Abstract
Renal capsule transplantation is a very helpful method to grow embryonic tissues or tumors in a vascular environment, allowing long-term engraftment and biological analyses. This chapter describes the surgical procedure for the transplantation of embryonic skeletal elements in the renal capsule of adult mice and points out the manipulations that can be applied for assaying the role of angiogenesis during bone development and repair.
Keywords: skeletal development, skeletal repair, angiogenesis, renal capsule transplantation
1. Introduction
Bone is a highly vascularized tissue with tight connections between blood vessels, bone marrow, and bone cells to maintain skeletal integrity. Angiogenesis plays a pivotal role in skeletal development and particularly during endochondral ossification as an angiogenic switch is required for the replacement of cartilage by bone marrow and bone [1–3]. Numerous tools and methodologies have been used to study the impact of angiogenesis on osteogenesis both in vitro and in vivo [4–8]. Although in vitro angiogenesis assays have provided direct evidence for bidirectional interactions between osteoblasts and endothelial cells, which are crucial for osteogenesis, other cell types, circulating factors, and extracellular matrix proteins are involved in bone vascularization. Therefore, in vivo angiogenic assays are also essential to study the role of supporting cells (smooth muscle cells, pericytes, fibroblastic cells) and other factors in the tissue environment. Moreover, in vitro assays do not allow the development of the hematopoietic compartment of bone, which is required for establishing the stromal compartment of bone and providing osteoclasts that are together necessary for bone formation and remodeling. Since the kidney is one of the most vascularized organs in the body, the renal capsule constitutes a permissive environment to grow cells, tumors or embryonic tissues [9–13]. The renal capsule of adult mice has been used as a host environment to dissect the role of angiogenesis in skeletal development [13, 14]. With the growing number of genetically modified mouse models, this approach can help distinguish the effects of specific gene mutations in skeletal tissues versus blood vessels and their impact on angiogenesis and subsequent bone development [15, 16]. Any skeletal element from the developing embryo can potentially be collected prior to its vascularization in vivo and transplanted in the adult host renal capsule. More recently, this model was used to grow human embryonic skeletal elements [17]. Vascularization of the grafts occurs within three days, and the renal capsule environment can support normal bone development and growth, including establishment of the bone marrow, cortical bone and surrounding periosteum (Figure 1). Finally, skeletal stem/progenitors within bone marrow and periosteum can be isolated from long bones grown in the renal capsule and mobilized to repair bone after skeletal injury thus extending the use of this system to study bone repair mechanisms (Figure 2) [18].
Figure 1: Steps of the surgical procedure and development of skeletal elements in the renal capsule.
(A) Anesthetized host mouse prior to transplantation; note the position of the skin incision. (B) Exteriorized kidney post-transplantation of one E14.5 femoral skeletal element (denoted by a white dotted line). The white arrow indicates the incision in the renal capsule. (C) Femoral skeletal elements at days 0 (d0), 5 (d5), 7 (d7) and 60 (d60) post-transplantation. By day 60, the skeletal element is fully ossified and has grown to reach almost the size of a two-month-old mouse femur (approximately 1cm in length). (D) PECAM immunostaining reveals blood vessels (black arrows) on longitudinal sections of femoral skeletal elements at d0 (E14.0), d5 and d7 post-transplantation. At the time of transplantation (d0), the perichondrium (pc) is vascularized but not the cartilage (c). The cartilage becomes vascularized by day 5 to form the primary ossification center. The bone marrow (bm) and periosteum (po) are well developed by day 7; the epiphyseal cartilage (c) is not yet invaded by blood vessels to form the secondary ossification center.
Figure 2: Steps of the surgical procedure and repair after cortical defect or fracture in long bones grown under the renal capsule.
(A-D) Steps of cortical defect repair and (E-H) steps of fracture repair. (A, E) Exteriorized kidney 6 weeks post-transplantation of an E14.5 femoral skeletal element. (B, F) Cortical defect (B) and bone fracture (F) at the time of procedure (white arrows point to the injury site). (C, G) Healing cortical defect (C) and bone fracture (G) 2 weeks after the procedure (white arrows indicate the repair site). (D, H) Longitudinal sections of cortical defect (D) and fracture (H) stained with Safranin’O showing a fully ossified callus (orange dotted line) composed of newly formed bone (black arrows).
2. Materials
2.1. Anesthetics and analgesic
Anesthetics: Prepare the solution of ketamine/medetomidine by mixing 1 volume of ketamine with 1 volume of medetomidine.
Anesthetic reversal solution: Atipamezole comes as a ready to use reagent.
Analgesics: Prepare the solution of buprenorphine in NaCl 0.09%.
2.2. Isolation of E14-E14.5 mouse femora
Pregnant female mice with embryos at E14-E14.5 (see Note 1).
Surgical instruments (Fine forceps Dumont #5 and #55, scissors, Fine Science Tools).
Ice cold 1XPBS: Phosphate buffer saline solution.
70% Ethanol.
24-well plate.
Petri dish 100 mm diameter.
Binocular microscope.
2.3. Renal capsule transplantation
24-Well plate.
Insulin micro-fine syringe (30G).
Electric shear.
Male mice (8–12 week old) (see Note 1).
Pregnant mouse female (E14-E14.5).
Binocular microscope.
Betadine Soap and Betadine solution.
Cotton swab.
1XPBS: Phosphate buffer saline solution.
Plastic Pasteur pipette.
“L”-shape glass rod (Home made): Using a fire, separate the narrow end of a glass Pasteur pipette (4–5cm in length); make a thin “L”-shaped glass rod with a rounded closed end of approximately 1mm in diameter.
Surgical instruments (Fine forceps, Fine Vanna Scissors, tweezers, hemostatic forceps, scissors)(Fine Science Tools) (see Note 2).
4–0 Absorbable sutures.
Clips and wound clipper.
2.4. Bone fracture and cortical defect in the renal capsule
Electric shear.
Binocular microscope.
Betadine Soap and Betadine solution.
Surgical instruments (Fine forceps, Fine Vanna Scissors, tweezers, scissors)(Fine Science Tools) (see Note 2).
Driller with 0.8mm drill bit.
4–0 absorbable sutures.
Clips and wound clipper.
2.5. Analysis of vascularization and angiogenesis
Glass jar.
4% Paraformaldehyde fixative solution.
0.5M EDTA pH 7.4.
70% Ethanol.
95% Ethanol.
100% Ethanol.
Xylene.
Paraffin.
Superfrost Microscope slides.
Histoclear.
1XPBS: Phosphate buffer saline solution.
Rotary microtome.
Deionized water.
Hydrophobic pen.
Hydrogen peroxidase (H2O2).
Methanol.
Ficin solution.
Glycine.
Ovalbumin.
Nonfat powdered milk
Normal Goat serum.
Rat anti-PECAM primary antibody (BD Biosciences).
Goat biotinylated anti-Rat secondary antibody (BD Biosciences).
Horseradish Peroxidase (HRP)- conjugated Streptavidin (BD Biosciences).
Diaminobenzidine (DAB): Prepare working solution according to supplier manual (Life Technologies).
Fast Green.
Permount.
Cover slides.
3. Methods
3.1. Isolation of mouse embryonic femora
Prepare the cartilage grafts by finely dissecting the skeletal elements of E14-E14.5 mouse embryos using 2 pairs of fine forceps (see Note 3).
Sacrifice pregnant mouse by cervical dislocation under anesthesia (IP injection of ketamine/medetomidine: 50 mg of ketamine and 0.5 mg of medetomidine per kg of body weight) and position the mouse in a supine posture.
Soak the abdomen with 70% Ethanol, and make a small incision at the midline. Continue with a V-shaped incision through the skin and pull the skin toward the head to expose the abdomen.
Cut the peritoneum to expose the abdominal cavity.
Locate the 2 uterine horns, the uterus and oviduct in the dorsal region of the abdomen cavity.
Explant the uterus by cutting the mesometrium and the surrounding fat tissue. Place the uterus in ice cold 1x PBS (see Note 4).
Discard the pregnant mouse and proceed for embryo dissection.
Separate each embryo by cutting between implantation sites along the uterine explant.
Make a small incision through the decidua tissue surrounding each embryo and with a pair of fine forceps, tear decidua apart and the embryo can be shelled out.
Once embryo is removed, Reichert’s membrane may still be attached as well as the ectoplacental cone (see Note 5).
Place the embryo in a clean petri dish with clean ice-cold 1xPBS and proceed to carefully dissecting the embryo under the binocular microscope.
Using fine forceps, carefully separate the upper and bottom parts of the embryo body by cutting through the abdomen. Discard the upper body.
Carefully peel the skin to visualize the femora.
Use the surrounding soft tissue to hold the hindlimb with the forceps and separate the hindlimb from the hip.
Using a pair of fine forceps, pinch the soft tissue on both sides of the femora and the soft tissue on both sides of the tibia. Pull to separate the femora and the tibia. Discard tibia.
Take off the surrounding soft tissue. Keep some to be used to grasp the femoral cartilage (see Note 6).
Place the femoral cartilage grafts in ice-cold 1xPBS or DMEM medium in a 24-well plate on ice for no longer than 2 hours for optimal development after transplantation.
3.2. Renal capsule transplantation
Weigh male (8–12 week old) mice and induce general anesthesia with an IP injection of ketamine/medetomidine (50 mg of ketamine and 0.5 mg of Medetomidine per kg of body weight).
Perform a subcutaneous injection of analgesics solution (0.1 mg buprenorphine in NaCl 0.09% per kg of body weight) (See Note 7).
With the mouse under anesthesia, shave the left flank with the electric shear.
Position the mouse on its side with the left shaved flank facing up under the binocular microscope (Fig. 1).
Swab the shaved area center-out with Betadine soap followed by Betadine solution.
Locate the left kidney and make a small longitudinal incision of approximately 1–1.5 cm through the skin and the body wall (Fig. 1) (see Note 8).
Expose the kidney outside the body by pulling with forceps the fat located at the distal pole of the kidney and simultaneously applying a slight pressure to both sides of the incision with the forefinger and thumb to pop the kidney out of the abdominal cavity. The exteriorized kidney will rest on the body wall. Keep the kidney moist by applying a PBS solution with a Pasteur pipette (see Note 9).
Prepare the graft site by making a small 2 mm hole in the renal capsule at the base of the kidney using small Vanna scissors (Fig. 1, arrow) (see Note 10).
Insert the “L”-shape glass rod into the hole and carefully slide it in between the capsule and the kidney parenchyma to make a small pouch for the graft (see Note 11).
Transfer the graft to the surface of the kidney using a pair of fine forceps (see Notes 12–14).
Insert the graft into the pouch by gently lifting the capsule with one pair of fine forceps and by placing the graft under the capsule with another pair of forceps. Once the graft is entirely covered with the capsule, guide it with the forceps to position it in the mid-axial part of the kidney (Fig. 1, white dotted line).
Reposition the kidney into the body cavity and close the body wall layer with 2 stiches using a 4–0 silk absorbable suture.
Align both sides of the skin incision together and close the skin with 2 or 3 clips using a wound clipper.
If needed, clean the skin of the mouse using a Betadine solution swab.
Inject the anesthetic reversal solution (0.1 mg atipamezole per kg body weight) via IP injection and place the mouse on a heating blanket set at approximately 37°C for recovery. Monitor the mice closely until fully awake. Let the mice ambulate freely to access food and water.
Monitor mice daily and remove skin staples after 2 weeks (see Note 15).
3.4. Bone fracture and cortical defect in the renal capsule
Six to 8 weeks after transplantation of E14-E14.5 femora in the renal capsule, weight and anesthetize the host mice as described in section 3.2.
Shave the left flank of the mice and clean the shaved area with Betadine soap then Betadine solution.
At the level of the left kidney, incise the skin and the body wall longitudinally on approximately 1–1,5 cm.
Expose the kidney with the bone transplant to make it accessible as described in section 3.2 (See Note 16) (Fig. 2A, 2E).
-
To induce a cortical defect, drill a hole of 0.8mm in diameter into one cortex in the diaphysis (Fig. 2B).
To induce a bone fracture, cut the bone in the mid diaphysis with scissors (See Note 17) (Fig. 2F).
Reposition the kidney into the body cavity and suture the body wall using a 4–0 silk absorbable suture.
Close the skin incision with 2 or 3 clips using a wound clipper.
If needed, clean the skin of the mouse using a Betadine solution swab.
Inject the anesthetic reversal solution (0.1 mg atipamezole per kg body weight) via IP injection and place the mouse on a heating blanket set at approximately 37°C for recovery. Monitor the mice closely until fully awake. Let the mice ambulate freely to access food and water.
Monitor mice daily and remove skin staples after 2 weeks (see Note 15).
3.5. Analysis of vascularization and angiogenesis
Blood vessels are visualized with anti-PECAM (CD31) immunostaining (see Notes 18 and 19).
Harvest renal capsule transplanted femora and fix the tissue with 4% Paraformaldehyde fixative solution for 24h (see Notes 20 and 21).
Decalcify samples in 0.5M EDTA for 24h- 7 days on a rocking platform shaker at 4°C. Change EDTA solution every day (see Notes 22 and 23).
Dehydrate skeletal tissues by immersing tissue in graded ethanol series followed by xylene three times for 20 minutes each at room temperature (see Note 24).
Embed the tissue in paraffin at 58 °C.
Cut 5–7μm thick tissue sections using a rotary microtome. Float the sections in a 56 °C water bath and mount the sections onto microscope slides.
Dry the slides at room temperature for 1h and proceed with anti-PECAM immunostaining (see Notes 25 and 26).
Rehydrate sections by immersing the slides in Histoclear 2 times for 5 minutes each.
Immerse the slides in 100% Ethanol 2 times for 5 minutes each.
Immerse the slides in 95% Ethanol for 5 minutes.
Immerse the slides in 70% Ethanol for 5 minutes.
Immerse the slides with deionized H2O for 5 minutes.
Rehydrate the slides with 1x PBS for 5 minutes using a glass jar with lid.
Surround the tissue with a hydrophobic barrier using a barrier pen.
Block endogenous peroxidase activity with fresh 0.3% of hydrogen peroxide (H2O2) diluted in methanol for 45 minutes at room temperature (see Notes 27 and 28).
Proceed for enzymatic antigen retrieval step by incubating sections with ready to use Ficin solution for 5 minutes at room temperature (see Notes 29).
Wash slides 3 times for 5 minutes each in 1x PBS.
Block non-specific staining by incubating sections with 0.1M glycine solution diluted in 1x PBS for 60 s at room temperature (see Notes 30).
Wash slides 3 times for 5 minutes each in 1x PBS.
Block nonspecific staining by incubating sections with 5% nonfat powdered milk solution diluted in 1x PBS for 10 minutes at room temperature.
Wash slides 3 times for 5 minutes each in 1x PBS.
Block nonspecific staining by incubating sections with 0.1% ovalbumin solution diluted in 1x PBS for 10 minutes at room temperature.
Wash slides 3 times for 5 minutes each in 1x PBS.
Block nonspecific staining by incubating sections with 5% normal goat serum diluted in 1x PBS for 30 minutes at room temperature (see Notes 31).
Apply rat anti-PECAM primary antibodies solution at 1:50 diluted in serum blocking solution (5% normal goat serum diluted in 1x PBS) and incubate overnight at 4°C (see Notes 32 and 33).
Rinse 1 time with 1x PBS to drain the excess of primary antibodies.
Wash slides 3 times for 5 minutes each in 1x PBS.
Block non-specific staining by incubating sections with 5% normal goat serum diluted in 1x PBS for 30 min at room temperature (see Notes 31).
Apply goat biotinylated anti- rat secondary antibodies solution at 1:200 diluted in serum blocking solution (5% normal goat serum diluted in 1x PBS) and incubate for 1h at room temperature.
Rinse the slides once with 1x PBS to remove excess secondary antibody.
Wash the slides 3 times for 5 minutes each in 1x PBS.
Apply HRP- conjugated streptavidin solution at 1:100 diluted in serum blocking solution (5% normal goat serum diluted in 1x PBS) and incubate for 45 min at room temperature.
Rinse the slides once with 1x PBS to remove excess HRP-streptavidin solution.
Wash the slides three times for 10 minutes each in 1x PBS.
Apply DAB substrate working solution and develop for 30 s to 1 minute (see Notes 34–36).
Rinse the slides for 2 minutes with ddH2O.
Wash the slides three times for 5 minutes each in ddH2O.
Proceed to counterstaining by incubating sections with 0.01% Fast Green solution diluted in deionized H2O for 15 s at room temperature.
Dehydrate the sections by immersing tissue in 70% ethanol for 3 min at room temperature.
Immerse the slides in 95% ethanol for 3 min at room temperature.
Immerse the slides in 100% ethanol for 3 min at room temperature.
Immerse the slides in Histoclear solution for 5 mins at room temperature.
Apply a drop of Permount, and coverslip.
Let slides dry at room temperature.
Acknowledgements
This work was supported by INSERM ATIP-AVENIR, Osteosynthesis and Trauma Care Foundation, ANR-18-CE14–0033 and NIH-NIAMS R01AR072707 grants to CC.
Notes.
Use donor and host mice from the same genetic background to avoid graft rejection. Host mice should be preferably male as remodeling of the graft is accelerated in female hosts.
All surgical instruments and reagents must be sterile to avoid any risks of infection.
For the transplantation of stylopods and zeugopods, E14-E14.5 embryonic stage is the ideal time point as hypertrophic cartilage is well differentiated and will efficiently attract host blood vessels, but endogenous blood vessels have not invaded the cartilage yet and will not for another 24hrs. For other skeletal elements that are less advanced in their development, such as autopods, later embryonic stages may be more appropriate.
Avoid excessive compression on the uterine explant, which could deform and compromise the embryonic tissues.
Embryo can be handled by grasping the attached Reichert’s membrane as well as the ectoplacental cone using fine forceps.
Use the soft tissue surrounding the stylopod (cartilage femoral graft) to handle the embryonic tissue. At E14-E14.5 embryonic stage, the stylopod is very soft. Excessive compression may deform and compromise the normal development under the renal capsule.
At least one additional injection of analgesic is performed the day following the surgery. Please refer to your institutional guidelines concerning animal care and welfare. All of our procedures received approval from the Paris Descartes University Ethical committee.
The kidney is retroperitoneal. It is not necessary to cut into the peritoneum. The cut through the body wall should start just above the hip level and should be long enough (1–1.5 cm) for the kidney to be “popped out” but not longer to avoid the risk of it falling back into the body cavity during the procedure (Fig. 1). Avoid cutting major vessels and nerves.
It is important to keep the capsule moist during the entire process; otherwise it will be easily torn.
The size of the incision in the capsule is determined by the size of the graft, but it should not exceed 4mm as it may cause a loss of the graft (Fig. 1).
The L-shape glass should be manipulated under the capsule tangential to the surface of the kidney to avoid tearing the capsule. Great care should be taken while creating the pouch to not damage the kidney parenchyma which if damaged will bleed.
The skeletal elements should be transplanted with intact perichondrium to allow optimal vascularization and development. Some remaining soft tissues can be kept around the graft, as it will not interfere with bone development and growth.
Numerous treatments and manipulations can be applied to the graft prior to transplantation or at the time of transplantation (for example incubation in a solution of blocking antibody, or placing beads soaked in a protein solution adjacent to the graft under the kidney capsule) [16]
Several grafts can be transplanted in one kidney capsule depending on the length of the study (3 grafts for up to one week of development in the renal capsule, 2 grafts for up to 2 weeks, one graft for longer time points). Bilateral grafting is not recommended.
Potential adverse effects include infection, parenchyma bleeding and graft rejection. Although these effects occur very rarely, mice should be monitored daily following the transplantation.
Avoid pulling on bone transplant to expose the kidney.
Maintain one extremity of the bone with forceps to create fracture or cortical defect.
Femurs or other long bones grown under the renal capsule repair after a cortical defect or a fracture (Figure 2C–D, G–H, respectively) following a similar regeneration process as observed in adult bone[18].
PECAM immunostaining can be realized on uninjured transplanted embryonic femur (Figure 1d) or after a cortical defect or fracture (data not shown) following the same protocol.
The volume of fixative solution should be 50 times greater than the size of the immersed tissue to ensure a proper fixation of the tissue.
Avoid fixing the tissue for more than 24 hours since tissue antigens may either be masked or destroyed.
Decalcification using chelator reagents such as EDTA works by capturing the calcium ions from the bone. EDTA acts slowly but is compatible with many immunostaining protocols.
The time of decalcification varies from 24h till 7 days depending on the mineral density of the sample determined by the size of the skeletal element and the time point of harvest.
Paraffin is immiscible with water. Tissue must be dehydrated before adding paraffin wax.
Slides with paraffin-embedded sections can be stored either at room temperature or at 2–8 °C for several years in slide storage boxes. However, PECAM immunostaining should be performed within a week after sectioning for optimum results.
This PECAM immunostaining protocol can also be performed on cryo-embedded tissues (starting the procedure at step 12).
Some cells or tissues contain endogenous peroxidase. Using HRP conjugated antibody may result in high, non-specific background staining. Incubation with Peroxide (H2O2) suppresses endogenous peroxidase activity and therefore reduces background staining.
Hydrogen peroxide should be stored in the refrigerator and protected from sunlight in order to slow it’s thermal decomposition. Always use fresh H2O2 working solution.
The Ficin enzymatic antigen retrieval method serves as a proteolytic digestion to expose the antigenic sites that are covered when the tissue is fixated making antibody-antigen binding easier during the staining procedure.
The non-specific staining blocking step is most often performed just prior to incubating the sample with the primary and secondary antibodies. Non-specific staining blocking solution reduces the background signal produced by non-specific interaction of primary and secondary antibodies with proteins in the tissue section.
Serum is required in the blocking solution to block immunoglobulin Fc receptors present on cells in the section. The serum should be of the same species as the secondary antibody.
Overnight incubation at 4°C with primary antibodies allows proper and optimal specific binding of antibodies to tissue targets and reduces nonspecific background staining.
A negative control is critical for an accurate interpretation of the immunostaining results. A negative control could be using the incubation buffer with no primary antibody to identify non-specific staining of the secondary reagents. Additional controls can be employed to support the specificity of staining generated by the primary antibody. These include absorption controls, isotype-matched controls (for monoclonal primary antibodies), and tissue-type controls.
Upon reaction with HRP (horseradish peroxidase), DAB substrate will produce a brown colored deposit. Signal development might be monitored under microscope.
DAB is extremely carcinogenic. Necessary precautions should be taken (wear gloves and use only glass containers).
DAB is photosensitive: Keep the DAB working solution away from light and always use freshly prepared DAB working solution.
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