A New Method of Bone Stromal Cell Characterization by Flow Cytometry

Abstract

The bone microenvironment cellular composition plays an essential role in bone health and is disrupted in bone pathologies, such as osteoporosis, osteoarthritis, and cancer. Flow cytometry protocols for hematopoietic stem cell lineages are well defined and well established. Additionally, a consensus for mesenchymal stem cell flow markers has been developed. However, flow cytometry markers for bone-residing cells– osteoblasts, osteoclasts, and osteocytes– have not been proposed. Here, we describe a novel partial digestion method to separate these cells from the bone matrix and present new markers for enumerating these cells by flow cytometry. We optimized bone digestion and analyzed markers across murine, non-human primate, and human bone. The isolation and staining protocols can be used with either cell sorting or flow cytometry. Our method allows for the enumeration and collection of hematopoietic and mesenchymal lineage cells in the bone microenvironment combined with bone-residing stromal cells. Thus, we have established a multi-fluorochrome bone marrow cell-typing methodology.

Basic Protocol 1: Partial Digestion for Murine Long Bone Stromal Cell Isolation

Alternate Protocol 1: Partial Digestion for Primate Vertebrae Stromal Cell Isolation

Alternate Protocol 2: Murine Vertebrae Crushing for Bone Stromal Cell Isolation

Basic Protocol 2: Staining of Bone Stromal Cells

Support Protocol 1: Fluorescence Minus One Control, Isotype Control, and Antibody Titration Protocol

Basic Protocol 3: Cell Sorting of Bone Stromal Cells

Alternate Protocol 3: Flow Cytometry Analysis of Bone Stromal Cells

Support Protocol 2: Preparing Compensation Beads

INTRODUCTION:

The composition of the bone microenvironment plays an important role in development and pathologies, with alterations in the marrow and stromal cell composition having profound effects on the bone structure. A balance between cells of the hematopoietic stem cell lineage, including osteoclasts, and mesenchymal stem cell lineage cells, including osteoblasts, is required for bone homeostasis. Flow cytometry and cell sorting are the gold standard methods for the enumeration and sorting of hematopoietic stem cell lineage cells. Additionally, a consensus for mesenchymal stem cell flow cytometry markers is established for these progenitor cells in the bone marrow (Dominici et al., 2006). However, flow cytometry markers for bone-residing stromal cells- osteoblasts, osteoclasts, and osteocytes- have not been established. This is partly to the difficulty in removing these cells from the bone matrix for analysis. In Basic Protocol 1, an optimized partial collagenase digestion protocol to isolate bone marrow stromal cells is described. Further, in Alternate Protocols 1 and 2, isolation of bone marrow stromal cells by partial digestion of primate vertebrae or grinding from murine vertebrae is discussed. Basic Protocol 2 describes the staining of isolated bone marrow stromal cells to identify hematopoietic stem cells, mesenchymal stem cells, osteoblasts, osteoclasts, and osteocytes using the marker panel shown in Figure 1. Finally, in Basic Protocol 3 and Alternate Protocol 3, the analysis of bone stromal cells by cell sorting and flow cytometry are described. This protocol was optimized for use with common three laser cell sorters or two laser flow cytometers using the brightest and most stable fluorophores available for each antibody. This method allows for the enumeration and, using cell sorting, isolation of hematopoietic stem cells, mesenchymal stem cells, and bone-residing cells– osteoblasts, osteoclasts, and osteocytes– in a single experiment. This method will thus profile a majority of the bone microenvironment cells. It can be used for assessing the cellular composition of the bone microenvironment during development and examining the influence of various pathologies on the microenvironment.

Figure 1. Staining Scheme for Bone Marrow and Residing Stromal Cells.

Based upon existing literature, a panel of markers was developed to identify and differentiate bone marrow stromal cells of the hematopoietic and mesenchymal lineages. Created with Biorender.com.

STRATEGIC PLANNING:

Choosing Analysis Method

Before starting experiments, determine whether flow cytometry (cell enumeration) or cell sorting (cell enumeration and separation) will provide the information needed for your experiments. If you need to perform downstream analyses on bone-residing stromal cells, cell sorting will be a better choice. Flow cytometry can be performed on fixed cells and has several potential storage/stoppage points. Cell sorting can only be completed on unfixed cells and must be completed within a few days of isolating cells. Please check the Time Considerations carefully to reserve time on a cell sorting machine.

Choice of Antibody Combinations

The selection of antibodies is essential to identify the proper cell population, especially for the rare bone-residing cells. For this protocol, multi-fluorophore panels were developed for two or three laser analysis machines using commercially available antibodies with brighter fluorophores assigned to rarer cell surface proteins. Markers for hematopoietic stem cells and mesenchymal stem cells are well defined (Dominici et al., 2006; Boxall and Jones, 2012). Due to fluorophore availability, not all the markers were tested in this protocol but could be used in multi-fluorophore panels with additional lasers for excitation. Antibody selection for bone-residing osteoclasts, osteoblasts, and osteocytes focused on surface marker expression. Osteoclast surface marker RANKL/CD254 (receptor activator of nuclear factor-κB ligand) or RANK/CD265 is associated with differentiation and activity of osteoclasts (Boyle et al., 1999). Thus, CD265 expression on cells expressing the macrophage surface markers CD68 and CD115 was postulated to represent osteoclasts or preosteoclast cells. Osteoblast surface proteins were proposed to be CD90/Thy-1 and alkaline phosphatase based on their roles in bone formation and bone matrix calcification (Trentz et al., 2010; Rutkovskiy et al., 2016; Picke et al., 2018). Osteocyte markers SPARC (secreted protein acidic and rich in cysteine) and Podoplanin (GP38) were derived from publications describing osteocyte ontogeny from osteoblasts (Bonewald, 2011; Zhu et al., 2011). Markers were chosen based on their expression on the cell surface, their ability to differentiate between bone stromal cell populations, and the commercial availability of fluorophore-conjugated antibodies. When designing panels, look for antibodies in bright fluorophores with minimal overlap. Additional markers can be included as more fluorophore conjugations become available and for use with analyzers with additional excitation lasers.

BASIC PROTOCOL 1

Partial Digestion for Murine Long Bone Stromal Cell Isolation

This protocol describes the partial digestion of murine long bones by collagenase I. While hindlimbs are often used, the procedure is the same for forelimbs, and these can be combined to maximize cell yield. This digestion method results in 3 – 6×10⁶ cells released from long bones per animal (See Figure 2 and Table 1). Animal care and euthanasia should be performed under an institutionally approved protocol.

Figure 2. Murine Digestion Yields across Age, Sex, and Genotype.

For this study, male and female Balb/c mice aged 8 weeks and C57Bl/6 mice aged 18 or 28 weeks were purchased to optimize the protocol across strains, sex, and age under an approved Wake Forest School of Medicine IACUC protocol. Long bones were isolated from mice and processed as described in Basic Protocol 1. The numbers of cells recovered from 3–5 individual animals are represented as mean±SEM.

Table 1.

Murine Long Bones and Vertebrae Digestion Yields

Tissue	Live Cell Yield (107 cells)	Percent Viable
Long Bones	11.9	85
Vertebrae	16.4	76

Spines, forelimbs, and hindlimbs were isolated from five male mice aged 28 weeks, and samples were pooled after digestion.

Materials:

• 1X PBS containing 100 U/mL Penicillin and 100 μg/mL Streptomycin (PBS+P/S)

• 1 mg/mL Collagenase I in PBS+P/S (Gibco, 17-100-017)

• 1% methanol-free formaldehyde in PBS (diluted from 10% Ultra Pure, methanol-free formaldehyde, Polysciences Inc, 04018–1)

• Flow Cytometry Staining Media (see recipe in Reagents and Solutions)

• Laminar chemical hood (Kewaunee Scientific Airflow Thin Wall Fume hood, or equivalent)

• Laminar biosafety cabinet (Baker Company SterilGard Hood Class II, or equivalent)

• Centrifuges (Eppendorf AG 5453 and Eppendorf 5810)

• Shaking incubator (Barnstead MaxQ 4000, or equivalent)

• No. 10 Safety Scalpel (Fisher, Futura SMS210)

• Surgical scissors

• Forceps

• 27 ½ G needles (Medlab Gear, BD305109)

• 15 mL and 50 mL conical tubes

• 40 μm cell strainer (Denville, 1006202)

• 10 mL syringe (Fisher, 1482919C)

• Pipettor (Gilson pipetman P2L, P20L, P200L and P1000L)

• Pipettes (10 mL: Fisher, 13-678-11E, 5 mL: Fisher, 13-678-11D)

• Pipette tips (VWR, BT1250 (1 mL), BT200 (200μL), BT20 (20 μL))

• 100 mm dishes (Corning 353003)

1. Sacrifice mice according to an approved animal protocol.

2. Remove skin from the leg by cutting down the medial side and then removing skin with fingers to expose the entire limb musculature.

3. Remove hindlimb at hip joint and place in PBS+P/S on ice.

   Whole limbs can be stored at 4°C for one week with cell viability remaining 85–90%. For long-term storage, remove muscle, flash freeze the bones, and store at −20°C until ready to use. Freezing legs drops cell viability to 20–30%.

4. In a biosafety or chemical hood with the light off, prepare a 10 mL syringe with PBS+P/S and attach a 27 ½ G needle.

   A biosafety hood is recommended for tissues containing biohazards, producing aerosols, or when sterility is a concern and cells will be recovered after sorting. A chemical hood can be used when sterility and biohazards are not concerns.

5. Place bones in a 100 mm dish. Remove extra musculature and the foot with small surgical scissors and a scalpel using a 100 mm dish lid as a workspace. Cut off the proximal and distal ends of the long bones (tibia, femur, humerus).

6. Insert syringe containing PBS+EDTA into the marrow cavity and flush the diaphysis with 2–3 mL of PBS+EDTA into a 50 mL tube. Keep flushed bone marrow on ice.

   Repeat for all bones of a single genotype. Use a new syringe for each different genotype or experimental animal group.

7. Take the flushed bone and holding with forceps, cut compact and trabecular bone into 1–3 mm chips with scissors.

   Cut bones over a 15 mL conical tube to collect pieces and minimize debris.

8. Transfer bone chips into a 50 mL conical tube containing 1–3 mL of 1 mg/mL collagenase I in PBS+P/S.

9. Incubate 30 minutes to an hour at 37°C in a shaking incubator at 200 rpm (0.04 g).

   The expected cell yield and viability are shown in Table 1. Figure 2 demonstrates yields from murine hindlimbs across ages, sexes, and genotypes.

10. Combine partially digested bone chips with bone marrow extract. Add PBS+EDTA to a final volume of 30 mL.

11. Mix the cell suspension vigorously by pipetting 20 times or vortexing.

12. Filter the cell suspension through a 40 μm cell strainer into a new 50 mL conical tube.

13. Centrifuge cell suspension at 175 g for 5 minutes at room temperature.

14. Discard the supernatant and resuspend in Flow Cytometry Staining Media to an approximate concentration of 1 million cells per mL. Proceed to Basic Protocol 2 to stain isolated cells.

    If not staining immediately, store at 4°C overnight. For long-term storage, store samples in 1% methanol-free formaldehyde in PBS for up to two weeks for flow cytometry. Cells can be fixed in 1% methanol-free formaldehyde in PBS before or after staining for flow cytometry. Unfixed cells are required for cell sorting.

ALTERNATE PROTOCOL 1

Partial Digestion for Primate Vertebrae Stromal Cell Isolation

This protocol describes the isolation of bone marrow stromal cells from the vertebrae of non-human primates or humans. Below, the protocol begins with the isolation from whole spines, but individual vertebrae can also be processed starting from step 6. While not tested, the trabecular regions from other primate bones could be used with this protocol with additional optimization of incubation times.

CAUTION: Tissues from some species of non-human primates are a Biosafety Level 2 (BSL-2) pathogen due to the risk of Herpes B Virus. Follow all local guidelines and regulations for the use and handling of potentially pathogenic tissues.

Additional Materials:

• Sterile water

• αMEM media containing 100 U/mL Penicillin and 100 μg/mL Streptomycin (αMEM+P/S)

• 3 mg/mL collagenase I in αMEM+P/S

• Balance

• Dremel 300 with Dremel 409 cut off wheel blades

• Mopec 810 autopsy saw

• Cut-resistant gloves

• Nitrile gloves

• Safety goggles

• Blue absorbent pads

• Water bottle for irrigation

• 70 μm cell strainer (Fisher, 08-771-2)

• Large plastic tube for thawing spines

1. Retrieve wrapped spine samples from −20°C freezer and thaw for 36 hours in a tub at 4°C.

   If samples remain partially frozen, thaw wrapped spines in room temperature to slightly warm water until thawed completely.

2. In a biosafety hood, unwrap spines and, using a scalpel, remove musculature and ligaments from one end of the spine until the intervertebral disc is revealed.

   A biosafety hood is recommended due to the potential for aerosolization and biohazards associated with primate tissues.

3. Using a bone saw, Dremel, or scalpel, remove the vertebrae from the remainder of the spine.

   If using a power tool, irrigate the area with water to prevent aerosolization of the bone. Re-wrap remaining spine in gauze covering the bone and keep on ice until ready to return to freezer.

4. Using a Dremel and a bone saw, remove the spinal processes from the vertebrae.

5. Using a Dremel and a scalpel, remove intervertebral discs from both sides of the vertebrae.

6. Using a Dremel and a bone saw, cut a wedge into the vertebral body containing both cortical and trabecular bone and representing approximately 1/6^th of the tissue mass. Further, subdivide this wedge into 8 to 10 small pieces to expose as much surface area as possible for digestion.

   Additionally, cores can be drilled from the vertebrae and will provide equivalent results.

7. Weigh the samples and place them in 50 mL conical tubes with 10 mL 3 mg/mL collagenase I in αMEM+P/S.

8. Place the sample in a 37°C shaking incubator overnight at 200 rpm (0.04 g).

   Optimization of incubation time is shown in Table 2. The expected cell yield and viability are shown in Table 3 for baboons and in Table 4 for humans.

9. Strain the sample through a 70 μm cell strainer and centrifuge at 122 g for 3–5 minutes.

   If there is additional debris, resuspend cells and pass through the 70 μm cell strainer again.

10. Resuspend cells in 2 mL of Flow Cytometry Staining Media and proceed to Basic Protocol 2 to stain isolated cells.

    If not staining immediately, store at 4°C overnight. For long-term storage, store samples in 1% methanol-free formaldehyde in PBS for up to two weeks for flow cytometry.

Table 2.

Optimization of Collagenase I Partial Digestion of Bone to Release Bone Stromal Cells from Non-Human Primates.

Time	Initial Weight (g)	Extracted Cell Concentration (105 cells/mL)
4 hours	2.88	2.19
6 hours	2.87	3.17
8 hours	2.13	7.06
22 hours	2.76	7.94

Archival female 10-year-old baboon vertebrae digested with 3 mg/mL collagenase I for the indicated times at 37°C, shaking.

Table 3.

Non-Human Primate Vertebrae Digestion Yields

Animal Pooled Samples	Initial Weight (g)	Extracted Cell Yield (107 cells)	Live Cell Yield (107 cells)	Percent Viable
B1	21.927	94.7	30	31.7
B2	5.934	4.87	1.04	21.4
B3	6.790	2.74	0.985	35.9

Each pool contains two female and one male baboon archival vertebrae with an average age of 19.6 in each pool. Ages range from 18 to 22 years across all baboons in the sample.

Table 4.

Human Vertebrae Digestion Yields

Subject Pooled Samples	Live Cell Yield (107 cells)	Percent Viable
H1	2.42	21
H2	10.8	37
H3	6.1	27

Each pool contains archival samples from 4 people with 2 males and 2 females in pools H1 and H3. Ages across the samples range from 64 to 92 years old. The pools average 81, 89, and 80 years respectively.

ALTERNATE PROTOCOL 2

Murine Vertebrae Grinding for Bone Stromal Cell Isolation

This protocol is used to isolate bone marrow-residing stromal cells from murine spines. Compared to bone marrow isolation methods using femurs and tibias (Basic Protocol 1), this procedure allows for a much greater yield of cells (Table 1). When sacrificing mice, avoid performing a cervical dislocation to prevent potential contamination of spinal bone marrow cells.

Additional Materials:

• 3 mg/mL collagenase I in αMEM+P/S

• UV sterilized mortar and pestle

• Alcohol swaps

• Bone scissors

1. Sacrifice mice according to an approved animal protocol.

   Do not use cervical dislocation, as this can injure the spine.

2. Open the peritoneal cavity and remove organs to visualize the spine. Cut on both sides of the spine from the base of the tail to the neck, making sure to cut through attached ribs.

3. Use a scalpel or scissors to clean off any extra musculature and bones.

   The spinal process and spine will be attached. When using spines from aged mice, be extra gentle as vertebrae are fragile.

4. Remove the ends of the spine near the brainstem and tail.

   Spines can be stored in cold PBS in 50 mL conical tubes at 4°C for up to 1 week.

5. Cut the remaining spine into three equal pieces using bone scissors.

6. Holding a spine piece vertically with forceps, use scissors to cut the spine open. Once open, move scissors into a horizontal position and open spreading apart the sides of the spine to expose and then remove the spinal cord.

7. When the spinal cords are removed from all three pieces, place the remaining spine into a mortar with 10 mL of cold PBS. Use a pestle to crush pieces until the liquid becomes pink.

8. Pipette fluid from mortar through 40 μm cell strainer into a 50 mL conical tube.

9. Repeat steps 7 and 8 until the PBS remains clear after crushing.

   The expected cell yield and viability are shown in Table 1.

10. Centrifuge cell suspension at 274 g for 5 minutes.

11. Discard the supernatant and resuspend in Flow Cytometry Staining Media to an approximate concentration of 1 million cells per mL. Proceed to Basic Protocol 2 to stain isolated cells.

    If not staining immediately, store at 4°C overnight. For long-term storage, store samples in 1% methanol-free formaldehyde in PBS for up to two weeks. Cells can be fixed in 1% methanol-free formaldehyde in PBS before or after staining for flow cytometry analysis. Unfixed cells are required for cell sorting.

BASIC PROTOCOL 2:

Staining of Bone Stromal Cells

This protocol describes the multi-fluorochrome staining for bone-residing stromal cells according to the schema in Figure 1. After staining, cells can be analyzed by cell sorting (Basic Protocol 3) or flow cytometry (Alternate Protocol 3).

Materials:

• PBS+P/S

• Trypan Blue

• Flow Cytometry Staining Media (see recipe in Reagents and Solutions)

• Cell Sorting Buffer (see recipe in Reagents and Solutions)

• Antibodies of interest directly conjugated to fluorochromes (see Tables 5 (murine) and 6 (primate) for antibodies and clones, as well as potential isotype control antibodies in Tables 8 (murine) and 9 (primate))

• FcR blocking solution (BioLegend TruStain FcX PLUS, 422302 (primate) or Miltenyi Biotec, 130-092-575 (murine))

• Live/Dead Stain (Zombie Aqua, BioLegend, 423102, or equivalent)

• 1% methanol-free formaldehyde in PBS

• Cell Counter and slides (BioRad TC20 or equivalent) or hemocytometer

• Microcentrifuge tubes (VWR C-3260)

• Round bottom 96 well plate (Falcon, 3910)

• Flow cytometry tubes (Polypropylene Microtiter Tubes, Fisher 02-681-385, or Falcon5mL Polystyrene Round Bottom Tube, Corning 352054, or similar)

• Cell sorting tubes (Falcon 352052)

• Multichannel Pipettes

• Pipettes (Gilson Pipetman P1000, P200, P20)

• Pipette tips (VWR, BT1250 (1 mL), BT200 (200μL), BT20 (20 μL))

Table 5.

Staining antibodies for murine bone stromal cells

Antigen	Isotype Control	Company	Catalog Number	RRID
MESENCHYMAL STEM CELL
FITC SCA1 (LY-6A/E)	FITC Rat IgG2a, κ	BioLegend	108106	AB_313343
PE CD146	PE Rat IgG2a, κ	BioLegend	134704	AB_2143527
APC CD29	APC Hamster IgG	BioLegend	102214	AB_492831
PerCP-Cy5.5 CD90	PerCP-Cy5.5 Rat IgG2b, κ	BioLegend	105338	AB_2571945
MACROPHAGE/ OSTEOCLAST
FITC CD11B	FITC Rat IgG2b, κ	BioLegend	101206	AB_312789
APC CD115	APC Rat IgG2a, κ	BioLegend	135510	AB_2085221
PerCP-Cy5.5 CD68	PerCP-Cy5.5 Rat IgG2b, κ	BioLegend	137010	AB_2260046
PE CD265 (RANK)	PE Rat IgG2a, κ	BioLegend	119806	AB_2205353
OSTEOBLAST
APC Alkaline Phosphatase	APC Mouse IgG1, κ	R&D Systems	FAB1448A	AB_357039
PerCP-Cy5.5 CD90	PerCP-Cy5.5 Rat IgG2b, κ	BioLegend	105338	AB_2571945
OSTEOCYTE
PE Podoplanin (GP38)	PE Hamster IgG	BioLegend	127408	AB_2161929
APC SPARC	APC Rat IgG2b, κ	R&D Systems	IC942R	AB_2814696
HEMATOPOIETIC STEM CELL
PE CD34	PE Rat IgG2a, κ	BioLegend	152204	AB_2629648
FITC Sca1 (Ly-6A/E)	FITC Rat IgG2a, κ	BioLegend	108106	AB_313343

Table 8.

Fluorescence minus one for primate fluorescence active cell sorting of bone stromal cells.

Antigen	FITC	PE	PerCP-Cy5.5	APC
MESENCHYMAL STEM CELLS
CD146 FMO	CD90	---	---	CD29
CD90 FMO	---	CD146	---	CD29
CD29 FMO	CD90	CD146	---	---
MACROPHAGE/ OSTEOCLAST
CD11b FMO	---	CD254	CD68	CD115
CD254 (RANKL) FMO	CD11b	---	CD68	CD115
CD68 FMO	CD11b	CD254	---	CD115
CD115 FMO	CD11b	CD254	CD68	---
OSTEOBLAST
Alkaline Phosphatase FMO	---	---	CD90	---
CD90 FMO	---	---	---	AlkPhos
OSTEOCYTE
SPARC FMO	---	GP38	---	---
Podoplanin GP38) FMO	---	---	---	SPARC
HEMATOPOIETIC STEM CELLS
CD34 FMO	---	---	---	CD45
CD45 FMO	CD34	---	---	---

Fluorescence minus one (FMO) control panel was conducted to identify fluorochrome bleed over with cell-specific antibody mixes. Results indicated no fluorochrome bleed over into the FMO channel. AlkPhos = Alkaline Phosphatase; SPARC = Secreted Protein Acidic and Rich in Cysteine

• 1.Bring the volume of fresh or fixed cells to 10 mL with PBS+P/S and mix thoroughly. Take a 10 μL sample, combine with an equal volume of trypan blue, and count cells with an automated cell counter (or using hemocytometer).

  Cells need to be diluted to 10 × 10⁶ cells per mL for the cell counter to count cells effectively. Dilute cells before counting as needed to ensure an accurate count.

• 2.Remove 1 × 10⁶ cells per planned staining group and centrifuge 122 g for 3–5 minutes.

  Samples include an unstained control and each staining group using the multicolor antibody groups in Table 5 for murine samples and Table 6 for primate samples. Unstained controls are needed to determine the negative population’s location for gating and to assess autofluorescence (described in Basic Protocols 3 and 4). For initial optimization, prepare staining group samples for FMO controls, isotype controls, and antibody titration as described in Support Protocol 1.

• 3.Resuspend cells in PBS at 1 × 10⁷ cells per mL.
• 4.For live/dead staining: Incubate cells with 1 μL live/dead staining solution per 1 mL cell solution for 20 minutes at room temperature. Wash cells and resuspend in Flow Cytometry Staining Media at 1 × 10⁷ cells per mL.

  This step can only be performed on unfixed cells.

• 5.Block samples with 5 μL appropriate FcR blocking solution per 10⁷ cells for 30 min on ice.
• 6.Aliquot 100 μL of samples into each staining group, including unstained control.

  Staining of samples can be performed in microfuge tubes. If more than 20 samples are prepared, it may be helpful to use a round bottom 96 well plates, prepare antibody master mixes, and use a multichannel pipette to mix samples. Also, prepare a tube or well for isotype controls if using. For sorting, higher volumes of cells will be used to increase the number of cells recovered.

• 7.Stain with primary antibody for 30 min on ice in the dark.

  Recommended multicolor antibody groups are shown in Table 5 for murine samples and Table 6 for primate samples. If secondary antibody staining is required, wash cells, centrifuge at 175 g for 5 minutes, and repeat steps 7 and 8 with the secondary antibody.

• 8.Wash twice in Flow Cytometry Staining Media, centrifuging at 175 g for 5 min.
• 9a.For cell sorting (Basic Protocol 3), cells should be resuspended in a flow cytometry tube containing 100 μL Cell Sorting Buffer.
• 9b.For flow cytometry (Alternate Protocol 3), resuspend sample in 100 μL 1% methanol-free formaldehyde in PBS (1 × 10⁶ cells per 100 μL) while mixing to prevent clumping and move to flow cytometry tube for flow cytometry.
• 10.Store covered in foil at 4°C until ready to run cell sorting (Basic Protocol 3) or flow cytometry analysis (Alternate Protocol 3).

  Samples can be stored for up to 24 hours before cell sorting. Samples can be stored for up to a week before flow cytometry analysis. Samples stained with the PerCP fluorophore will degrade the quickest.

Table 6.

Staining antibodies for primate bone stromal cells.

Antigen	Company	Catalog Number	RRID
MESENCHYMAL STEM CELL
APC CD29	BioLegend	303018	AB_2130080
PE CD146	BioLegend	361006	AB_2562981
FITC CD90	BioLegend	328108	AB_893429
MACROPHAGE/ OSTEOCLAST
PE RANKL (CD254)	BioLegend	347504	AB_2256264
FITC CD11b	BioLegend	101206	AB_312789
PerCP-Cy5.5 CD68	BioLegend	333814	AB_10683171
APC CD115	BioLegend	347306	AB_2562441
OSTEOBLAST
APC Alkaline Phosphatase	R&D Systems	FAB1448A	AB_357039
FITC CD90	BioLegend	328108	AB_893429
OSTEOCYTE
FITC SPARC	R&D Systems	IC941G	AB_2827438
PE Podoplanin (GP38)	BioLegend	337004	AB_1595457
HEMATOPOIETIC STEM CELL
FITC CD34	BioLegend	343504	AB_1731852
APC CD45	BioLegend	304012	AB_314400

SUPPORT PROTOCOL 1

Fluorescence Minus One Control, Isotype Control, and Antibody Titration Protocol

This protocol describes controls that can be used to compensate for spectral overlap and to improve staining. Single color compensation controls are run with each experiment as well as an unstained control to determine autofluorescence. Antibody titration is performed to determine the optimal staining concentration of antibodies for high signal to noise peak expression while retaining identification of cell markers prior to collecting experimental data. Fluorescence minus one controls (FMO) are conducted to assess the proper gating of positive cell populations due to the spread of the data along the log axis at the optimal antibody concentration for each multi-fluorophore staining group (Roederer, 2001). If there is a spectral spread between channels that is not compensated by the single color stained beads, then FMOs must be run every time. FMO analysis should be performed on fixed and unfixed cells representative of the cells being analyzed in experiments. Isotype controls, while controversial (Herzenberg et al., 2006; Maecker and Trotter, 2006; Hulspas et al., 2009), can be run for each experimental group (animal age, genotype, subject, etc.) to serve as controls for non-specific staining, which can be a problem for rare cell populations. Isotype control antibodies must be at the same final concentration and fluorophore to protein ratio as the experimental antibody.

Additional Materials:

• Isotype antibodies (see Table 9 for murine samples and Table 10 for primate samples)

Table 9.

Isotype control murine antibody staining.

Isotype Control	Company	Catalog Number	RRID
MESENCHYMAL STEM CELL
FITC Rat IgG2a, k	BioLegend	400506	AB_273619
PE Rat IgG2a, k	BioLegend	400508	AB_326530
APC Hamster IgG	BioLegend	402012	AB_2814701
PerCP-Cy5.5 Rat IgG2b, k	BioLegend	400632	AB_893677
MACROPHAGE/ OSTEOCLAST
FITC Rat IgG2b, k	BioLegend	400634	AB_893662
APC Rat IgG2a, k	BioLegend	400512	AB_2814702
PerCP-Cy5.5 Rat IgG2b, k	BioLegend	400632	AB_893677
PE Rat IgG2a, k	BioLegend	400508	AB_326530
OSTEOBLAST
APC Rat IgG1, k	BD Biosciences	555751	AB_398613
PerCP-Cy5.5 Rat IgG2b, k	BioLegend	400632	AB_893677
OSTEOCYTE
PE Hamster IgG	BioLegend	400923	AB_2814703
APC Rat IgG2b, k	BioLegend	400612	AB_326556
HEMATOPOIETIC STEM CELL
PE Rat IgG2a, k	BioLegend	400508	AB_326530
FITC Rat IgG2a, k	BioLegend	400506	AB_273619

Isotype control mixes were utilized as a negative control to determine the contribution of non-specific background to staining and correct for non-specific binding. Results indicated no non-specific binding for cell identification schema.

Table 10.

Isotype control primate antibody staining.

Isotype Control	Company	Catalog Number	RRID
MESENCHYMAL STEM CELL
APC Mouse IgG1, κ	BioLegend	400120	AB_2888687
PE Mouse IgG1, κ	BioLegend	400114	AB_326435
FITC Mouse IgG1, κ	BioLegend	400110	AB_2861401
MACROPHAGE/ OSTEOCLAST
PE Mouse IgG2b, κ	BioLegend	400314	N.A
FITC Rat IgG2b, κ	BioLegend	400634	AB_893662
PerCP-Cy5.5 Mouse IgG 2b,k	BioLegend	400338	N.A.
APC Rat IgG1, κ	BioLegend	400412	AB_326518
OSTEOBLAST
APC Mouse IgG1, κ	BioLegend	400120	AB_2888687
FITC Mouse IgG1, κ	BioLegend	400110	AB_2861401
OSTEOCYTE
FITC Mouse IgG1, κ	BioLegend	400110	AB_2861401
PE Rat IgG2a, κ	BioLegend	400508	AB_326530
HEMATOPOIETIC STEM CELL
FITC Mouse IgG1, κ	BioLegend	400110	AB_2861401
APC Mouse IgG1, κ	BioLegend	400120	AB_2888687

Isotype control mixes were utilized as a negative control to determine the contribution of non-specific background to staining and correct for non-specific binding. Results indicated no non-specific binding for cell-identification schema. N.A. represents not available.

1. Follow Basic Protocol 2 to prepare and stain cells. Add groups for FMO analysis, Isotype Controls, and Antibody Titration.

2. For Antibody titration, prepare dilutions of the antibodies described in Table 5 for murine samples and Table 6 for primate samples at 25%, 50%, 100%, and 200% of manufacturer-recommended volume.

   This step should be done as optimization prior to running samples to generate experimental data. Once approximate concentrations are chosen, dilutions within a smaller range can be tested.

3. For FMO controls, set up staining groups as shown in Table 7 for murine samples and Table 8 for primate samples.

   The goal is to create mixes for each staining group, excluding a single antibody. These samples are used to check for fluorescence bleed into other channels. This step should be done as optimization prior to running samples to generate data.

4. For Isotype controls, set up staining groups as shown in Table 9 for murine samples and Table 10 for primate samples.

   Run isotype controls with each new experimental cohort. Isotype antibodies should be used at the same concentration as the experimental antibody which they are controlling. Most manufacturers will recommend a matched isotype control antibody for each experimental antibody.

Table 7.

Fluorescence minus one for murine fluorescence active cell sorting of bone stromal cells.

Antigen	FITC	PE	PerCP-Cy5.5	APC
MESENCHYMAL STEM CELLS
Sca1 (Ly6A/E) FMO	---	CD146	CD90	CD29
CD146 FMO	Sca1	---	CD90	CD29
CD90 FMO	Sca1	CD146	---	CD29
CD29 FMO	Sca1	CD146	CD90	---
MACROPHAGE/ OSTEOCLAST
CD11b FMO	---	CD265	CD68	CD115
CD265 (RANK) FMO	CD11b	---	CD68	CD115
CD68 FMO	CD11b	CD265	---	CD115
CD115 FMO	CD11b	CD265	CD68	---
OSTEOBLAST
Alkaline Phosphatase FMO	---	---	CD90	---
CD90 FMO	---	---	---	AlkPhos
OSTEOCYTE
SPARC FMO	---	GP38	---	---
Podoplanin GP38) FMO	---	---	---	SPARC
HEMATOPOIETIC STEM CELLS
CD34 FMO	Sca1	---	---	---
Sca1 (Ly6A/E) FMO	---	CD34	---	---

Fluorescence minus one (FMO) control panel was conducted to identify fluorochrome bleed over with cell-specific antibody mixes. Results indicated no fluorochrome bleed over into the FMO channel. AlkPhos = Alkaline Phosphatase; SPARC = Secreted Protein Acidic and Rich in Cysteine

BASIC PROTOCOL 3:

Cell Sorting of Bone Stromal Cells

This protocol describes the sorting of stained, isolated bone stromal cells for downstream analyses. At the end of the experiment, individual hematopoietic lineage and mesenchymal lineage cells will be separated into tubes. The sorting report will include the number of cells isolated and percentages of stained cells.

Materials:

• Stained composition beads (see Support Protocol 2)

• Fluorescence-activated cell sorter machine (BC AstriosEQ, BD FACS Aria, or equivalent)

• 1.Take samples (from Basic Protocol 2) and prepared compensation beads (from Support Protocol 2) on ice to the cell sorting facility.
• 2.Prepare collection tubes with 2 mL Cell Sorting Buffer.
• 3.Prepare side scatter (SSC) and forward scatter (FSC) gates based on unstained controls. PMT voltages are set according to procedures outlined in (Maecker and Trotter, 2006) ensuring that weakly stained cells will be separated from electronic noise and ensuring that all positive signals are on scale.

  The gates will be set to exclude the unstained control’s background autofluorescence and locate the negative population.

• 4.Run compensation beads and live/dead controls to generate a compensation matrix. Also, use beads to adjust FSC and SSC.

  Name all the parameters for consistency between experiments. A spillover spreading matrix (SSM) 0–5 is excellent, 6–9 acceptable, and above 10 voltage gains will need to be adjusted. For positive and negative separation, 3–4 median channel logs are desired between the peaks, but fewer median channel logs may be used for markers with lower expression.

• 5.Run a single-stained bead sample to ensure positive peaks are on scale.

  Do not record data until all beads are reviewed and on the scale.

• 6.Rerun single-stained bead samples to perform compensation set up and record files for compensation controls.
• 7.Set a gate around the SSC-H vs. FSC-H scatter to remove debris.
• 8.Set a gate around the singlet population using FSC-H vs. FSC-A and use it for further analysis (Murine: Figure 3A, Primate: Figure 4A). This will remove any doublets.
• 9.Gate live, Zombie Aqua (AmCyan channel) negative cells (Zombie Aqua-A vs. SSC-A; Murine: Figure 3B, Primate: Figure 4B). This cell population will be used in all the following bone-residing cell analyses.

  For hematopoietic stem cells

• 10a.Within the live, singlet population gated above, for murine samples, gate on the Sca1 high and CD34 high population (Figure 3C). For primate samples, gate the CD45 high and CD34 high population (Figure 4C). These markers will identify a broad range of hematopoietic stem cells.

  For mesenchymal stem cells

• 10b.Within the live, singlet population gated above, for murine samples, gate Sca1 high and CD29 high population and then gate the CD90 high and CD146 high population (Figure 3D). For primate samples, within the live, singlet population gated above, gate on CD146 high and CD90 high population and then gate the CD146 high and CD29 high population (Figure 4D). CD146 is considered a definitive marker for mesenchymal stem cells (Correa et al., 2016), while Sca1 is found on multiple murine progenitor cell populations.

  For osteoblasts

• 10c.Within the live, singlet population gated above, gate on the alkaline phosphatase high and CD90 high population (Murine: Figure 3E, Primate: Figure 4E). Only mature osteoblasts express alkaline phosphatase.

  For osteocytes

• 10d.Within the live, singlet population gated above, gate on the SPARC high and podoplanin high population (Murine: Figure 3F, Primate: Figure 4F).

  For macrophages

• 10e.Within the live, singlet population gated above, for murine samples, gate on the CD11b high and CD68 high population followed by the CD265 low and CD115 low (Figure 3G). For primate samples, gate on the CD11b high and CD68 high population followed by the CD254 low and CD115 low (Figure 4G). CD265/RANK or CD254/RANKL should be low on macrophages, while CD11b should still be expressed highly.

  For osteoclasts

• 10f.Within the live, singlet population gated above, for murine samples, gate on the CD68 high and CD115 high population followed by the CD11b low and CD265 high population (Figure 3H). For primate samples, gate on the CD68 high and CD117 high population followed by the CD11b low and CD254 high population (Figure 4H). CD11b should be lost as monocytes/macrophages differentiate into osteoclasts. CD265/RANK or CD254/RANKL is highly expressed on differentiated osteoclasts.
• 11.Run samples according to your sorter’s manufacturer’s protocol.

  Can readjust FSC and SSC setting for cells but do not change fluorescent detectors’ PMT voltages.

• 12.Place sorted samples on ice for downstream experiments.
• 13.Retain sorting reports with cell counts and percentage of stained cells data.

  Examples of cell percentages in the bone marrow are shown in Figures 5 and 6.

Figure 3. Representative Gating for Murine Bone-Residing Stromal Cells.

To isolate murine stromal cells, gating was performed to differentiate A) Singlets, B) Live/Dead Staining, C) hematopoietic stem cells (HSC), D) mesenchymal stem cells (MSC), E) osteoblasts, F) osteocytes, G) macrophages, and H) osteoclast.

Figure 4. Representative Gating for Primate Bone-Residing Stromal Cells.

To isolate primate stromal cells, gating was performed to differentiate A) Singlets, B) Live/Dead Staining, C) hematopoietic stem cells (HSC), D) mesenchymal stem cells (MSC), E) osteoblasts, F) osteocytes, G) macrophages, and H) osteoclast.

Figure 5. Murine Long Bone Stromal Cell Percentages across Age, Sex, and Genotype.

Isolated bone-residing stromal cells were stained and analyzed by cell sorting as described in Basic Protocols 2 and 3. The percentages of bone stromal cells from 3–5 individual animals are represented as mean±SEM.

Figure 6. Primate Vertebrae Bone Stromal Cell Percentages.

Archival baboon lumbar vertebrae (collected during routine necropsy at the Southwest National Primate Research Center (SNPRC) housed at Texas Biomedical Research Institute) and human lumbar vertebrae (from donated, deidentified cadaveric tissue to the Willed Body Program) were processed and stained prior to cell sorting. The percentages of bone stromal cells in three pooled samples (N=3 in each pool) are represented as mean±SEM.

ALTERNATE PROTOCOL 3

Flow Cytometry Analysis of Bone Stromal Cells

This protocol describes the counting of stained, isolated bone-residing stromal cells by flow cytometry and data analysis to examine the percentages of stained cells in the total cell sample.

Additional Materials:

• Stained composition beads (see Support Protocol 2)

• Flow Cytometer (BD FACS Canto II, or equivalent)

• Flow cytometry analysis software (FlowJo v10.6.1, or equivalent)

• 1.Run compensation beads to generate a compensation matrix

  Name all the parameters for consistency between experiments. An SSM 0–5 is great, 6–9 acceptable, and above 10 voltage gains will need to be adjusted. For positive and negative separation, 3–4 median channel logs are desired between the peaks, but fewer median channel logs may be used for markers with lower expression. Also, run live/dead controls with this step if samples were created with unfixed cells during staining in Basic Protocol 2.

• 2.Generate unstained gating controls for side scatter (SSC) and forward scatter (FSC) using unstained samples.
• 3.Using compensation beads, adjust FSC and SSC to visualize beads for each channel or fluorophore used and compensation samples. These can be used to set initial PMT voltages.
• 4.Run a single-stained bead sample to ensure positive peaks are on scale. PMT voltages can be decreased to shift peaks left and back onto the scale.

  Do not record data until all beads are reviewed.

• 5.Rerun single-stained bead samples to perform compensation set up and record files for compensation controls. Set an FSC vs. SSC gate to remove cell debris. Gate FSC-A vs. FSC-A around the singlet population and use it for further analysis.
• 6.Run samples according to flow cytometer manufacturer’s protocol recording at least 100,000 events per sample. Export files for further analysis.

Data Analysis

• 7.Open flow cytometry data in an appropriate gating software. Create a scatter gate of SSC-H vs. FSC-H to remove any debris.
• 8.Use FSH-H vs. FSH-A to create a singlets gate to remove doublets (Murine: Figure 3A, Primate: Figure 4A). This will be the main gate for further analysis of singlets.

  SSC-A vs. live/dead stain (AmCyan channel) is next to gate live cells from the singlets if samples were generated (Murine: Figure 3B, Primate: Figure 4B). The negative cells will be gated as “live” cells for further analysis in the following steps.

  For hematopoietic stem cells

• 9a.Within the population gated above, for murine samples, gate on the Sca1 high and CD34 high population (Figure 3C). For primate samples, within the population gated above, gate the CD45 high and CD34 high population (Figure 4C). These markers will identify a broad range of hematopoietic stem cells.

  For mesenchymal stem cells

• 9b.Within the population gated above, for murine samples, gate Sca1 high and CD29 high population and then gate the CD90 high and CD146 high population (Figure 3D). For primate samples, within the population gated above, gate on CD146 high and CD90 high population and then gate the CD146 high and CD29 high population (Figure 4D). CD146 is considered a definitive marker for mesenchymal stem cells (Correa et al., 2016), while Sca1 is found on multiple murine progenitor cell populations.

  For osteoblasts

• 9c.Within the population gated above, gate on the alkaline phosphatase high and CD90 high population (Murine: Figure 3E, Primate: Figure 4E). Only mature osteoblasts express alkaline phosphatase.

  For osteocytes

• 9d.Within the population gated above, gate on the SPARC high and podoplanin high population (Murine: Figure 3F, Primate: Figure 4F).

  For macrophages

• 9e.Within the population gated above, for murine samples, gate on the CD11b high and CD68 high population followed by the CD265 low and CD115 low (Figure 3G). For primate samples, gate on the CD11b high and CD68 high population followed by the CD254 low and CD115 low (Figure 4G). CD265/RANK or CD254/RANKL should be low on macrophages, while CD11b should still be expressed highly.

  For osteoclasts

• 9f.Within the population gated above, for murine samples, gate on the CD68 high and CD115 high population followed by the CD11b low and CD265 high population (Figure 3H). For primate samples, gate on the CD68 high and CD117 high population followed by the CD11b low and CD254 high population (Figure 4H). CD11b should be lost as monocytes/macrophages differentiate into osteoclasts. CD265/RANK or CD254/RANKL is highly expressed on differentiated osteoclasts.
• 10.The cell percentages or mean fluorescence intensities calculated by the analysis software can be graphed to profile changes in the numbers of bone marrow-residing cells.

SUPPORT PROTOCOL 2

Preparing Compensation Beads

This protocol describes preparing compensation beads for cell sorting and flow cytometry. Compensation beads are used to set voltages and gating parameters while running samples to obtain accurate data across experiments. The use of compensation beads for this protocol is crucial as the cells being examined are rare within the sample. Further information on compensation beads in flow cytometry has been described by (Perfetto et al., 2006).

Additional Materials:

• Compensation beads (ThermoFisher Scientific UltraComp eBeads, 01-2222-41, or equivalent)

1. Prepare a tube or well for each fluorochrome.

2. Mix beads vigorously by inverting at least 10 times or pulse vortexing.

3. Add one drop of beads to each tube or well.

4. Add 2 μL of each antibody in 2 mL Flow Cytometry Staining Media and mix well by inversion or pulse vortexing.

   Use volumes above those optimized after antibody titration.

5. Incubate at 4°C for 30 minutes in the dark.

6. Add 2 mL Flow Cytometry Staining Media to each tube and centrifuge at 400 g for 5 minutes.

7. Remove supernatant and add 400 μL of Flow Cytometry Staining Media to each sample.

8. Vortex prior to running the sample.

   Beads can be stored for up to 5 days at 4°C, except for live/dead stained compensation beads which needs to be analyzed the same day.

REAGENTS AND SOLUTIONS:

Flow Cytometry Staining Media

• 500 mL Hank’s Buffered Salt Solution (ThermoFisher, 88284)

• 15 mL heat-inactivated fetal calf serum (Gibco, 21540079)

• 2 mL 500 mM EDTA pH 8.0 (EDTA powder: Sigma E134)

• 1 mL 1M HEPES pH 7.0 (Gibco, 15-630-080)

• Store at 4°C for up to 12 months

• Heat inactivate fetal calf serum for 30 minutes at 56°C before use.

• Final concentrations: 3% FCS, 2 mM EDTA and 10 mM HEPES.

Cell Sorting Buffer

• 500 mL PBS

• 100 μL 500 mM EDTA pH 8.0 (Fisher, S311)

• 1% (w/v) bovine serum albumin (BSA, Fisher BP1600)

• Store at 4°C for up to 12 months

COMMENTARY:

Background Information:

The bone marrow comprises two distinct stem cell lineages: hematopoietic and mesenchymal. Hematopoietic stem cells give rise to all blood cell types, macrophages, and osteoclasts. In contrast, mesenchymal stem cell differentiation pathways are responsible for generating stromal cells, osteoblasts, and osteocytes, that compose the bone marrow microenvironment (Laurenti and Göttgens, 2018). The bone-forming osteoblasts balance bone-resorbing osteoclasts to maintain bone homeostasis during development and in response to structural stressors. However, this balance is often dysregulated in disease states leading to an overabundance of bone in osteosclerosis and a loss of bone in osteoporosis. Additionally, the relative concentration of osteoclasts and osteoblasts has profound effects on the development of bone metastatic cancers, with enhanced osteoclasts resulting in tumor-induced osteolysis and increased osteoblast activity resulting in osteoblastic lesions (Jinnah et al., 2018). The progenitor cells also play a role in the homing and engraftment of cancer cells to the bone marrow. Thus, there is a need to characterize the composition of the bone microenvironment in healthy and disease states. There have been two major obstacles to characterizing bone marrow microenvironmental composition changes. First, classification studies were severely underpowered and, second, biased due to the lack of high-throughput methodologies, compounded with variance and lack of quantitation and reproducibility (Zeng and Sanes, 2017).

Thus, we sought to develop a multi-fluorophore cell identification panel to accurately identify cell-specific alterations of key bone marrow niche cell types based on markers proposed in the literature. Markers for hematopoietic stem cells and mesenchymal stem cells are already well defined (Dominici et al., 2006; Boxall and Jones, 2012). For hematopoietic stem cells, we chose CD34 and CD45 for primates, including Sca1 for mice, to identify hematopoietic progenitors capable of multiple differentiation lineages. Additional markers for hematopoietic stem cells could include CD117/c-kit and lineage negative (Lin and Goodell, 2011; Rundberg Nilsson et al., 2013; Mayle et al., 2013). CD117 was not included due to its high expression on mast cells and endothelial cells (Foster et al., 2018, 2021; Li et al., 2003). For mesenchymal stem cells, we chose the markers CD29, CD90, and CD146 for primates, with the addition of Sca1 for mice. CD29/integrin β1 and CD90/Thy1 are highly expressed by mesenchymal lineage cells (Shekaran et al., 2014; Morris, 2018). CD146 is expressed on mesenchymal stem cells in the bone marrow and peripheral tissues (Somoza et al., 2015; Correa et al., 2016). Additional markers for mesenchymal stem cells could include CD73 and CD105 (Nery et al., 2013; Dominici et al., 2006). CD105/endoglin is also associated with microvessel formation and endothelial cell differentiation (Lee et al., 2012; Dallas et al., 2008) and thus may also stain cells of the hematopoietic lineage. Therefore, CD105 excluded from this panel. We focused on potential surface marker expression for bone-residing osteoclasts, osteoblasts, and osteocytes. Osteoclast surface marker CD265/RANK (receptor activator of nuclear factor-κB) and CD254/RANKL are associated with differentiation and activity of osteoclasts (Boyle et al., 1999). Thus, CD265 expression on cells expressing the macrophage surface markers CD68 and CD115 was postulated to represent osteoclasts or preosteoclast cells. Osteoblast surface proteins were proposed to be CD90/Thy-1 and alkaline phosphatase based on their roles in bone formation and bone matrix calcification (Trentz et al., 2010; Rutkovskiy et al., 2016; Picke et al., 2018). Osteocyte markers SPARC and Podoplanin (GP38) were derived from publications describing osteocyte ontogeny from osteoblasts (Bonewald, 2011; Zhu et al., 2011). Markers were chosen based on their expression on the cell surface, their ability to differentiate between bone stromal cell populations, and the commercial availability of fluorophore-conjugated antibodies. Antibodies were carefully selected with fluorophores to make a multi-fluorophore panel and have brighter fluorophores assigned to rarer cell surface proteins when possible. With this newly established multi-fluorochrome bone marrow cell-typing methodology, the cellular composition can be tracked during development and disease progression.

Critical Parameters:

For best results, fresh tissues should be used for digestion. The protocols described were optimized using frozen tissues; however, the cell viability after isolation is significantly decreased. If a high number of cells are required for downstream analysis after sorting, then additional samples would be needed if frozen, archival tissues are used. For cell staining, it is recommended that the antibody titration step be performed to optimize staining if any different antibodies or animal genotypes are tested. The inclusion of compensation beads in every experiment will improve the reproducibility of results between experiments. For flow cytometry, a high number of events is recommended for robust data, with at least 100,000 events being counted per sample.

Troubleshooting:

See Table 11 for possible problems, causes, causes, and solutions.

Table 11.

Troubleshooting Guide for Bone Stromal Cell Staining and Cytometry

Problem	Possible Cause	Solution
Low Signal/ High noise	Contamination with B or T cells	Gate or sort B and T cells out of the target population by staining samples with PE/Cy7 or PE/Cy5 conjugated antibodies to CD3, CD4, CD19, and CD45 (McLaughlin et al., 2008; Perfetto et al., 2004; Baumgarth and Roederer, 2000). In this case, the cells can be removed from the analysis by gating allowing for a higher signal to noise ratio. Depending upon the cytometer, other fluorophores can be used to create a “dump” channel where non-target populations can be measured or sorted. Additional markers for B and T cells can also be included.
Low Cell Isolation	Insufficient collagenase digestion	Try incubating in collagenase I for longer times or at a higher concentration. Another potential issue is too small vessel size for the tissue pieces to move freely. Tissues can also be cut smaller to increase the surface area exposed to the collagenase.
High Autofluorescence	Fixation method	Autofluorescence can occur with formaldehyde fixation for hematopoietic lineage cells. Use another fixative to reduce autofluorescence.
Loss of Fluorescence Signal	Fixation duration	Fixation at 1% formaldehyde should not affect fluorescence signal (including for PE and APC) if samples are run within two weeks. If longer storage is required or if the signal degrades, fixation can be performed overnight, followed by centrifugation and resuspension in PBS for longer storage.

Understanding Results:

This protocol was developed to reliably isolate bone marrow-derived cells for downstream fluorescence-activated cell-type identification and isolate mineralized bone cells including osteoblasts, osteoclasts, and osteocytes. In initial optimization for murine long bones, digestions were conducted with 0.5, 1, 2, and 3 mg/mL collagenase I at 37 °C for 30 min, 1 hour, 2 hours, and 4 hours to optimize cell yield and cell viability. The highest cell yield and cell viability were found at 1 mg/mL collagenase I digestion for 1 hour at 37 °C. Yields are consistent across mouse age, sex, and genotype (Figure 2). Fewer cells were isolated from female mice, although this is likely due to the smaller size of the starting material. Using this optimized procedure, 3 to 10 × 10⁶ cells are routinely recovered from each mouse. Additionally, protocols were developed and optimized to isolate cells from primate and murine vertebrae. Both procedures resulted in high numbers of cells recovered, as demonstrated in Tables 1–4. The number of cells recovered mainly indicates the efficacy of bone digestion. However, the length of digestion will also affect the viability of cells, and thus, we have optimized the protocol to maximize cell number and viability.

Using these freshly isolated samples, specific antibody panels for hematopoietic stem cells, mesenchymal stem cells, osteoblasts, osteoclasts, and osteocytes were created to identify bone marrow and stromal cells (Figure 1). These cells were analyzed by either cell sorting or flow cytometry using the gating schemes shown in Figures 3 and 4. Using these analysis protocols, the numbers of progenitor cells vary with sex and age. Aged female mice demonstrate increased osteoclast numbers, while osteoblast numbers are only reduced in older mice (both male and female) (Figure 5). This data is consistent with known results of bone aging and sex differences. Between primates- humans and baboons- the cellular composition in the bone marrow and stromal compartment were consistent (Figure 6). These data indicate that the results from non-human primates could be translatable to humans in future interventional studies. Either the total cell number or percentage of cells analyzed can be used to demonstrate cellular composition in the bone microenvironment. The cell count can be particularly important for downstream applications where the exact cell number is essential for reproducible results.

Time Considerations:

• Basic Protocol 1: Partial Digestion for Murine Long Bone Stromal Cell Isolation: 1–3 hrs

• Alternate Protocol 1: Partial Digestion for Primate Vertebrae Stromal Cell Isolation: 57–58 hrs

• Alternate Protocol 2: Murine Vertebrae Crushing for Bone Stromal Cell Isolation: 1–3 hrs

• Basic Protocol 2: Staining of Bone Stromal Cells: 4–5 hrs

• Support Protocol 1: Fluorescence Minus One Control, Isotype Control, and Antibody Titration Protocol: 4–6 hrs

• Basic Protocol 3: Cell Sorting of Bone Stromal Cells: 4–8 hrs

• Alternate Protocol 3: Flow Cytometry Analysis of Bone Stromal Cells: 4–8 hrs

• Support Protocol 2: Preparing Compensation Beads: 45 min

  Times will vary based on the number of samples

DATA AVAILABILITY STATEMENT:

The data, tools, and material (or their source) that support the protocol are available from the corresponding author upon reasonable request.