In Vivo Osteo-organoid Approach for Harvesting Therapeutic Hematopoietic Stem/Progenitor Cells

Summary

Here, we established in vivo osteo-organoids triggered by bone morphogenetic protein-2-loaded gelatin scaffolds to harvest therapeutic hematopoietic stem/progenitor cells for the reconstruction of a damaged hematopoietic and immune system. Overall, this approach can provide a promising cell source for cell therapies.

Abstract

Hematopoietic stem cell transplantation (HSCT) requires a sufficient number of therapeutic hematopoietic stem/progenitor cells (HSPCs). To identify an adequate source of HSPCs, we developed an in vivo osteo-organoid by implanting scaffolds loaded with recombinant human bone morphogenetic protein-2 (rhBMP-2) into an internal muscle pouch near the femur in mice. After 12 weeks of implantation, we retrieved the in vivo osteo-organoids and conducted flow cytometry analysis on HPSCs, revealing a significant presence of HSPC subsets within the in vivo osteo-organoids.

We then established a sublethal model of hematopoietic/immune system injury in mice through radiation and performed hematopoietic stem cell transplantation (HSCT) by injecting the extracted osteo-organoid-derived cells into the peripheral blood of radiated mice. The effect of hematopoietic recovery was evaluated through hematological, peripheral blood chimerism, and solid organ chimerism analyses. The results confirmed that in vivo osteo-organoid-derived cells can rapidly and efficiently reconstruct damaged peripheral and solid immune organs in irradiated mice. This approach holds potential as an alternative source of HSPCs for HSCT, offering benefits to a larger number of patients.

Introduction

Hematopoietic stem cell transplantation stands as the conventional therapy for a variety of hematological malignancies, as well as numerous inherited and autoimmune disorders^1,2,3,4. Nevertheless, the restricted quantity and origin of hematopoietic stem/progenitor cells (HSPCs) have emerged as a substantial impediment to the clinical implementation of hematopoietic stem cell transplantation (HSCT)^5,6.

Large-scale in vitro cell expansion is a commonly employed method for harvesting therapeutic cells^5,7. Various studies have developed conditions that stimulate HSPCs to self-renew in vitro, typically through the use of a combination of self-renewal agonists (such as cytokines and growth factors) and serum albumin, resulting in the ex vivo expansion of HSPCs⁸. However, it is important to note that current methods are still time-consuming and challenging to maintain the self-renewal capacity of expanded HSPCs⁹.

In contrast to the aforementioned method, harvesting cells in vivo presents a new and innovative strategy. This approach involves the establishment of an in vivo osteo-organoid that mimics the native bone marrow structure^10,11. To achieve this, we form in vivo osteo-organoids using bone morphogenetic protein-2 (BMP-2)-loaded gelatin scaffolds to obtain abundant and high-quality autologous cell cocktails, including HSPCs. By therapeutically applying these osteo-organoids, we have successfully treated irradiation damage and demonstrated that HSPCs derived from the osteo-organoids can rapidly and stably reconstitute the experimentally impaired immune system.

Protocol

Male and female C57BL/6 mice, aged 8-10 weeks, were included in the study. All mice were housed in the animal facility of East China University of Science and Technology. All the experimental procedures were approved by the Institutional Animal Care and Use Committees of East China University of Science and Technology (ECUST-21010).

1. Fabrication of bioactive scaffold

1. Preparation
   1. Put the cleaned and dried surgical scissors and tweezers into a 1,000 mL beaker. Wrap the mouth of the beaker with absorbent paper and tie it tightly with cotton thread.
   2. Place the beaker in the high-pressure sterilizer, close the lid tightly, set the sterilization program to 121 °C for 30 min, and start the sterilization program.
   3. After the completion of the sterilization process, remove and transfer the sterilized beaker to a drying oven set at 60 °C for drying. Once completely dried, spray the beaker with 75% (v/v) alcohol and place it on the clean bench.
   4. Thaw the rhBMP-2 stock solution (1.0 mg/mL) 2 h before the procedure.
   5. Once thawed to room temperature, transfer the rhBMP-2 stock solution and unopened gelatin sponge to the sterile clean bench for the fabrication of bioactive scaffolds.
      1. Cut the gelatin sponge (60 mm x 20 mm x 5 mm) into 48 uniformly sized pieces (5 mm x 5 mm x 5 mm) (Figure 1B) with the sterilized scissors and transfer them to a 48-well plate.
      2. Mix the rhBMP-2 stock solution by pipetting up and down gently to ensure even mixing, taking care to avoid bubbles. Aspirate 30 µL of the solution and add it dropwise to the gelatin sponge.
      3. Seal the 48-well plate using one layer of parafilm to facilitate the subsequent freeze-drying process. Take the plate containing the gelatin sponge out from the clean bench. Place it at -20 °C to be frozen. Simultaneously, turn on the freeze-dryer and precool it to a temperature range of -50 °C to -60 °C.
      4. After freezing for 2 h, place the well plate with gelatin sponges in the freeze-dryer for 12 h. To maintain the dryness and sterility of the samples, apply an additional layer of parafilm to seal the well plate after freeze-drying. Store it in a -20 °C refrigerator.

2. Surgical implantation of bioactive scaffolds

NOTE: All surgical instruments are sterilized for use.

1. Fast the mice for 1 h before anesthesia. Apply a lubricating vet ointment around the eyes of mice to reduce postoperative discomfort caused by dryness in the eye area. Anesthetize the mice by intraperitoneal injection of 1% (w/v) solution of pentobarbital sodium (at a dose of 48 mg/kg).
   NOTE: This protocol uses the ethically approved method of anesthesia with sodium pentobarbital intraperitoneal injection. Other methods, such as isoflurane inhalation or intraperitoneal injection of ketamine-xylazine, can also be used after approval.
2. Confirm the depth of anesthesia in mice by looking for the following signs in a supine mouse: steady heartbeat and respiration, relaxed muscles, immobile limbs, whiskers unresponsive to touch, and absence of a pedal reflex. In addition, pinch the skin of the mice with toothed forceps or stimulate the toes or paws of mice. If the mice display sensitive reactions, the anesthesia is too light; administer additional anesthesia. After the anesthetic plane is achieved, transfer the mice to the surgical area in the right lateral recumbent position to implant the bioactive scaffold into the left leg easily.
3. Shave the hair on the left leg using a shaving machine. Disinfect the shaved area using alternating rounds of chlorhexidine and alcohol 3 times each, and create incisions on the hind leg, located on the lateral side of the vastus lateralis muscle. Use ophthalmic scissors to make incisions approximately 3.0 mm in length, extending along the direction of the tibia.
   NOTE: To maintain sterile conditions during the surgery, the used surgical items should be stored in designated locations. Sterile supplies should be placed in a clean area to prevent contact with individuals other than the operator. Reusable surgical instruments should be promptly disinfected with 75% (v/v) ethanol. Minimize the duration of surgery to reduce the exposure of wounds to air and decrease the risk of infection.
4. To create the pocket for the implant, open the fascia at the distal aspect of the vastus lateralis muscle, towards the tibia.
5. Use ophthalmic forceps to shape the bioactive scaffold into a cylinder, approximately 5 mm in height and 5 mm in diameter. Implant the scaffold into the muscle pocket along the muscle slit created by ophthalmic scissors and close the implant pocket with an interrupted suture pattern through the fascia. Close the skin incision using a 4-0 suture.
   NOTE: The occurrence of convulsions or urination in animals during the surgical procedure indicates excessively deep anesthesia, necessitating prompt implementation of emergency measures.
6. Repeat steps 2.3-2.5 and implant another bioactive scaffold in the right femoral muscle pouch of the mice. Use iodine swabs to clean the surgical and injection sites of the mice after the surgery to prevent infection.
   NOTE: Only one implant can be inserted into each vastus lateralis muscle.
7. Place the mice on a constant temperature platform at 37 °C until they awake after the experiment. Turn the mice over every 10-15 min to prevent blood pooling. After surgery, in addition to maintaining warmth, monitor the postoperative mice without leaving the animal unattended until it has regained sufficient consciousness to maintain sternal recumbency. Pay close attention to important indicators of mice, such as heart rate and respiratory rate.
   NOTE: Trembling of the forelimbs in mice signifies the initiation of the recovery phase.
8. Place the mice back in a sterile cage that has been disinfected with alcohol spraying and ultraviolet irradiation. Do not place the animal that has undergone surgery with other animals until it has fully recovered. In addition, administer meloxicam to manage postoperative pain for at least one day, with the option to extend up to three days for more intensive pain management. Check the animal and the wound regularly during the 72-hour post-surgery period to identify any potential adverse effects. If infection is detected later, euthanize the animals by first anesthetizing the mice via intraperitoneal injection of a 1% (w/v) solution of pentobarbital sodium (at a dose of 48 mg/kg), followed by cervical dislocation after anesthesia.

3. Characterization of in vivo osteo-organoids 12 weeks after implantation

NOTE: The bioactive scaffolds will develop in vivo to form osteo-organoids after implantation.

1. Collection of in vivo osteo-organoids
   1. When the in vivo osteo-organoids have been implanted for 12 weeks, anesthetize the mice by intraperitoneal injection of 1% (w/v) solution of pentobarbital sodium (at a dose of 48 mg/kg), followed by cervical dislocation after anesthesia.
      NOTE: To minimize harm to animals, mice should be anesthetized prior to euthanasia by cervical dislocation.
   2. Spray 75% (v/v) alcohol on the skin at the site of dissection.
   3. Disconnect the hind limb from the trunk and remove the muscle from the osteo-organoids using miniature scissors carefully.
   4. Remove the attached soft tissues with gauze.
   5. Fix some osteo-organoids in a 4% (v/v) paraformaldehyde (PFA) solution for 24 h at room temperature for histological analysis. Utilize the remaining samples for macroscopic photography and flow cytometry analysis of HPSCs.
2. Macroscopic images
   1. Place the osteo-organoids on a blue background plate and photograph them with a digital camera.
3. Histological analysis
   1. Preparation for histological staining
      1. Transfer the osteo-organoids from the PFA solution to ultrapure water and soak for 1 h.
      2. Replace ultrapure water with 0.5 M ethylenediamine tetraacetic acid (EDTA) solution for decalcification for 1 week.
      3. Place decalcified tissues in an embedding cassette and submerge the cassette in ultrapure water for 1 h.
      4. For gradient dehydration, immerse the embedding cassette in ethanol at varying concentrations (v/v): 75%, 80%, 90%, and 95% for a duration of 1 h each, followed by immersion in 100% (v/v) ethanol I and II for 1 h each.
         NOTE: "100% (v/v) ethanol I and II" indicates that the embedding cassette should be immersed once in the "container I" filled with 100% (v/v) ethanol and then immersed a second time in the "container II" filled with 100% (v/v) ethanol.
      5. After dehydration, place the embedded tissues sequentially in a mixture of 100% (v/v) xylene and absolute ethanol (xylene: ethanol volume ratio of 1:1) for 1 h and 100% (v/v) xylene I and II for 30 min each.
      6. Immerse the tissues in paraffin I and II at a constant temperature of 60 °C for 2 h each.
      7. To embed the tissues soaked in wax using an embedding machine, place the melted wax into the embedding mold. Before the wax solidifies, remove the tissues from the dehydration container and place them in the embedding mold; label them accordingly. Cool the mold at -20 °C freezing condition and after the wax solidifies, remove the wax block from the embedding mold and trim it.
      8. Slice the embedded wax blocks into 4.5 µm thick sections using a paraffin slicer and then affix the sections to glass slides.
      9. Briefly bake the sections at 40 °C and then collect and store them in a sample box.
   2. H&E staining
      1. Place the sections in a sample rack and bake in a drying oven at 60 °C for 30 min.
      2. Deparaffinize the sections in xylene and ethanol. Sequentially place the paraffin sections into xylene I for 10 min, xylene II for 10 min, xylene III for 10 min, absolute ethanol I for 5 min, absolute ethanol II for 5 min, 90% (v/v) ethanol for 5 min, 80% (v/v) ethanol for 5 min, 70% (v/v) ethanol for 5 min, 50% (v/v) ethanol for 5 min and transfer into pure water for a gradient dewaxing process.
      3. Immerse the sections in the hematoxylin staining solution for 2 min, then rinse them in running water for 10 min.
      4. Immerse the sections in eosin staining solution for 2 min.
      5. Sequentially place the sections into 75% (v/v) ethanol for 2 min, 85% (v/v) ethanol for 2 min, absolute ethanol for 5 min, absolute ethanol for 5 min, and xylene for 5 min for dehydration.
      6. Take the sections out of xylene and seal them with neutral balsam.
      7. Observe the sections under the microscope and capture representative images at a magnification of 10x after the neutral balsam has dried.
4. Flow cytometry analysis for HPSCs
   1. Place the in vivo osteo-organoids obtained from step 3.1 in a mortar and add an appropriate amount of staining buffer. Cut the osteo-organoids into fragments with a diameter of 1-3 mm using ophthalmic scissors and crush them in the staining buffer using the mortar and pestle. Vertically press rather than grind the fragments of the in vivo osteo-organoids using a pestle to prevent a decline in cell survival rate.
   2. Collect the cell suspension into a centrifuge tube and centrifuge them at 300 × g for 5 min at 4 °C.
   3. Remove the supernatant and add 1 mL of red blood cell lysis buffer to lyse the cells for 3-5 min.
   4. Wash the cells with 1 mL of staining buffer and filter them through a 300 mesh nylon filter into a 5 mL flow cytometry tube.
   5. Centrifuge at 300 × g for 5 min at 4 °C and discard the supernatant. Stain the remaining approximately 100 µL (cells) at the bottom of the flow cytometry tube at 4 °C for 30 min using antibodies, including a live/dead staining kit, Peridinin chlorophyll protein-Cyanine5.5 (PerCp-Cy5.5)-anti-lineage cocktail (1:10), Phycoerythrin-Cyanine5 (PE-Cy5)-anti-c-kit (1:200), Alexa Fluor 700(AF700)-anti-Stem cells antigen-1(Sca-1) (1:200), Brilliant Violet 711(BV711)-anti-CD16/32 (1:200), bio-anti-CD34 (1:200), BV421-anti-CD127 (1:200), PE-CF594-anti-CD135 (1:200), BV510-anti-CD48 (1:200), and PE-Cy7-anti-CD150 (1:200).
   6. Resuspend the cells with 1 mL of Hanks' Balanced Salt Solution (HBSS) without Ca²⁺ and Mg²⁺ and centrifuge at 300 × g for 5 min at 4 °C.
   7. Aspirate the supernatant and stain with PE-streptavidin secondary antibody (1:200) for an additional 60 min at 4 °C.
   8. Resuspend the cells with 1 mL of HBSS (no Ca²⁺, Mg²⁺) and centrifuge at 300 × g for 5 min at 4 °C.
   9. Aspirate the supernatant and resuspend the cells with 300 µL of HBSS (no Ca²⁺, Mg²⁺).
   10. Vortex the solution to perform further flow cytometry analysis. Further analyze the obtained data using flow cytometry software to obtain quantitative data on HSPCs.
       NOTE: The data collection and analysis of flow cytometry refer to our previous articles and supporting materials (Supplemental Figure S1) for the typical gating strategies used to identify HSPCs¹⁰.
   11. Establish a side scatter-area (SSC-A) vs forward scatter-area (FSC-A) dot plot and gate the surrounding region of the cell population of interest (P1) in the SSC-A vs FSC-A dot plot during data acquisition on a flow cytometer. Double-click on the P1 gate; select the forward scatter-height (FSC-H) for the X-axis and the forward scatter-width (FSC-W) for the Y-axis to perform the first cell adhesion treatment. Double-click on the single cells gate; select the side scatter-height (SSC-H) for the X-axis and the side scatter-width (SSC-W) for the Y-axis to perform the second cell adhesion treatment. Use the rectangular gating tool to select the single cells.
   12. Exclude dead cells. Distinguish live cells from dead cells by detecting viability markers. Establish a dot plot of SSC-A vs Live/Dead and gate the region corresponding to live cells, naming it "live cells".
   13. Screen the cells labeled with specific antibodies in "live cells". Exclude the cell population that may be affected by the positive expression of lineage-specific antibodies in the myeloid hematopoietic stem/progenitor cell lineage by using lineage-negative (lin^-) gating. Gate the lin^- cells.
   14. Screen for different subpopulations of cells within the lin^- cells. Double-click on the lin^- cells gate; select Sca-1-AF700 for the X-axis and c-kit-PE-Cy5 for the Y-axis. Gate the c-kit⁺Sca-1^- cells as the LKS^- cells, the c-kit⁺Sca-1⁺ cells as the LKS⁺ cells, and the cells with low expression of c-kit and Sca-1 as the LK^loS^lo cells. Double-click on the LKS^- gate and select CD34-PE for the X-axis and FCγR-BV711 for the Y-axis. Double-click on the LKS⁺ gate; select FIt3-PE-CF594 for the X-axis and IL7Rα-BV421 for the Y-axis and select CD150-PE-Cy7 for the X-axis and CD48-BV510 for the Y-axis.
   15. Based on the specificity of antibodies, identify positive or negative gates to distinguish cell subpopulations: hematopoietic stem cells (Lin⁻c-kit⁺Sca-1⁺CD48⁻CD150⁺) and differentiated hematopoietic progenitors: multipotent progenitors (Lin⁻c-kit⁺Sca-1⁺CD48⁻CD150⁻), common lymphoid progenitors (Lin⁻c-kit^-Sca-1^-Flt3⁺IL7Rα⁺), common myeloid progenitors (Lin⁻c-kit⁺Sca-1⁻CD34⁺FCγR⁻), granulocyte-monocyte progenitors (Lin⁻c-kit⁺Sca-1⁻CD34⁺FCγR⁺), and megakaryocyte erythroid progenitors (Lin⁻c-kit⁺Sca-1⁻CD34⁻FCγR⁻).

4. Mouse irradiation model

NOTE: C57BL/6 mice were irradiated with X-rays using an X-ray irradiator.

1. To achieve sublethal impairment of the hematopoietic/immune system, expose wild-type (WT) mice in the 5 Gray (Gy) phosphate buffered saline (PBS)-treated group, the 5 Gy native bone marrow (BM)-treated group, and the 5 Gy osteo-organoids-treated group to 5 Gy irradiation (1.25 Gy/min) 24 h prior to transplantation.
   NOTE: Make sure that WT mice in the 0 Gy PBS-treated group are treated with PBS and do not undergo irradiation.

5. Cell therapy process

NOTE: All procedures should be done in sterile conditions.

1. Acquisition of in vivo osteo-organoid-derived- and native bone marrow- derived cells
   1. Implant two bioactive scaffolds into the muscle pouch of C57BL/6.SJL-Ptprca Pepcb/BoyJ (CD45.1) mice and incubate them for 12 weeks to form the in vivo osteo-organoids. Harvest two in vivo osteo-organoids and two femurs from one CD45.1 mice mentioned above. Subsequently, clear the soft tissue on the surface of the osteo-organoids and femurs using forceps and gauze.
   2. Separately place the osteo-organoids and femurs into a mortar and add 1 mL of HBSS (no Ca²⁺, Mg²⁺).
      NOTE: HBSS without Ca²⁺ and Mg²⁺ can reduce cell aggregation. The solution used for transplantation must not contain serum components to avoid any potential rejection reactions.
   3. Cut the samples up with surgical scissors and gently crush them in a mortar and pestle.
   4. Filter the cell suspension through a 40 µm cell strainer and centrifuge at 300 × g for 5 min at 4 °C.
   5. Remove the supernatant and resuspend the cell pellets with 300 µL of HBSS (no Ca²⁺, Mg²⁺). Mix thoroughly to obtain a suspension of osteo-organoid-derived cells or whole bone marrow cells.
      NOTE: To fully utilize the cell cocktails derived from the in vivo osteo-organoid, we chose to use whole bone marrow cells for therapy. The typical number of total cells in a 30 µg BMP-2 induced in vivo osteo-organoid is 30,000,000-40,000,000.
2. Orbital venous sinus injection
   NOTE: CD45.1⁺ whole bone marrow cells collected from in vivo osteo-organoids or femurs were injected through the orbital venous sinus into the sublethally irradiated C57BL/6 (CD45.2) recipient, respectively. Orbital venous sinus injection and tail vein injection are commonly used methods in animal experiments. Orbital venous sinus injection was chosen due to its simplicity and low failure rate for beginners. During the process of orbital injection, certain protective measures need to be taken, such as pre-anesthetizing the mice, using a heated plate to reduce the risk of hypothermia, and restraining mechanically the mice to prevent movement.
   1. Place the CD45.2 mice in a preanesthesia chamber and induce anesthesia using isoflurane. Apply a lubricating vet ointment around the eyes of mice to prevent dryness and counteract the ocular irritations caused by isoflurane.
      NOTE: To minimize harm to the animals, isoflurane anesthesia is administered to mice before orbital venous sinus injection.
   2. Transfer the mice to the anesthesia table for further anesthesia.
   3. To reduce the likelihood of hypothermia in mice, place the irradiated, anesthetized mouse on a thermostatic heating pad set at 37 °C.
   4. Place the mouse in left lateral recumbency with its head facing to the right after being fully anesthetized.
   5. To protrude the eye, place a finger on the top of the head and along the jawline. Gently pull the skin back and down.
   6. Use a 1 mL syringe fitted with a 25 G needle to aspirate 200 µL of the in vivo osteo-organoid-derived cell suspension or native bone marrow-derived cell suspension of CD45.1⁺ mice obtained in step 5.1.5.
   7. Carefully introduce the needle, bevel down, at an angle of approximately 30°, into the medial canthus.
      NOTE: Injections are usually given with the needle bevel up, but for retro-orbital injections, it is better to position the needle bevel down to reduce the risk of eye damage.
   8. Use the needle to follow the edge of the eyeball down until the needle tip is at the base of the eye.
   9. Slowly and smoothly inject the injectate.
   10. After the injection is complete, place the mouse back into its cage for recovery.

6. Evaluation of treatment effects

1. Hematological analysis
   NOTE: To analyze the hematology of recipients in immune system reconstitution assays, we collected peripheral blood samples from recipients subjected to different treatments: 0 Gy PBS treatment, 5 Gy PBS treatment, 5 Gy native BM treatment, and 5 Gy osteo-organoid treatment. The blood samples were obtained through retro-orbital puncture 2 days before and 2, 4, 8, 12, and 16 weeks after transplantation. A portion of the blood sample of the same group was used for hematological analysis, while the other part was used for chimerism analysis. There are different blood sampling methods, such as through submandibular vein or the tail vein. Retro-orbital puncture is one of the common methods for blood sampling in mice and is permitted under ethical conditions.
   1. Repeat steps 5.2.1-5.2.5 to anesthetize the mice and protrude the eyes of the mice.
      NOTE: To mitigate harm to the animals, isoflurane anesthesia is administered to the mice before retro-orbital puncture blood sampling.
   2. Position the capillary tube with a diameter of 0.5 mm in the medial canthus of the eye and direct it caudally at a 30°-45° angle relative to the plane of the nose.
   3. Rotate the capillary tube gently while applying pressure, allowing it to penetrate through the conjunctival membranes. Allow the blood to flow into the capillary tube via capillary action.
   4. Apply continuous pressure after the blood starts flowing to sustain the flow.
   5. After drawing 100 µL of blood, release the pressure on the neck immediately. Simultaneously, remove the capillary tube to allow the eye to return to the normal position to prevent postoperative bleeding from the puncture site.
   6. Promptly expel the blood into a centrifuge tube that had been washed with 1.5% (w/v) ethylenediamine tetraacetic acid dipotassium salt dihydrate (EDTA-K2) solution and then had been added 10 µL of 1.5% (w/v) EDTA-K2 solution. Gently shake the centrifuge tube to ensure thorough mixing of EDTA-K2 solution and blood, preventing clotting.
   7. After wiping the wound with sterile cotton and applying slight pressure to stop bleeding, observe the mice for 1-2 min after hemostasis to ensure there are no abnormalities and then return them to their cages.
   8. Perform hematological analysis on a portion of each group's blood samples using a hematology analyzer, including the counts of white blood cells (WBC), red blood cells (RBC), and platelets (PLT).
      NOTE: Analyze all blood samples within 2 h of collection.
2. Analysis of peripheral blood chimerism
   NOTE: Similar to hematological analysis, peripheral blood samples from 5 Gy native BM-treated and 5 Gy osteo-organoid-treated groups were collected by retro-orbital puncture at 2, 4, 8, 12, and 16 weeks after transplantation.
   1. Use the other portion of the peripheral blood sample obtained from step 6.1 for analysis of peripheral blood chimerism.
   2. Add 1 mL of erythrocyte lysis buffer to the sample and lyse for 3-5 min. Invert the sample upside down gently to facilitate better lysis of red blood cells.
   3. Centrifuge at 300 × g for 5 min at 4 °C.
   4. Discard the supernatant and add 1 mL of HBSS (no Ca²⁺, Mg²⁺) to resuspend the pellet.
   5. Centrifuge at 300 × g for 5 min at 4 °C and discard the supernatant. Stain the remaining approximately 100 µL of cells at the bottom of the centrifuge tube at 4 °C for 30 min using antibodies, including a live/dead staining kit, PE-anti-CD45.1 (1:200), Fluorescein Isothiocyanate (FITC)-anti-CD45.2 (1:200), PE-Cy7-anti-B220 (1:200), AF700-anti-CD11b (1:200), PerCp-Cy5.5-anti-CD8a (1:200), PE-Dazzle594-anti-CD4 (1:200), and Allophycocyanin (APC)-anti-CD3e (1:200).
   6. Resuspend the cells with 1 mL of HBSS (no Ca²⁺, Mg²⁺) and centrifuge at 300 × g for 5 min at 4 °C.
   7. Aspirate the supernatant and resuspend the cells with 300 µL of HBSS (no Ca²⁺, Mg²⁺).
   8. Vortex the suspension to perform further flow cytometry analysis.
   9. Further analyze the obtained data using flow cytometry software to obtain information about the chimerism rate of HSPCs in the peripheral blood of recipient mice in both the 5 Gy native BM-treated group and the 5 Gy osteo-organoid-treated group.
      NOTE: For the analysis of flow cytometric data, refer to Supplemental Figure S2 for the typical gating strategies used to identify donor-derived CD45.1⁺ cells in peripheral blood of CD45.2 recipients.
   10. Repeat steps 3.4.11-3.4.12 to gate the region corresponding to live cells.
   11. Analyze peripheral blood chimerism by initially identifying cell populations and subsequently discerning donor and recipient cells.
       1. Screen specific antibody-labeled cells within live cells. Select CD11b-AF700 for the X-axis and B220-PE-Cy7 for the Y-axis. Gate the B220⁺CD11b^- cells as B cells, the B220^-CD11b⁺ cells as myeloid cells, and the B220^-CD11b^- cells as double-negative (DN) cells.
       2. Double-click on the DN cells gate, using CD3-APC for the X-axis and SSC-A for the Y-axis. Gate the CD3⁺ cells as T cells.
       3. Double-click on the CD3⁺ cells gate, selecting CD4-PE-Dazzle594 for the X-axis and CD8-PerCp-Cy5.5 for the Y-axis. Gate the CD8⁺CD4^- cells as cytotoxic T cells (CTL) and the CD8^-CD4⁺ cells as regulatory T cells (Treg).
   12. Finally, peripheral blood chimerism analysis is performed on the donor and recipient cells. Within the designated cell populations, double-click to select CD45.1-PE for the X-axis and CD45.2-FITC for the Y-axis. Gate the CD45.1⁺ cells and perform cell counting to determine the proportion of CD45.1⁺ cells within the cell population, thus reflecting the chimerism rate.
3. Analysis of solid organ chimerism
   NOTE: At 16 weeks post transplantation, BM, spleen, and thymus chimeras were analyzed in the 5 Gy BM-treated group and the 5 Gy osteo-organoid-treated group.
   1. Repeat steps 5.1.1-5.1.4 to obtain the cell pellets derived from the BM.
   2. Place a 300 mesh nylon filter above a cell culture dish (6 cm) containing 1 mL of HBSS (no Ca²⁺, Mg²⁺).
   3. Place the dissected spleen or thymus on the nylon filter.
   4. Grind the mentioned organs using the front rubber stopper of a 5 mL syringe until no obvious tissue structure remains, only residual fibrous tissue. Handle the cells gently to ensure cell viability.
   5. Collect the filtrate and centrifuge at 300 × g for 5 min at 4 °C.
   6. Remove the supernatant to obtain the cell precipitates derived from the spleen or thymus.
   7. Add 1 mL of red blood cell lysis buffer to the cell pellets obtained in steps 6.3.1 and 6.3.6, and lyse for 3-5 min.
   8. Repeat steps 6.2.3-6.2.8 for staining and flow cytometry analysis.
   9. Further analyze the obtained data using flow cytometry software to obtain information about the chimerism rate of HSPCs in the solid organs (BM, spleen, and thymus) of recipient mice in both the 5 Gy native BM-treated group and the 5 Gy osteo-organoid-treated group.
      NOTE: For the analysis of flow cytometric data, refers to Supplemental Figure S2 for the typical gating strategies used to identify donor derived CD45.1⁺ cells in BM, spleen, and thymus of CD45.2 recipients.
   10. Replace peripheral blood in steps 6.2.10-6.2.12 with BM, spleen, and thymus to obtain the chimerism rate of solid organs.

Results

As per the protocol, we have created a bioactive scaffold by dripping BMP-2 into a degradable gelatin sponge under sterile conditions. The scaffold was then implanted into the lower limb muscles of mice to establish in vivo osteo-organoids. After an incubation period of 12 weeks, we conducted macroscopic photography, histological analysis, and flow cytometry analysis on the osteo-organoids (Figure 1A). The gelatin sponge was cut into cubes with dimensions of 5 mm x 5 mm x 5 mm, and ...

Discussion

In this protocol, we present an approach to establish in vivo osteo-organoids with bone marrow-like structures by implanting gelatin sponge scaffolds loaded with BMP-2. We demonstrate that these in vivo osteo-organoids can stably produce therapeutic HSPCs over a long period of time (more than 12 weeks). Compared to existing in vitro expansion or in vivo incubation methods that load cells, this protocol can obtain cell cocktails with diverse cell types, including HSPCs and various immun...

Materials

Name	Company	Catalog Number	Comments
AF700-anti-CD11b (M1/70)	eBioscience	56-0112-82	Store at 4 °C. Dilute 1:200 for staining.
AF700-anti-Sca-1 (D7)	BioLegend	108141	Store at 4 °C. Dilute 1:200 for staining.
APC-anti-CD3e (145-2C11)	Tonbo	20-0031-U100	Store at 4 °C. Dilute 1:200 for staining.
bio-anti-CD34 (RAM34)	eBioscience	13-0341-82	Store at 4 °C. Dilute 1:200 for staining.
BV421-anti-CD127 (IL-7Rα)	BioLegend	135023	Store at 4 °C. Dilute 1:200 for staining.
BV510-anti-CD48 (HM48-1)	BioLegend	103443	Store at 4 °C. Dilute 1:200 for staining.
BV711-anti-CD16/32 (93)	BioLegend	101337	Store at 4 °C. Dilute 1:200 for staining.
Capillary tube	Shanghai Huake Labware Co.	DC616297403604-100mm/0.5mm
Cell strainer	CORNING	352340
Ethanol	GENERAL-REAGENT	01158566
Ethylenediamine tetraacetic acid (EDTA) solution	Servicebio	G1105
Ethylenediaminetetraacetic acid dipotassium salt dihydrate (EDTA-K2)	Solarbio	E8651
FITC-anti-CD45.2 (104)	BioLegend	109806	Store at 4 °C. Dilute 1:200 for staining.
Flowjo	Becton, Dickinson & Company		A flow cytometry.
Gelatin sponge	Jiangxi Xiangen Co.		Use under sterile conditions.
HBSS without Ca2+ and Mg2+	Gibco	14170112	HBSS without Ca2+ and Mg2+ can prevent cell aggregation.
Hematology analyzer	Sysmex	pocH-100i Diff
iodine swabs	Xiangtan Mulan Biological Technology Co., Ltd.	01011	To prevent the infection after operation.
Isoflurane	RWD	R510-22-10	To avoid adverse effects of anesthesia waste gases on the environment and laboratory personnel, a gas recovery system should be used in conjunction.
Kraft paper			absorbent paper
LIVE/DEAD Fixable Near IR Dead Cell Staining Kit(used in 3.4.5)	Thermo Fisher Scientific	L34962	A live/dead staining kit. Store at -20 °C. Dissolve in 50 μL of DMSO for working solution.
lubricating vet ointment	Pfizer		To prevent dryness and counteract the ocular irritations caused by isoflurane.
Neutral balsam	Solarbio	G8590
nylon filter	Shanghai Shangshai Wire Mesh Manufacturing Co., Ltd.		Used for cell filtration.
Paraffin liquid	Macklin	P815706
Paraformaldehyde (PFA) solution	Servicebio	G1101	Immersion fixation is used for routine animal tissues. The volume of fixative used is generally 10-20 times the tissue volume, and fixation at room temperature for 24 hours is sufficient.
PE-anti-CD45.1 (A20)	BioLegend	110708	Store at 4 °C. Dilute 1:200 for staining.
PE-CF594-anti-CD135 (A2F10.1)	BD Biosciences	562537	Store at 4 °C. Dilute 1:200 for staining.
PE-Cy5-anti-c-kit (2B8)	BD Biosciences	105809	Store at 4 °C. Dilute 1:200 for staining.
PE-Cy7-anti-B220 (RA3-6B2)	BioLegend	103222	Store at 4 °C. Dilute 1:200 for staining.
PE-Cy7-anti-CD150 (TC15-12F12.2)	BioLegend	115914	Store at 4 °C. Dilute 1:200 for staining.
PE-Dazzle594-anti-CD4 (GK1.5)	BioLegend	100456	Store at 4 °C. Dilute 1:200 for staining.
Pentobarbital sodium salt	Sigma-Aldrich	57-33-0	Prepare for use at a concentration of 1% (w/v).
PerCp-Cy5.5-anti-CD8a (53-6.7)	BioLegend	100734	Store at 4 °C. Dilute 1:200 for staining.
PerCp-Cy5.5-anti-lineage cocktail	BD Biosciences	561317	Store at 4 °C. Dilute 1:10  for staining.
Red blood cell lysis buffer	Beyotime	C3702	Store at 4 °C. Use in clean bench.
rhBMP-2	Shanghai Rebone Biomaterials Co.		The concentration of rhBMP-2 in the stock solution is 1.0 mg/mL.
Staining buffer	BioLegend	420201	Store at 4 °C.
Xylene	GENERAL-REAGENT	01018114
Zombie UV Fixable Viability Kit (used in 6.2.5)	BioLegend	423108	A live/dead staining kit. For reconstitution, bring the kit to room temperature; add 100 µL of DMSO to one vial of Zombie UV dye until fully dissolved.