate Buffered Saline (PBS)	Gibco/ Thermo Fisher Scientific	10010023	Alternatively, freshly prepared PBS that has been filtered and autoclaved can be used
Sterile Standard Scalpels	Integra Miltex/ Fisher Scientific	12-460-455
Sterile Standard Scalpels	Integra Miltex/ Fisher Scientific	12-460-455
Sucrose Powder 1 KG	Thermo Fisher Scientific	036508-A1
Superfrost Plus Miscroscope Slides	Fisherbrand/ Fisher Scientific	1255015	Adhesive slides
Sure-Tek Tissue Processing/ Embedding Casettes	Fisherbrand Histosette/ Fisher Scientific	22-048-141
Tissue Plus Optimal Cutting Temperature O.C.T Compound	Fisher Scientific	23-730-571
Tissue Sponges	Epredia/ Fisher Scientific	84-53	Tissue sponges protect tissues from overproocessing
Toluidine Blue	Fisher Scientific	AC348600050