Fat-derived cells Enhance the Bone-Forming Capacity of Hypertrophic Cartilage Matrices


Treating particular bone defects or injuries present a substantial challenges for clinicians. The method of choice usually involves the use of an “autologous” bone graft (“autologous” simply means that the graft comes from the patient’s own bone). However, autologous bone grafts have plenty of limitations. For example, if a patient has a large enough bone defect, there is no way the orthopedist and take bone from a donor site without causing a good deal of risk to the donor site. Even with small bone grafts, so-called “donor site morbidity” remains a risk. Having said that, plenty of patients have had autologous bone grafts that have worked well, but larger bone injuries or defects are not treatable with autologous bone grafts.

The answer: bone substitute materials. Bone substitute materials include tricalcium phosphate, hydroxyapatite, cement, ceramics, bioglass, hydrogels, polylactides, PMMA or poly(methy methacrylate) and other acrylates,, and a cadre of commercially available granules, blocks, pastes, cements, and membranes. Some of these materials are experimental, but others do work, even if do not work every time. The main problem with bone substitute materials is that, well, they are not bone, and, therefore lack the intrinsic ability to induce the growth of new bone (so-called osteoinductive potential) and their ability to integrate into new bone is also a problem at times.

We must admit that a good deal of progress has been made in this area and it’s a good thing too. Many of our fabulous men and women-at-arms have returned home with severe injuries from explosives and wounds from large-caliber weapons that have shattered their bones. These courageous men and women have been the recipient of these technologies. However, the clinician is sometimes left asking herself, “can we do better?”

A new paper from the laboratories of Ivan Martin and Claude Jaquiery from the University Hospital of Basel suggests that we can. This paper appeared in Stem Cells Translational Medicine and describes the use of a hypertrophic cartilage matrix that was seeded with cells derived from the stromal vascular faction of fat to not only make bone in the laboratory, but to also heal skull defects in laboratory animals. While this work benefitted laboratory animals, it was performed with human cells and materials, which suggests that this technique, if it can be efficiently and cheaply scaled up, might be usable in human patients.
The two lead authors of this paper, Atanas Todorov and Matthias Kreutz and their colleagues made hypertrophic cartilage matrices from human bone marrow mesenchymal stem cells (from human donors) that were induced to make cartilage. Fortunately, protocols have been very well worked out and making cartilage plugs with chondrocytes that are enlarged (hypertrophic) is something that has been successfully done in many laboratories. After growing the mesenchymal stem cells in culture, half a million cells were induced to form cartilage with dexamethasone, ascorbic acid 2-phosphate, and the growth factor TGF-beta1. After three weeks, the cartilage plugs were subjected to hypertrophic medium, which causes the cartilage cells to enlarge.

Chondrocyte enlargement is a prolegomena to the formation of bone and during development, many of our long bones (femur, humerus, fibula, radius, etc.), initially form as cartilage exemplars that are replaced by bone as the chondrocytes enlarge. Ossification begins when a hollow cylinder forms in the center of the bone (known as the periosteal collar). The underlying chondrocytes degenerate and die, thus releasing the matrix upon which calcium phosphate crystals accrete. The primary ossification center commences with the calcification of the central shaft of the bone and erosion of the matrix by the invasion of a blood vessel. The blood vessels bring osteoprogenitor cells that differentiate into osteoblasts and begin to deposit the bone matrix.

Next, Todorov and his crew isolated the stromal vascular fraction from fat that was donated by 12 volunteers who had fat taken from them by means of liposuction. The fat is then minced, digested with enzymes, centrifuged, filtered and then counted. This remaining fraction is called the stromal vascular fraction or SVF, and it consists of a pastiche of blood vessel-forming cells, mesenchymal stem cells, and bone-forming cells (and probably a few other cells types too). These SVF cells were seeded onto the hypertrophic cartilage plugs and used for the experiments in this paper.

First, the SVF-seeded plugs were used to grow bone in laboratory rodents. The cartilage plugs were implanted into the backs for nude mice. Different cartilage plugs were used that had been seeded with gradually increasing number of SVF cells. The implanted plugs definitely made ectopic bone, but the amount of bone they made was directly proportional to the number of SVF cells with which they had been seeded. Staining experimental also showed that some of the newly-grown bone came from the implanted SVF cells.

Ectopic bone formation. Grafts based on devitalized hypertrophic cartilage pellets were embedded in fibrin gel without or with stromal vascular fraction cells from adipose tissue and implanted subcutaneously in nude mice. (A): Representative hematoxylin and eosin, Masson-Tri-Chrome, and Safranin-O (Saf-O) staining and in situ hybridization for human ALU sequences (dark blue = positive) after 12 weeks in vivo. Saf-O stainings are blue-green because of lack of glycosaminoglycans and counterstaining with fast green. Osteoid matrix and bone marrow are visible. Scale bars = 200 µm. (B): Stainings for metalloproteinase (MMP)13 and MMP9, as well as for the N-terminal neoepitope at the major MMP cleavage site (DIPEN) after 12 weeks in vivo (red/pink = positive). Scale bars = 50 µm. +, osteoid matrix; ⋆, bone marrow. Abbreviations: ALU, Arthrobacter luteus; H&E, hematoxylin and eosin; Masson, Masson’s trichrome; MMP, metalloproteinase; Saf-O, Safranin-O; SVF, stromal vascular fraction.
Ectopic bone formation. Grafts based on devitalized hypertrophic cartilage pellets were embedded in fibrin gel without or with stromal vascular fraction cells from adipose tissue and implanted subcutaneously in nude mice. (A): Representative hematoxylin and eosin, Masson-Tri-Chrome, and Safranin-O (Saf-O) staining and in situ hybridization for human ALU sequences (dark blue = positive) after 12 weeks in vivo. Saf-O stainings are blue-green because of lack of glycosaminoglycans and counterstaining with fast green. Osteoid matrix and bone marrow are visible. Scale bars = 200 µm. (B): Stainings for metalloproteinase (MMP)13 and MMP9, as well as for the N-terminal neoepitope at the major MMP cleavage site (DIPEN) after 12 weeks in vivo (red/pink = positive). Scale bars = 50 µm. +, osteoid matrix; ⋆, bone marrow. Abbreviations: ALU, Arthrobacter luteus; H&E, hematoxylin and eosin; Masson, Masson’s trichrome; MMP, metalloproteinase; Saf-O, Safranin-O; SVF, stromal vascular fraction.

In the second experiment, Todorov and Kreutz used these SVF-seeded cartilage plugs to repair skull lesions in rats. Once again, the quantity of bone produced was directly proportional to the number of SVFs seeded into the cartilage matrices prior to implantation. In both experiments, the density of SVF cells positively correlates with the bone-forming cells in the grafts and the percentage of SVF-derived blood vessel-forming cells correlates with the amount of deposited mineralized matrix.

Bone repair capacity. Devitalized hypertrophic cartilage pellets were embedded in fibrin gel without or with stromal vascular fraction (SVF) cells from adipose tissue and implanted in rat calvarial defects. (A): Mineralized volume quantified by microtomography (n = 9 grafts assessed per group). (B): Bone area assessed in histological sections, expressed as percentage of total defect area (n = at least 24 sections assessed per group). ∗∗∗∗, p < .0001. (C, D): Representative three-dimensional microtomography reconstructions (left) and hematoxylin/eosin (H&E) staining (right) of the calvarial defects filled with devitalized grafts, implanted without (C) or with (D) activation by SVF cells after 4 weeks. Dotted circles indicate the defect borders (4 mm diameter). Scale bars = 500 µm. (E): Microtomography (left) and H&E staining (middle and right) displaying the bridging between hypertrophic matrix and bone of the calvarium, or the fusion of single pellets (right) in activated grafts. White bar = 850 µm; black bars = 500 µm. Dotted lines indicate the edge of the calvarium. (F): In situ hybridization for Arthrobacter luteus sequences showing the presence of human cells (dark blue, positive) in the explants. Scale bar = 200 µm. Abbreviations: C, calvarium; dev, fibrin gel without stromal vascular fraction; dev + SVF, fibrin gel with stromal vascular fraction; P, hypertrophic matrix; SVF, stromal vascular fraction.
Bone repair capacity. Devitalized hypertrophic cartilage pellets were embedded in fibrin gel without or with stromal vascular fraction (SVF) cells from adipose tissue and implanted in rat calvarial defects. (A): Mineralized volume quantified by microtomography (n = 9 grafts assessed per group). (B): Bone area assessed in histological sections, expressed as percentage of total defect area (n = at least 24 sections assessed per group). ∗∗∗∗, p < .0001. (C, D): Representative three-dimensional microtomography reconstructions (left) and hematoxylin/eosin (H&E) staining (right) of the calvarial defects filled with devitalized grafts, implanted without (C) or with (D) activation by SVF cells after 4 weeks. Dotted circles indicate the defect borders (4 mm diameter). Scale bars = 500 µm. (E): Microtomography (left) and H&E staining (middle and right) displaying the bridging between hypertrophic matrix and bone of the calvarium, or the fusion of single pellets (right) in activated grafts. White bar = 850 µm; black bars = 500 µm. Dotted lines indicate the edge of the calvarium. (F): In situ hybridization for Arthrobacter luteus sequences showing the presence of human cells (dark blue, positive) in the explants. Scale bar = 200 µm. Abbreviations: C, calvarium; dev, fibrin gel without stromal vascular fraction; dev + SVF, fibrin gel with stromal vascular fraction; P, hypertrophic matrix; SVF, stromal vascular fraction.

This is not the first time scientists have proposed the use of cartilage plugs to induce the formation of new bone. Van der Stok and others and Bahney and colleagues were able to repair segmental bone defects in laboratory rodents. Is this technique transferable to human patients? Possibly. Hypertrophic cartilage is relatively easy to make and it is completely conceivable that hypertrophic cartilage wedges can be sold as “off-the-shelf” products for bone treatments. SVF can also be derived from the patient or can be derived from donors.

Furthermore, the protocols in this paper all used human cells and grew the products in immunodeficient rats and mice. Therefore, in addition to scaling this process up, we have a potentially useful protocol that might very well be adaptable to the clinic.

The efficacy of this technique must be confirmed in larger animal model system before human trials can be considered. Hopefully human trials are in the future for this fascinating technique.

Stem Cells from Fat Improve Blood Vessel Responses after Injury


When tissues are injured, the blood vessels that feed them are often shocked and damaged as well. “Vasoactivity” refers the ability of blood vessels to dilate or constrict. When tissues are harmed, blood vessels tend to shrink in order to squelch blood loss at the site of damage. This same response, however, and deprive the damaged tissues of much-needed oxygen and lead to “ischemia,” which is the insufficient supply of blood and oxygen to an organ.

James B. Hoying and his colleagues at the University of Louisville in Kentucky used the “stromal vascular fraction” or SVF from fat in order to treat damaged blood vessels to determine if they could mitigate the decrease in vasoactivity as a result of injury.

The SVF refers to the stem fraction from fat after the fat has been minced, digested with enzymes, and centrifuged (it’s more complicated than that, but this is a short summary). The cells that remain include mesenchymal stromal cells, growth factors, immune cells, pre-fat cells and fat cells, blood-cell-making stem cells, and blood vessel-making cells (endothelial cells). The SVF, therefore, contains a cocktail of cell types and growth factors that are available for regenerative medicine.

Hoying and his team discovered that when fluorescent SVF cells were injected into a laboratory mouse, they cells distributed to a variety of tissues. Further and more detailed examinations showed that these cells were finding their ways into organs and tissues because they traveled through the circulatory system and could be found in the walls of blood vessels.

Next, the composition of the SVF was examined. About 25% of the cells in the SVF were endothelial cells, 22% were various types of blood cells, 20% were “CD11b” cells, which means that these cells had a protein called CD11b on their cell surfaces. That protein was formerly canned “Mac-1” and is was normally found on the surfaces of phagocytic cells called macrophages. Therefore, this CD11b faction could very well be macrophages, but other cell types have this protein on their surfaces as well.

Macrophages

Next, Hoying and others injected these SVF-derived cells into the large leg vein (saphenous) of the leg. Such injections consistently caused these vessels to relax and dilate. Secondly, the SVF-derived cells caused the vessels to relax in a CD11b-dependent manner. In other words, the more CD11b cells there were in the SVF preparation, the greater the amount of vasoactivity they induced. If fractions were depleted of their CD11b, they could not induce vasoactivity.

When Hoying and others examined the SVF-treated vessels, they saw CD11b+ cells lining the inner layer of the vessels. Thus these cells were getting right up against the inside of the vessel and signaling to the underlying smooth muscle to relax.

Finally, Hoying and others clamped the saphenous veins of laboratory mice. Such clamping will induce tissue ischemia and inflammation in the vessels. Can SVF cells calm the inflammation and make the vessels more vasoactive? The answer is an unqualified yes.  See below.  The veins from SVF-treated animals show signficantly greater dilation than those from untreated or CD11b-depleted SVF-treated animals.

SVF cells relax vasomotor tone in inflamed saphenous arteries. (A): Schematic of the experimental plan involving the cell treatment of locally inflamed (cuffed) saphenous arteries of mice injected with syngeneic adipose SVF cells constitutively expressing luciferase and GFP reporter transgenes or SVF cells depleted of CD11b+ cells. Also shown is a gross view and a histological cross-section of a cuffed saphenous artery. (B): Hematoxylin and eosin-stained histological cross-sections of normal (noncuffed) and cuffed mouse saphenous arteries untreated or injected with SVF cells or SVF-11bΔ cells. Rightmost panels: Higher magnification images of the adjacent images. Scale bars = 25 μm in the left and right columns and 100 μm in the middle column. (C): Lumen diameters of untreated (n = 9) and cell-injected cuffed saphenous arteries measured from histological sections. Cell treatments included complete SVF cell isolates (C + SVF, n = 7) or SVF isolates depleted of CD11b+ cells (C + SVF-11bΔ, n = 7). Data are shown as the mean ± SEM; ∗, p < .05, determined by one-way analysis of variance. (D): Visualization of luciferase-positive SVF cells within histological paraffin sections of cuffed saphenous arteries from untreated, SVF-injected, and SVF-11bΔ-injected mice via immunostaining for luciferase. Brown stain indicates positive luciferase immune-staining and the presence of SVF cells. Tissues were harvested 1 week after cell delivery. Scale bars = 100 μm. Abbreviations: C, cuff; GFP, green fluorescent protein; PE, polyethylene; SVF, stromal vascular fraction; SVF-11bΔ, CD11b+ cell-depleted adipose SVF cells.
SVF cells relax vasomotor tone in inflamed saphenous arteries. (A): Schematic of the experimental plan involving the cell treatment of locally inflamed (cuffed) saphenous arteries of mice injected with syngeneic adipose SVF cells constitutively expressing luciferase and GFP reporter transgenes or SVF cells depleted of CD11b+ cells. Also shown is a gross view and a histological cross-section of a cuffed saphenous artery. (B): Hematoxylin and eosin-stained histological cross-sections of normal (noncuffed) and cuffed mouse saphenous arteries untreated or injected with SVF cells or SVF-11bΔ cells. Rightmost panels: Higher magnification images of the adjacent images. Scale bars = 25 μm in the left and right columns and 100 μm in the middle column. (C): Lumen diameters of untreated (n = 9) and cell-injected cuffed saphenous arteries measured from histological sections. Cell treatments included complete SVF cell isolates (C + SVF, n = 7) or SVF isolates depleted of CD11b+ cells (C + SVF-11bΔ, n = 7). Data are shown as the mean ± SEM; ∗, p < .05, determined by one-way analysis of variance. (D): Visualization of luciferase-positive SVF cells within histological paraffin sections of cuffed saphenous arteries from untreated, SVF-injected, and SVF-11bΔ-injected mice via immunostaining for luciferase. Brown stain indicates positive luciferase immune-staining and the presence of SVF cells. Tissues were harvested 1 week after cell delivery. Scale bars = 100 μm. Abbreviations: C, cuff; GFP, green fluorescent protein; PE, polyethylene; SVF, stromal vascular fraction; SVF-11bΔ, CD11b+ cell-depleted adipose SVF cells.

This an interesting and exciting finding not only because of the ability of these fat-based cells to maintain vasoactivity even under pro-inflammatory conditions, but because it is the macrophage cell population that is doing the work.  In most stem preparations, macrophages are excluded.  This paper shows that macrophages have greater therapeutic capabilities than previously thought, and should also be tested for sanative properties.