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22 pages, 31372 KiB  
Article
Silactins and Structural Diversity of Biosilica in Sponges
by Hermann Ehrlich, Alona Voronkina, Konstantin Tabachniсk, Anita Kubiak, Alexander Ereskovsky and Teofil Jesionowski
Biomimetics 2024, 9(7), 393; https://doi.org/10.3390/biomimetics9070393 - 27 Jun 2024
Viewed by 1251
Abstract
Sponges (phylum Porifera) were among the first metazoans on Earth, and represent a unique global source of highly structured and diverse biosilica that has been formed and tested over more than 800 million years of evolution. Poriferans are recognized as a unique archive [...] Read more.
Sponges (phylum Porifera) were among the first metazoans on Earth, and represent a unique global source of highly structured and diverse biosilica that has been formed and tested over more than 800 million years of evolution. Poriferans are recognized as a unique archive of siliceous multiscaled skeletal constructs with superficial micro-ornamentation patterned by biopolymers. In the present study, spicules and skeletal frameworks of selected representatives of sponges in such classes as Demospongiae, Homoscleromorpha, and Hexactinellida were desilicified using 10% HF with the aim of isolating axial filaments, which resemble the shape and size of the original structures. These filaments were unambiguously identified in all specimens under study as F-actin, using the highly specific indicators iFluor™ 594-Phalloidin, iFluor™ 488-Phalloidin, and iFluor™ 350-Phalloidin. The identification of this kind of F-actins, termed for the first time as silactins, as specific pattern drivers in skeletal constructs of sponges opens the way to the fundamental understanding of their skeletogenesis. Examples illustrating the biomimetic potential of sophisticated poriferan biosilica patterned by silactins are presented and discussed. Full article
(This article belongs to the Special Issue Advances in Biomimetics: The Power of Diversity)
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Figure 1

Figure 1
<p>Digital microscopy imagery of <span class="html-italic">Ephydatia muelleri</span> freshwater demosponge oxeas with symmetrical tips after removal of organic material using HNO<sub>3</sub> treatment. Desilicification of such spicules with HF led to isolation of organic axial filaments, which were identified as F-actin (see <a href="#biomimetics-09-00393-f002" class="html-fig">Figure 2</a>).</p>
Full article ">Figure 2
<p>Bright field (<b>a</b>,<b>c</b>,<b>e</b>) and fluorescence microscopy imagery of the axial filaments obtained after demineralization of oxeas of <span class="html-italic">E. muelleri</span> freshwater demosponge using HF under the conditions of the “sliding drop technique” [<a href="#B24-biomimetics-09-00393" class="html-bibr">24</a>] and stained with 594-Phalloidin (<b>b</b>); also with 488-Phalloidin (<b>d</b>) and 350-Phalloidin (<b>f</b>) for comparison.</p>
Full article ">Figure 3
<p>Bright field (<b>a</b>) and fluorescence microscopy (<b>b</b>) imagery of the axial filaments of <span class="html-italic">E. muelleri</span> demosponge oxeas isolated in bulk after HF treatment, dialyzed, and finally stained with 594-Phalloidin. (<b>c</b>) SDS-PAGE: arrows indicating the actin (45 kDa) and silicatein (25 kDa) bands well visible after both Coomassie blue (left gel) and silver reagent (right gel) staining of the axial filaments sample <span class="html-italic">of E. muelleri</span> under study.</p>
Full article ">Figure 4
<p>Digital microscopy images of organic-freed acantoxeas and oxeas isolated from the endemic <span class="html-italic">O. rotunda</span> freshwater demosponge. Demineralization of such spicules with HF led to isolation of organic axial filaments, which were identified using such diverse phalloidin indicators as F-actin (see <a href="#biomimetics-09-00393-f005" class="html-fig">Figure 5</a>).</p>
Full article ">Figure 5
<p>Bright field (<b>a</b>,<b>c</b>,<b>e</b>) and fluorescence microscopy imagery of the axial filaments obtained after demineralization of acantoxeas and oxeas of the <span class="html-italic">O. rotunda</span> freshwater demosponge with HF under the conditions of the “sliding drop technique” [<a href="#B24-biomimetics-09-00393" class="html-bibr">24</a>] and stained for comparative purposes with 594-Phalloidin (<b>b</b>), 488-Phalloidin (<b>d</b>), and 350-Phalloidin (<b>f</b>).</p>
Full article ">Figure 5 Cont.
<p>Bright field (<b>a</b>,<b>c</b>,<b>e</b>) and fluorescence microscopy imagery of the axial filaments obtained after demineralization of acantoxeas and oxeas of the <span class="html-italic">O. rotunda</span> freshwater demosponge with HF under the conditions of the “sliding drop technique” [<a href="#B24-biomimetics-09-00393" class="html-bibr">24</a>] and stained for comparative purposes with 594-Phalloidin (<b>b</b>), 488-Phalloidin (<b>d</b>), and 350-Phalloidin (<b>f</b>).</p>
Full article ">Figure 6
<p>Bright field images of <span class="html-italic">Biemna</span> sp. marine demosponge spicules (<b>a</b>) and their axial filaments isolated in bulk after HF treatment (<b>b</b>). Fluorescence microscopy image (<b>c</b>) of dialyzed axial filaments stained with 594-Phalloidin showing the red color characteristic for phalloidin labeled F-actin. (<b>d</b>) SDS-PAGE: bands indicating the presence of both actin (45 kDa) and silicateins (25 kDa) in axial filaments extracted after HF-based desilicification of <span class="html-italic">Biemna</span> sp. remain well visible after silver reagent staining in two selected samples. For comparison, see <a href="#biomimetics-09-00393-f003" class="html-fig">Figure 3</a>c.</p>
Full article ">Figure 7
<p>HF-based desilicification of the <span class="html-italic">S. domuncula</span> marine demosponge tylostyle (<b>a</b>) led to isolation of the axial filaments (<b>b</b>), which were identified as F-actin using 594-Phalloidin staining (fluorescence microscopy image (<b>c</b>)). F-actin branching of the axial filament fragment within the “club-like” structure is well visible. See also <a href="#app1-biomimetics-09-00393" class="html-app">Figure S10</a>.</p>
Full article ">Figure 8
<p>Bright field images of axial filaments isolated from spicules of marine demosponges <span class="html-italic">P. arctica</span> (<b>a</b>), <span class="html-italic">S. borealis</span> (<b>c</b>), and <span class="html-italic">T. norvegica</span> (<b>e</b>) using HF-based treatment as presented above (see <a href="#biomimetics-09-00393-f005" class="html-fig">Figure 5</a>). Right: fluorescence microscopy images of respective species’ axial filaments stained with 594-Phalloidin (<b>b</b>), 488-Phalloidin (<b>d</b>), and 350-Phalloidin (<b>f</b>).</p>
Full article ">Figure 8 Cont.
<p>Bright field images of axial filaments isolated from spicules of marine demosponges <span class="html-italic">P. arctica</span> (<b>a</b>), <span class="html-italic">S. borealis</span> (<b>c</b>), and <span class="html-italic">T. norvegica</span> (<b>e</b>) using HF-based treatment as presented above (see <a href="#biomimetics-09-00393-f005" class="html-fig">Figure 5</a>). Right: fluorescence microscopy images of respective species’ axial filaments stained with 594-Phalloidin (<b>b</b>), 488-Phalloidin (<b>d</b>), and 350-Phalloidin (<b>f</b>).</p>
Full article ">Figure 9
<p>Polybranched microarchitecture of <span class="html-italic">Geodia cydonium</span> marine demosponge spicules are well visible, especially in SEM image (<b>a</b>). Both types of spicules, radially oriented sterrasters as well as linear megascleres after demineralization using HF, show the presence of correspondingly structured axial filaments, which have been identified as F-actin-based filaments through specific staining with 594-Phalloidin for <span class="html-italic">Erylus granularis</span> (Geodiidae) (<b>b</b>) and 350-Phalloidin for <span class="html-italic">G. cydonium</span> (<b>c</b>).</p>
Full article ">Figure 10
<p>Cell-free 18 cm-long skeleton of <span class="html-italic">E. aspergillum</span> glass sponge (<b>a</b>) used in the study. Bright field (<b>b</b>,<b>d</b>,<b>f</b>) images of selected skeletal fragments demineralized with HF, with characteristic square geometry of organic filaments. These filaments are identified as F-actin structures using fluorescence microscopy after staining with 488-Phalloidin (<b>c</b>), 350-Phalloidin (<b>e</b>), and 594-Phalloidin (<b>g</b>), corresponding to the bright field images.</p>
Full article ">Figure 11
<p>Schematic view of F-actin growth models previously described in the literature vs. siliceous structures observed in Hexactinellida sponges: (<b>a</b>) branching of bovine actin [<a href="#B41-biomimetics-09-00393" class="html-bibr">41</a>]; (<b>b</b>) uncinate spicule of Tretodictyidae sponge; (<b>c</b>) cortical axon branching [<a href="#B47-biomimetics-09-00393" class="html-bibr">47</a>]; (<b>d</b>) discoscopule spicule (Hexactinellida, Tretodictyidae); (<b>e</b>) actin branching in lamellipodia of <span class="html-italic">Xenopus laevis</span> keratocytes [<a href="#B48-biomimetics-09-00393" class="html-bibr">48</a>]; (<b>f</b>) oxyhexactin spicule (Hexactinellida, Euretidae); (<b>g</b>) astrocytes actin branching in rat nervous system [<a href="#B49-biomimetics-09-00393" class="html-bibr">49</a>]; (<b>h</b>) discohexaster spicule (Hexactinellidae, Tretodictyidae); (<b>i</b>) endothelial actine cytoskeleton in mouse retina [<a href="#B46-biomimetics-09-00393" class="html-bibr">46</a>]; (<b>j</b>) farreoid skeleton (Hexactinellida, Farreidae, <span class="html-italic">Farrea</span> sp.); (<b>k</b>) honeycomb actin structures in mouse lenses [<a href="#B50-biomimetics-09-00393" class="html-bibr">50</a>] and within diatom frustule [<a href="#B26-biomimetics-09-00393" class="html-bibr">26</a>]; (<b>l</b>) honeycomb skeleton of Aphrocallistidae glass sponge <span class="html-italic">Aphrocallistes</span> sp. (see also [<a href="#B22-biomimetics-09-00393" class="html-bibr">22</a>]).</p>
Full article ">Figure 12
<p>State-of-the-art overview on silactins’ distribution within skeletal structures of three poriferan classes. (<b>a</b>) Unique radial orientation of silactin microfilaments of <span class="html-italic">Pachymatisma normani</span> (Geodiidae) marine demosponge became well visible after HF-based desilicification of corresponding sterrasters and following staining with 488-phalloidin marker.</p>
Full article ">
21 pages, 26525 KiB  
Article
Transcriptomic Responses of a Lightly Calcified Echinoderm to Experimental Seawater Acidification and Warming during Early Development
by Ye Zhao, Mingshan Song, Zhenglin Yu, Lei Pang, Libin Zhang, Ioannis Karakassis, Panagiotis D. Dimitriou and Xiutang Yuan
Biology 2023, 12(12), 1520; https://doi.org/10.3390/biology12121520 - 13 Dec 2023
Cited by 1 | Viewed by 1408
Abstract
Ocean acidification (OA) and ocean warming (OW) are potential obstacles to the survival and growth of marine organisms, particularly those that rely on calcification. This study investigated the single and joint effects of OA and OW on sea cucumber Apostichopus japonicus larvae raised [...] Read more.
Ocean acidification (OA) and ocean warming (OW) are potential obstacles to the survival and growth of marine organisms, particularly those that rely on calcification. This study investigated the single and joint effects of OA and OW on sea cucumber Apostichopus japonicus larvae raised under combinations of two temperatures (19 °C or 22 °C) and two pCO2 levels (400 or 1000 μatm) that reflect the current and end-of-21st-century projected ocean scenarios. The investigation focused on assessing larval development and identifying differences in gene expression patterns at four crucial embryo–larval stages (blastula, gastrula, auricularia, and doliolaria) of sea cucumbers, using RNA-seq. Results showed the detrimental effect of OA on the early development and body growth of A. japonicus larvae and a reduction in the expression of genes associated with biomineralization, skeletogenesis, and ion homeostasis. This effect was particularly pronounced during the doliolaria stage, indicating the presence of bottlenecks in larval development at this transition phase between the larval and megalopa stages in response to OA. OW accelerated the larval development across four stages of A. japonicus, especially at the blastula and doliolaria stages, but resulted in a widespread upregulation of genes related to heat shock proteins, antioxidant defense, and immune response. Significantly, the negative effects of elevated pCO2 on the developmental process of larvae appeared to be mitigated when accompanied by increased temperatures at the expense of reduced immune resilience and increased system fragility. These findings suggest that alterations in gene expression within the larvae of A. japonicus provide a mechanism to adapt to stressors arising from a rapidly changing oceanic environment. Full article
(This article belongs to the Section Marine Biology)
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<p>Effect of OA and/or OW on larval development and growth of larvae of <span class="html-italic">A. japonicus.</span> (<b>a</b>) Body length of <span class="html-italic">A. japonicus</span> larvae (blastula: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 8.724, <span class="html-italic">p</span> = 0.019; gastrula: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 6.017, <span class="html-italic">p</span> = 0.01; auricularia: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 113.820, <span class="html-italic">p</span> &lt; 0.0001; doliolaria: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 800.676, <span class="html-italic">p</span> &lt; 0.0001). (<b>b</b>) Stage duration of <span class="html-italic">A. japonicus</span> larvae (blastula: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 114.723, <span class="html-italic">p</span> = 0.525; gastrula: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 9.333, <span class="html-italic">p</span> = 0.006; auricularia: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 521.214, <span class="html-italic">p</span> = 0.002; doliolaria: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 84.000, <span class="html-italic">p</span> &lt; 0.0001). (<b>c</b>) Growth rate of <span class="html-italic">A. japonicus</span> larvae (blastula: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 20.506, <span class="html-italic">p</span> = 0.115; gastrula: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 4.370, <span class="html-italic">p</span> = 0.027; auricularia: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 21.516, <span class="html-italic">p</span> &lt; 0.0001; doliolaria: <span class="html-italic">n</span> = 4, LSD, F<sub>3,12</sub> = 132.188, <span class="html-italic">p</span> &lt; 0.0001). OA: ocean acidification; OW: ocean warming. The “**” symbol above the bar chart indicates a significant difference (<span class="html-italic">p</span> &lt; 0.01), and the “*” symbol above the bar chart indicates a significant difference (<span class="html-italic">p</span> &lt; 0.05).</p>
Full article ">Figure 2
<p>Principal component analysis (PCoA) of samples in four treatments at four larval stages of <span class="html-italic">A. japonicus</span> based on the Bray–Curtis distance.</p>
Full article ">Figure 3
<p>Volcano plots of DEGs for comparison of OA vs. CON (<b>a</b>–<b>d</b>), OW vs. CON (<b>e</b>–<b>h</b>), and OWA vs. CON (<b>i</b>–<b>l</b>) at four larval stages of <span class="html-italic">A. japonicus.</span> The principle “a <span class="html-italic">p</span>-value ≤ 0.05 and the absolute value of log<sub>2</sub>Ratio ≥ 1” was used as a threshold to screen DEGs. DEGs: differentially expressed genes; OWA: ocean warming and acidification; CON: control group.</p>
Full article ">Figure 4
<p>Histogram of GO pathways enriched by DEGs. The gene numbers enriched in GO terms of the categories “Biological Process”, “Cellular component” and “Molecular function” are shown. GO: gene ontology.</p>
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<p>Bubble chart of top 20 KEGG pathways enriched by DEGs. Significantly enriched KEGG pathways are marked in bold and identified with q Value ≤ 0.05. KEGG: Kyoto Encyclopedia of Genes and Genomes.</p>
Full article ">Figure 6
<p>qRT-PCR results of eight DEGs in <span class="html-italic">A. japonicus</span> larvae under OA and/or OW. Data are represented as mean ± SD. The “**” symbol above the bar chart indicates a significant difference (<span class="html-italic">p</span> &lt; 0.01), and the “*” symbol above the bar chart indicates a significant difference (<span class="html-italic">p</span> &lt; 0.05). qRT-PCR: quantitative real-time polymerase chain reaction.</p>
Full article ">
17 pages, 4176 KiB  
Article
Modeling the Differentiation of Embryonic Limb Chondroprogenitors by Cell Death and Cell Senescence in High Density Micromass Cultures and Their Regulation by FGF Signaling
by Cristina Duarte-Olivenza, Juan M. Hurle, Juan A. Montero and Carlos I. Lorda-Diez
Cells 2023, 12(1), 175; https://doi.org/10.3390/cells12010175 - 31 Dec 2022
Cited by 2 | Viewed by 1836
Abstract
Considering the importance of programmed cell death in the formation of the skeleton during embryonic development, the aim of the present study was to analyze whether regulated cell degeneration also accompanies the differentiation of embryonic limb skeletal progenitors in high-density tridimensional cultures (micromass [...] Read more.
Considering the importance of programmed cell death in the formation of the skeleton during embryonic development, the aim of the present study was to analyze whether regulated cell degeneration also accompanies the differentiation of embryonic limb skeletal progenitors in high-density tridimensional cultures (micromass cultures). Our results show that the formation of primary cartilage nodules in the micromass culture assay involves a patterned process of cell death and cell senescence, complementary to the pattern of chondrogenesis. As occurs in vivo, the degenerative events were preceded by DNA damage detectable by γH2AX immunolabeling and proceeded via apoptosis and cell senescence. Combined treatments of the cultures with growth factors active during limb skeletogenesis, including FGF, BMP, and WNT revealed that FGF signaling modulates the response of progenitors to signaling pathways implicated in cell death. Transcriptional changes induced by FGF treatments suggested that this function is mediated by the positive regulation of the genetic machinery responsible for apoptosis and cell senescence together with hypomethylation of the Sox9 gene promoter. We propose that FGF signaling exerts a primordial function in the embryonic limb conferring chondroprogenitors with their biological properties. Full article
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<p>(<b>A</b>–<b>A’</b>) In situ hybridization of the embryonic leg bud at stage 25, showing the expression of <span class="html-italic">HoxA13</span>. <b>A’</b> the tissue selected for the experiments is illustrated in red. (<b>B</b>–<b>D</b>) are dark field low magnification views of micromasses cultured for 24 (<b>B</b>), 48 (<b>C</b>), and 72 h (<b>D</b>). (<b>B’</b>–<b>D’</b>) illustrate the same cultures after Alcian blue cartilage staining. Note the appearance of Alcian blue-positive nodules on Day 2 of culture. (<b>E</b>) 3-day-old micromass section immunolabeled for SOX9. (<b>F</b>) Flow cytometry histogram showing the cell cycle of 2 days cultures grown in DMEM. Scale bar in <b>B</b>–<b>D’</b> = 1 mm; scale bar in E = 100 µm.</p>
Full article ">Figure 2
<p>(<b>A</b>) Semithin section of a 3-day-old culture showing the presence of dark apoptotic cells (arrows) and vacuolated senescent cells (arrowheads) around a chrondrogenic nodule (Ch). (<b>B</b>) Detailed view of a micromass tissue showing senescent cells positive for SAβ-gal histochemistry (eosin counterstaining). (<b>C</b>) Graphic representation of the rate of cell death evaluated by flow cytometry during the first 3 days of culture grown in DMEM-only. (<b>D</b>) Total tissue mass of micromass cultures at Days 1, 3, and 7 evaluated by MTT staining. (<b>E</b>) Low-magnification view of 1-day culture after SAβ-gal histochemistry (dark blue staining) showing the distribution of senescent cells. (<b>F</b>) Perinodular restricted distribution of senescent cells positive for SAβ-gal in a 3-day-old micromass. (<b>G</b>) Optical section of 3-day-old micromass showing the arrangement of apoptosis (green TUNEL labeling) in the perinodular tissue. Note the intense positivity for SOX9 (red labeling) in chondrogenic aggregates. (<b>H</b>) A 3-day-old micromass section immunolabeled for γH2AX (green) and SOX9 (red) to show the preferential distribution of γH2AX in perinodular tissue. Note the very reduced nuclear yH2AX labeling in the aggregated cells highly positive for SOX9 (arrows). (<b>I</b>–<b>I”</b>) Detailed view of a SOX9-positive cell (<b>I</b>) with a couple of dots positive for γH2AX (<b>I’</b>). (<b>I”</b>) is the merged image. <b>(J</b>–<b>J”)</b> Detailed view of an internodular cell with poor SOX9 labeling (<b>J</b>) but massive γH2AX labeling (<b>J’</b>). (<b>J”</b>) is the merged image (<b>J”</b>). (<b>K</b>–<b>L</b>) Single channel and merged image of double labeling with γH2AX(green) and phalloidin (red) showing the aligned cytoplasmic actin filaments in healthy progenitors in contrast to the irregular aggregation of actin clumps (arrow) in the cytoplasm of cells highly positive for γH2AX (<b>K</b>). Scale bar in A and H = 20 µm; scale bar in B = 25 µm; scale bar in E and F = 300 µm; scale bar in G = 100 µm; scale bar I-J-K-L= 5 µm.</p>
Full article ">Figure 3
<p>(<b>A</b>–<b>A’</b>) Graphic representation of the rate of cell death evaluated by flow cytometry in cultures treated for 2 days with 25 (light columns) or 50 ng/mL of FGF2 (red columns) to compare the effects of a continuous treatment with FGF (+FGF) and the effect of FGF withdrawal (-FGF) for 3 h (<b>A</b>) or 6 h (<b>A’</b>). The rate of cell death in the control untreated micromasses was considered 100 and is indicated by the dotted line. (<b>B</b>–<b>B’</b>) Confocal view of the perinodular arrangement of TUNEL-positive cells (green labeling) in a labeled control 2-days culture. The sample is also labeled with phalloidin (red labeling). (<b>B</b>) is a merged image, and (<b>B’</b>) shows only the green channel. (<b>C</b>–<b>C’</b>) Confocal view showing the widespread arrangement of TUNEL-positive cells (green labeling) after 3 h of FGF withdrawal in micromasses treated for 2 days with FGF2 (25 ngr/mL). Red labeling corresponds to phalloidin. (<b>C</b>) is a merged image, and (<b>C’</b>) shows only the green channel. (<b>D</b>,<b>E</b>) Detailed view of the SAβ-gal arrangement in control (<b>D</b>) and experimental micromasses subjected to FGF-withdrawal (<b>E</b>). Note the perinodular distribution of SAβ-gal in the control in contrast with the intranodular distribution in the experimental culture. (<b>F</b>) Graphic representation of the rate of cell death evaluated by flow cytometry in 2-day micromasses subjected to treatments with SU5402 (800 ng/mL), SU5402 plus 25 ng/mL of FGF2, U0126 (7.6 ng/mL), and U0126 plus FGF2. Scale bar in B−C´ = 250 µm; scale bar in D−E = 50 µm. * <span class="html-italic">p</span> &lt; 0.05 (versus control); ##, <span class="html-italic">p</span> &lt; 0.01 (versus SU5402+FGF2).</p>
Full article ">Figure 4
<p>(<b>A</b>,<b>B</b>) Changes in the rate of cytosine methylation of the <span class="html-italic">Scleraxis</span> (<b>A</b>) and <span class="html-italic">Sox9</span> (<b>B</b>) promoters during the first three days of micromass culture. (<b>C</b>,<b>D</b>) Changes in the rate of cytosine methylation of the <span class="html-italic">Sox9</span> (<b>C</b>) and <span class="html-italic">Scleraxis</span> (<b>D</b>) promoters during the first two days of culture induced by the addition of FGF2 (25 ng/mL) to the culture medium. * <span class="html-italic">p</span> &lt; 0.05; *** <span class="html-italic">p</span> &lt; 0.001.</p>
Full article ">Figure 5
<p>(<b>A</b>–<b>C</b>) In situ hybridizations of 2-day-old micromass cultures showing the expression of <span class="html-italic">Bmp7</span> (<b>A</b>), <span class="html-italic">Wnt5a</span> (<b>B</b>), and <span class="html-italic">Dkk1</span> (<b>C</b>). (<b>D</b>) Graphic representation of the rate of cell death evaluated by flow cytometry in cultures treated for 2 days with BMP7 (200 ng/mL); BMP7 plus FGF2 (25 ng/mL); NOGGIN (200 ng/mL); and NOGGIN plus FGF2 (25 ng/mL). The rate of cell death in untreated control cultures was considered to be 100% and indicated by the dotted line. (<b>E</b>) Graphic representation of the rate of cell death evaluated by flow cytometry in cultures treated for 2 days with WNT5A (100 ng/mL), WNT5A plus FGF2(25 ng/mL), DKK1 (200 ng/mL), and DKK-1 plus FGF2 (25 ng/mL). The rate of cell death in untreated control cultures was considered to be 100 and is indicated by the dotted line. Scale bar in A, B, C = 0.5 mm. *** <span class="html-italic">p</span> &lt; 0.001.</p>
Full article ">
15 pages, 2022 KiB  
Article
Gene Expression Detects the Factors Influencing the Reproductive Success and the Survival Rates of Paracentrotus lividus Offspring
by Serena Federico, Francesca Glaviano, Roberta Esposito, Bruno Pinto, Maissa Gharbi, Anna Di Cosmo, Maria Costantini and Valerio Zupo
Int. J. Mol. Sci. 2022, 23(21), 12790; https://doi.org/10.3390/ijms232112790 - 24 Oct 2022
Cited by 1 | Viewed by 1493
Abstract
The increase in the demand for Paracentrotus lividus roe, a food delicacy, causes increased pressure on its wild stocks. In this scenario, aquaculture facilities will mitigate the effects of anthropogenic pressures on the wild stocks of P. lividus. Consequently, experimental studies should [...] Read more.
The increase in the demand for Paracentrotus lividus roe, a food delicacy, causes increased pressure on its wild stocks. In this scenario, aquaculture facilities will mitigate the effects of anthropogenic pressures on the wild stocks of P. lividus. Consequently, experimental studies should be conducted to enhance techniques to improve efficient aquaculture practices for these animals. Here, we for the first time performed molecular investigations on cultured sea urchins. We aimed at understanding if maternal influences may significantly impact the life of future offspring, and how the culture conditions may impact the development and growth of cultured specimens. Our findings demonstrate that the outcomes of in vitro fertilization of P. lividus are influenced by maternal influences, but these effects are largely determined by culture conditions. In fact, twenty-three genes involved in the response to stress and skeletogenesis, whose expressions were measured by Real Time qPCR, were differently expressed in sea urchins cultured in two experimental conditions, and the results were largely modified in offspring deriving from two groups of females. The findings herein reported will be critical to develop protocols for the larval culture of the most common sea urchin, both for research and industrial production purposes for mass production. Full article
(This article belongs to the Special Issue 21st Anniversary of IJMS: Advances in Molecular Genetics and Genomics)
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<p>Average survival rates of larvae obtained from the two pools of individuals (A+B vs. C+D) in smaller (<b>A</b>) and larger (<b>B</b>) tanks.</p>
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<p>Average larval survival rates evaluated during the first week in (<b>A</b>) smaller and (<b>B</b>) largertanks, referring to the recruits of females A+B vs. those of females C+D. The linear regressions are superimposed.</p>
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<p>(<b>A</b>) Interactomic analysis by STRING (<a href="https://string&#x2013;db.org/" target="_blank">https://string–db.org/</a>; accessed on 30 April 2022). The network graphically displays the relationship between genes. The biological relationships between genes are indicated by different colors. Known interactions: reported by database = light blue and determined experimentally = pink. Expected interactions: gene proximity = green; gene fusion = red; and genes with similar pattern = light blue. (<b>B</b>) <span class="html-italic">Homo sapiens</span> gene names and the corresponding <span class="html-italic">P. lividus</span> orthologous genes. The most significant relations among genes (confidence score cut-off = 900) displaying experimental evidence are highlighted.</p>
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<p>Heatmaps (Heatmapper available at <a href="http://www.heatmapper.ca" target="_blank">www.heatmapper.ca</a>, accessed on 28 July 2022) showing the expression profiles and hierarchical clustering of twenty-three genes analyzed by <span class="html-italic">Real Time qPCR</span> in <span class="html-italic">P. lividus</span> embryos deriving from females A, B, C and D. Color code: red = up-regulated genes with respect to the control; green = down-regulated genes.</p>
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21 pages, 1644 KiB  
Review
The Role of Protein Kinase CK2 in Development and Disease Progression: A Critical Review
by Daniel Halloran, Venu Pandit and Anja Nohe
J. Dev. Biol. 2022, 10(3), 31; https://doi.org/10.3390/jdb10030031 - 27 Jul 2022
Cited by 8 | Viewed by 4449
Abstract
Protein kinase CK2 (CK2) is a ubiquitous holoenzyme involved in a wide array of developmental processes. The involvement of CK2 in events such as neurogenesis, cardiogenesis, skeletogenesis, and spermatogenesis is essential for the viability of almost all organisms, and its role has been [...] Read more.
Protein kinase CK2 (CK2) is a ubiquitous holoenzyme involved in a wide array of developmental processes. The involvement of CK2 in events such as neurogenesis, cardiogenesis, skeletogenesis, and spermatogenesis is essential for the viability of almost all organisms, and its role has been conserved throughout evolution. Further into adulthood, CK2 continues to function as a key regulator of pathways affecting crucial processes such as osteogenesis, adipogenesis, chondrogenesis, neuron differentiation, and the immune response. Due to its vast role in a multitude of pathways, aberrant functioning of this kinase leads to embryonic lethality and numerous diseases and disorders, including cancer and neurological disorders. As a result, CK2 is a popular target for interventions aiming to treat the aforementioned diseases. Specifically, two CK2 inhibitors, namely CX-4945 and CIBG-300, are in the early stages of clinical testing and exhibit promise for treating cancer and other disorders. Further, other researchers around the world are focusing on CK2 to treat bone disorders. This review summarizes the current understanding of CK2 in development, the structure of CK2, the targets and signaling pathways of CK2, the implication of CK2 in disease progression, and the recent therapeutics developed to inhibit the dysregulation of CK2 function in various diseases. Full article
(This article belongs to the Special Issue 2022 Feature Papers by JDB’s Editorial Board Members)
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<p>CK2 is expressed at embryonic day 11 and is critical for many developmental processes. Specifically, CK2 is necessary for limiting neurogenesis and preventing the excessive differentiation of neurons. In addition, CK2 expression promotes long-term memory storage. Further, this protein is essential for skeletogenesis, chondrogenesis, adipogenesis, and proper limb formation. CK2 contributes to spermatogenesis, and the inhibition of its expression leads to infertility. Finally, CK2 is important for B cell differentiation and development, myotube formation, the regulation of the epithelial-to-mesenchymal transition (EMT), and the establishment of circulation between the fetus and the mother during the third trimester.</p>
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<p>There are three classifications of CK2 substrates. Class I substrates are identified as proteins that are equally phosphorylated by the holoenzyme and individually by the catalytic subunits. Class II substrates are specifically phosphorylated by the catalytic submits of CK2 but not by the holoenzyme. Class III substrates are preferentially targeted and phosphorylated by the holoenzyme but not by the catalytic subunits of CK2.</p>
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<p>The localization of CK2 changes when cells are exposed to ionizing radiation (IR). Specifically, CK2α is translocated from the cytoplasm to the nucleus in multiple cell lines, including A549, H460, PC9, and M059K. Further, upon radiation exposure, the overall kinase activity of CK2 increases, emphasizing its role in the DNA-damage-repair response, as it colocalizes with DNA in the nucleus of cells.</p>
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<p>The localization of the catalytic subunits of CK2 is altered during hypoxia. Specifically, these subunits are translocated to the nucleus transiently to aid the cellular response to hypoxia.</p>
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<p>Simplified schematic demonstrating three therapeutics targeting the activity of CK2. (<b>A</b>). CX-4945 is a competitive ATP inhibitor that binds preferentially to ATP-pocket domains. Here, it binds to both CK2α and CK2α’, thereby preventing ATP from binding and activating CK2. CX-4945 has prevented tumor progression by inhibiting signaling pathways such as Smad and PI3K/AKT. (<b>B</b>). CIBG-300 functions by binding to conserved phosphorylation sequences on the substrates of CK2, such as Akt. This association prevents the phosphorylation of these substrates by CK2, leading to decreased cell signaling, proliferation, and survival. (<b>C</b>). CK2.3 is uptaken by cells and binds to CK2, therefore preventing its association with BMPRIa. As a result, BMPRII can phosphorylate BMPRIa, leading to the downstream activation of signaling pathways that induce mineralization and cell survival.</p>
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13 pages, 4281 KiB  
Article
Toxicity of Vanadium during Development of Sea Urchin Embryos: Bioaccumulation, Calcium Depletion, ERK Modulation and Cell-Selective Apoptosis
by Roberto Chiarelli, Rosaria Scudiero, Valeria Memoli, Maria Carmela Roccheri and Chiara Martino
Int. J. Mol. Sci. 2022, 23(11), 6239; https://doi.org/10.3390/ijms23116239 - 2 Jun 2022
Cited by 6 | Viewed by 1859
Abstract
Vanadium toxicology is a topic of considerable importance as this metal is widely used in industrial and biomedical fields. However, it represents a potential emerging environmental pollutant because wastewater treatment plants do not adequately remove metal compounds that are subsequently released into the [...] Read more.
Vanadium toxicology is a topic of considerable importance as this metal is widely used in industrial and biomedical fields. However, it represents a potential emerging environmental pollutant because wastewater treatment plants do not adequately remove metal compounds that are subsequently released into the environment. Vanadium applications are limited due to its toxicity, so it is urgent to define this aspect. This metal is associated with sea urchin embryo toxicity as it perturbs embryogenesis and skeletogenesis, triggering several stress responses. Here we investigated its bioaccumulation and the correlation with cellular and molecular developmental pathways. We used cytotoxic concentrations of 1 mM and 500 μM to perform quantitative analyses, showing that vanadium accumulation interferes with calcium uptake during sea urchin development and provokes a disruption in the biomineralization process. At the end of the whole treatment, the accumulation of vanadium was about 14 and 8 μg for embryos treated respectively with 1 mM and 500 μM, showing a dose-dependent response. Then, we monitored the cell signaling perturbation, analyzing key molecular markers of cell survival/cell death mechanisms and the DNA fragmentation associated with apoptosis. This paper clarifies vanadium’s trend to accumulate directly into embryonic cells, interfering with calcium uptake. In addition, our results indicate that vanadium can modulate the ERK pathway and activate a cell-selective apoptosis. These results endorse the sea urchin embryo as an adequate experimental model to study metal-related cellular/molecular responses. Full article
(This article belongs to the Special Issue Molecular Research on Reproductive Toxicity)
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<p>Pictures of representative embryos at 36 h of development/treatment. Control embryo (<b>A</b>), 1 mM V-treated embryo (<b>B</b>), 500 μM V-treated embryo (<b>C</b>). Bar: 100 μm.</p>
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<p>Amount of V and Ca incorporated during the time after 12, 18, 24, 30, 36 and 42 h of development/treatment. Embryos were cultured in 1 mM or 500 μM of V. V (<b>A</b>) and Ca (<b>B</b>) content were detected by Inductively Coupled Plasma Mass Spectrometry (ICP-MS), determining the metal quantity in about 250,000 embryos. Experiments were performed in triplicate and data are expressed as means ± standard deviation (<span class="html-italic">n</span> = 3 ± SD).</p>
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<p>Immunoblotting detection and quantitative analysis for pERK and ERK 1/2. (<b>A</b>) Total lysates of control and V-treated (1 mM, 500 μM) embryos after 24, 30, 36 and 42 h of development/treatment. Actin was used as a loading control. Histograms show the densitometric analysis of bands identified for (<b>B</b>) pERK and (<b>C</b>) ERK 1/2. Relative protein expression, reported as arbitrary units, was calculated as the band density ratio to that of actin. Experiments were performed in triplicate and data are expressed as means ± standard deviation (<span class="html-italic">n</span> = 3 ± SD). Data were analyzed by one-way ANOVA. Treatments with the same lowercase letter do not differ (Tukey HSD).</p>
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<p>Immunoblotting detection and quantitative analysis for CHOP 10/GADD 153 and cleaved caspase 7. (<b>A</b>) Total lysates of control and V-treated (1 mM, 500 μM) embryos after 24, 30, 36 and 42 h of development/treatment. Actin was used as a loading control. Histograms show the densitometric analysis of bands identified for (<b>B</b>) CHOP-10/GADD 153 and (<b>C</b>) cleaved caspase 7. Relative protein expression, reported as arbitrary units, was calculated as the band density ratio to that of actin. Experiments were performed in triplicate and data are expressed as means ± standard deviation (<span class="html-italic">n</span> = 3 ± SD). Data were analyzed by one-way ANOVA. Treatments with the same lowercase letter do not differ (Tukey HSD).</p>
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<p>Fluorescent TUNEL assay and densitometric analysis. Pictures of demonstrative embryos at 24, 30, 36 and 42 h of development/treatment. DNA fragmentation (<b>A1</b>–<b>N1</b>). Nuclei marked with propidium iodide (<b>A2</b>–<b>N2</b>). Merge of both signals (<b>A3</b>–<b>N3</b>). Control embryos (<b>A1</b>–<b>A3</b>,<b>D1</b>–<b>D3</b>,<b>H1</b>–<b>H3</b>,<b>K1</b>–<b>K3</b>); 1 mM V-treated embryos (<b>B1</b>–<b>B3</b>,<b>E1</b>–<b>E3</b>,<b>I1</b>–<b>I3</b>,<b>L1</b>–<b>L3</b>); 500 μM V-treated embryos (<b>C1</b>–<b>C3</b>,<b>F1</b>–<b>F3</b>,<b>J1</b>–<b>J3</b>,<b>M1</b>–<b>M3</b>). Positive control embryo at 42 h of development (<b>G1</b>–<b>G3</b>). Negative control embryo at 42 h of development (<b>N1</b>–<b>N3</b>). Bar = 100 μm. Histograms showing data related to the quantitative analysis of fluorescence from apoptotic DNA. Experiments were performed in triplicate and data are expressed as means ± standard deviation (<span class="html-italic">n</span> = 3 ± SD). Data were analyzed by one-way ANOVA. Treatments with the same lowercase letter do not differ (Tukey HSD).</p>
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12 pages, 739 KiB  
Review
Computation of Fetal Kicking in Various Fetal Health Examinations: A Systematic Review
by Yuwei Liu, Rongrong Xuan, Yuhuan He, Feng Ren and Yaodong Gu
Int. J. Environ. Res. Public Health 2022, 19(7), 4366; https://doi.org/10.3390/ijerph19074366 - 5 Apr 2022
Cited by 8 | Viewed by 2489
Abstract
Fetal movement has always been considered an essential indicator to evaluate the health of the unborn fetus. Many factors affect fetal movement. The frequency of fetal kicking is an important measurement of whether fetal development is progressing and healthy. Various instruments and methods [...] Read more.
Fetal movement has always been considered an essential indicator to evaluate the health of the unborn fetus. Many factors affect fetal movement. The frequency of fetal kicking is an important measurement of whether fetal development is progressing and healthy. Various instruments and methods of detecting fetal movement have been used and each method has its advantages and disadvantages. Although limited by the fetal environment in utero, the finite element method and musculoskeletal model can be used to calculate fetal lower limb movement. This review aims to summarize the current detection techniques for fetal movement, especially in the lower limbs. These will be outlined by describing the different measurements of fetal movement, and the related biomechanical analyses of fetal lower limb skeletogenesis and the associated muscular development to better evaluate and calculate the movements of the fetus in the womb. Full article
(This article belongs to the Special Issue Physical Activity for Public Health)
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<p>PRISMA flowchart of the process including the literature search and screening.</p>
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23 pages, 5533 KiB  
Review
The Evolution of Biomineralization through the Co-Option of Organic Scaffold Forming Networks
by Smadar Ben-Tabou de-Leon
Cells 2022, 11(4), 595; https://doi.org/10.3390/cells11040595 - 9 Feb 2022
Cited by 13 | Viewed by 3442
Abstract
Biomineralization is the process in which organisms use minerals to generate hard structures like teeth, skeletons and shells. Biomineralization is proposed to have evolved independently in different phyla through the co-option of pre-existing developmental programs. Comparing the gene regulatory networks (GRNs) that drive [...] Read more.
Biomineralization is the process in which organisms use minerals to generate hard structures like teeth, skeletons and shells. Biomineralization is proposed to have evolved independently in different phyla through the co-option of pre-existing developmental programs. Comparing the gene regulatory networks (GRNs) that drive biomineralization in different species could illuminate the molecular evolution of biomineralization. Skeletogenesis in the sea urchin embryo was extensively studied and the underlying GRN shows high conservation within echinoderms, larval and adult skeletogenesis. The organic scaffold in which the calcite skeletal elements form in echinoderms is a tubular compartment generated by the syncytial skeletogenic cells. This is strictly different than the organic cartilaginous scaffold that vertebrates mineralize with hydroxyapatite to make their bones. Here I compare the GRNs that drive biomineralization and tubulogenesis in echinoderms and in vertebrates. The GRN that drives skeletogenesis in the sea urchin embryo shows little similarity to the GRN that drives bone formation and high resemblance to the GRN that drives vertebrates’ vascular tubulogenesis. On the other hand, vertebrates’ bone-GRNs show high similarity to the GRNs that operate in the cells that generate the cartilage-like tissues of basal chordate and invertebrates that do not produce mineralized tissue. These comparisons suggest that biomineralization in deuterostomes evolved through the phylum specific co-option of GRNs that control distinct organic scaffolds to mineralization. Full article
(This article belongs to the Section Cell and Gene Therapy)
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<p>Partial Eumetazoan phylogenetic tree containing phyla and species discussed in this paper and their biomineralization programs. (<b>A</b>), calcification in cnidarian and protostome phyla. Numbers in red is the estimated time of divergence in million years (MA) based on [<a href="#B27-cells-11-00595" class="html-bibr">27</a>,<a href="#B29-cells-11-00595" class="html-bibr">29</a>]. Light blue indicates the usage of different CaCO3 polymorphs including but not limited to calcite, aragonite or vaterite. Dark blue indicates the usage of calcite, and purple indicates the usage of apatite. Black indicates non-mineralizing species. (<b>B</b>), skeleton formation in deuterostomes. Color code as in (<b>A</b>), with the addition of green, indicating the usage of cartilage by the skeletogenic tissue. Squares indicate the evolutionary origin of a specific skeletal tissue.</p>
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<p>Possible models for the evolution of biomineralization GRNs from ancestral GRNs. (<b>A</b>), a model where an ancestral GRN that drove the formation of an organic scaffold was co-opted for biomineralization by activating regulatory and differentiation genes that assist in calcification. In this model we expect to see a strong similarity between the biomineralization of GRN to the GRN that drive the formation of the organic scaffold in non-biomineralization species. (<b>B</b>), a model where an ancestral GRN that drove the activation of genes that participate in calcification was co-opted for biomineralization through the activation of phylum specific regulatory and differentiation genes that control organic scaffold formation. In that case, we expect to see a similarity between the regulatory and differentiation genes that drive calcification in the two phyla.</p>
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<p>Sea urchin skeletogenic GRN and the evolution of echinoderm skeletogenic GRN. (<b>A</b>), a scheme showing larval skeleton formation in the sea urchin embryo. The skeletogenic cells (red) form a ring with two lateral skeletogenic clusters where the spicule form. Enlargement, showing the mineral (gray) concentrated in vesicles and transported to the spicule tubular compartment where it is engulfed within a thin layer of extracellular matrix. (<b>B</b>), sea urchin larval skeletogenic GRN and differentiation genes with various functions. (<b>C</b>–<b>E</b>), embryonic territories in the different echinoderm clades. Color codes are explained in the figure and in the text. C, embryonic territories in the sea urchin (SU) and the pencil sea urchin (PU). D, embryonic territories in the sea cucumber (SC) and brittle star (BS). (<b>E</b>), embryonic territories in the sea star (SS). (<b>F</b>), expression of the skeletogenic regulatory genes in the mesoderm of embryos of different echinoderm clades. Color code explained in figure. (<b>G</b>), expression of skeletogenic regulatory genes in adult skeletogenic cells in three echinoderm clades.</p>
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<p>Bone biomineralization and vascular tubulogenesis GRNs in vertebrates. (<b>A</b>), schematic diagram showing a cross section of the dorsal part of a vertebrate embryo and the different embryonic territories that contribute to skeletal and vascular tissues. (<b>B</b>), tissues and cell types that participate in endochondral ossification in vertebrates. Immature cartilage (green) is generated by proliferating and resting chondrocytes. Mature cartilage (blue) is generated by hypertrophic chondrocytes, bone matrix and mineralization (gray) are generated by osteoblasts, maintained by osteocytes and reabsorbed by osteoclasts (purple). (<b>C</b>), GRN and differentiation genes in the different bone forming cells. Color code matches the territories and cells in (<b>B</b>,<b>D</b>), schematic diagram of vertebrate blood vessel showing the different cell types that constitute it. Blood cells occupy the lumen which is engulfed by endothelial cells. The endothelial cells are bound to the basement membrane from the inner side of the vessel and pericytes from the outside. Image courtesy of Yarden Ben-Tabou de-Leon (artist). (<b>E</b>), endothelial cell GRN and differentiation genes.</p>
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<p>Proposed models of the evolution of the biomineralization GRNs in echinoderms and in vertebrates. (<b>A</b>), an ancestral vascular GRN that generated blood vessels where hemocytes flow evolved in vertebrates to make the endothelial GRN and was co-opted for biomineralization in echinoderms. The ancestral vascular GRN included the VEGF pathway and transcription factors of the ETS family. Co-option for biomineralization included the activation of the transcription factor Alx1, the evolution of novel echinoderm spicule matrix (SM) proteins and the activation of genes of the biomineralization toolkit. The vertebrate endothelial GRN and the echinoderm skeletogenic GRN show similarities and trans-differentiation potential to the hemocyte GRN. (<b>B</b>), an ancestral GRN that drove cartilage formation was co-opted for biomineralization in the vertebrate phylum. The ancestral cartilage GRN included the transcription factors SoxE and SoxD that drove the expression of Col2. The co-option for biomineralization was through the activation of the transcription factors Runx and Sp7, the evolution of novel bone matrix (BM) proteins and the activation of Col1 and genes of the biomineralization toolkit.</p>
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25 pages, 6729 KiB  
Review
Regulation of FGF-2, FGF-18 and Transcription Factor Activity by Perlecan in the Maturational Development of Transitional Rudiment and Growth Plate Cartilages and in the Maintenance of Permanent Cartilage Homeostasis
by Anthony J. Hayes, John Whitelock and James Melrose
Int. J. Mol. Sci. 2022, 23(4), 1934; https://doi.org/10.3390/ijms23041934 - 9 Feb 2022
Cited by 16 | Viewed by 3292
Abstract
The aim of this study was to highlight the roles of perlecan in the regulation of the development of the rudiment developmental cartilages and growth plate cartilages, and also to show how perlecan maintains permanent articular cartilage homeostasis. Cartilage rudiments are transient developmental [...] Read more.
The aim of this study was to highlight the roles of perlecan in the regulation of the development of the rudiment developmental cartilages and growth plate cartilages, and also to show how perlecan maintains permanent articular cartilage homeostasis. Cartilage rudiments are transient developmental templates containing chondroprogenitor cells that undergo proliferation, matrix deposition, and hypertrophic differentiation. Growth plate cartilage also undergoes similar changes leading to endochondral bone formation, whereas permanent cartilage is maintained as an articular structure and does not undergo maturational changes. Pericellular and extracellular perlecan-HS chains interact with growth factors, morphogens, structural matrix glycoproteins, proteases, and inhibitors to promote matrix stabilization and cellular proliferation, ECM remodelling, and tissue expansion. Perlecan has mechanotransductive roles in cartilage that modulate chondrocyte responses in weight-bearing environments. Nuclear perlecan may modulate chromatin structure and transcription factor access to DNA and gene regulation. Snail-1, a mesenchymal marker and transcription factor, signals through FGFR-3 to promote chondrogenesis and maintain Acan and type II collagen levels in articular cartilage, but prevents further tissue expansion. Pre-hypertrophic growth plate chondrocytes also express high Snail-1 levels, leading to cessation of Acan and CoI2A1 synthesis and appearance of type X collagen. Perlecan differentially regulates FGF-2 and FGF-18 to maintain articular cartilage homeostasis, rudiment and growth plate cartilage growth, and maturational changes including mineralization, contributing to skeletal growth. Full article
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<p>Localisation of perlecan in human foetal knee joints (12 weeks gestational age). Immunolocalisation of HSPG2 with perlecan domain-1 MAb A76 in a 12-week-old gestational age human foetal knee demonstrating perlecan as a major extracellular matrix proteoglycan of the tibial and femoral cartilaginous rudiments (<b>a</b>) and menisci (M). Perlecan is also prominently localised around the margins of small chondroprogenitor cell niches in the stromal tissue surrounding the rudiment (<b>b</b>). Nomarski differential interference contrast images demonstrate the differing stromal, surface, and central cartilaginous rudiment cell morphologies (<b>c</b>). Detail of a chondroprogenitor niche at the interface of the stromal and rudiment surface with perlecan prominently located around the niche (<b>d</b>). Chromogen NovaRED, nuclei stained with haematoxylin. Photosegments modified from [<a href="#B16-ijms-23-01934" class="html-bibr">16</a>] with permission © Melrose 2016.</p>
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<p>Perlecan localisation in ovine knee and hip joints. Immunolocalisation of perlecan in cartilaginous tissues of a two-year-old ovine knee femoral condyle (<b>a</b>) and tibial plateau (<b>b</b>), patella (<b>c</b>) and in the humeral head of a hip joint (<b>d</b>). Higher power images demonstrate the pericellular localisation of perlecan (small arrows) around chondrocytes in regions of the femoral (<b>e</b>) and tibial articular cartilages (<b>f</b>) (boxed areas in (<b>a</b>,<b>b</b>)). Perlecan is also present as a gradient throughout the femoral long bone growth plate ECM of the hip in the resting and proliferative zones (double headed arrow) and is prominently expressed pericellularly by the hypertrophic columnar hip chondrocytes located in the bottom of photosegment (<b>g</b>). NovaRED chromogen, perlecan localised with MAb A7L6 to perlecan domain IV. Photo segments (<b>a</b>–<b>g</b>) modified from [<a href="#B9-ijms-23-01934" class="html-bibr">9</a>] reproduced under Open Access Creative Commons Attribution 4.0 International licence images © the authors (2010).</p>
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<p>Schematic depiction of the domain organization of a hypothetical HS chain showing the FGF-2 and FGFR binding domain. Figure modified from [<a href="#B46-ijms-23-01934" class="html-bibr">46</a>,<a href="#B47-ijms-23-01934" class="html-bibr">47</a>].</p>
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<p>Immunolocalisation of FGF-18 and type X collagen in growth plate cartilage. Upregulation of FGF-18 immunolocalised in hypertrophic growth plate chondrocytes (<b>a</b>), with their enlarged morphologies clearly visualized by alcian blue staining (<b>b</b>) and in a Masson’s trichrome image viewed under Nomarski differential interference contrast optics (<b>c</b>). Gene profiling shows that FGF-18 initially stimulates chondrogenesis in bone marrow chondroprogenitor cells up to day 30 in micromass pellet culture but by day 31, type II collagen expression ceases and osteogenic differentiation (Mef2c) is initiated (<b>d</b>). FGF-2, however, maintains chondrogenesis throughout the full period of pellet culture (up to day 41). Type X collagen synthesis occurs in FGF-18-stimulated cultures from day 31 and is clearly evident at the chondro-osseous junction (<b>e</b>). FGF-18 also upregulates <span class="html-italic">Snail1</span> expression in the hypertrophic growth plate chondrocytes. Images (<b>a</b>–<b>d</b>) ©Melrose 2016, reproduced from [<a href="#B50-ijms-23-01934" class="html-bibr">50</a>] with permission. Images Image (<b>e</b>) supplied courtesy of DrYao Hao, © Yao Hao 2019, Institute of Genetic Medicine, International Centre for Life, Newcastle University, UK.</p>
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<p>Immunolocalisation of perlecan in ovine AF and NP chondrons using laser scanning confocal microscopy and 3D rendered image stacks. Nuclear DAPI counterstaining is shown in blue. Fluorescent perlecan immunolocalisations were undertaken as described in [<a href="#B140-ijms-23-01934" class="html-bibr">140</a>]. A string of outer AF cells (<b>a</b>) and 3D reconstructions of perlecan in an NP chondron with 3D volume indicated by white boundary box (<b>b</b>). Immunolocalisation of perlecan in a stacked confocal image of an NP chondron (<b>c</b>) and in a 0.5 µm single z-stack image depicting punctate nuclear perlecan deposits (<b>d</b>). These deposits are obscured in the stacked image (<b>c</b>) by overlying tissue. Key: 1. Pericellular matrix, 2. Type VI Collagenous capsule, 3. Nuclear deposits of perlecan, 4. Vesicular perlecan transported out of the cell into the chondron matrix (*). n = nucleus. Figure reproduced under Open Access CC-BY-SA licence from [<a href="#B137-ijms-23-01934" class="html-bibr">137</a>] © the authors 2020.</p>
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20 pages, 5493 KiB  
Article
Phylogenetic Diversity of Ossification Patterns in the Avian Vertebral Column: A Review and New Data from the Domestic Pigeon and Two Species of Grebes
by Tomasz Skawiński, Piotr Kuziak, Janusz Kloskowski and Bartosz Borczyk
Biology 2022, 11(2), 180; https://doi.org/10.3390/biology11020180 - 24 Jan 2022
Viewed by 3963
Abstract
Despite many decades of studies, our knowledge of skeletal development in birds is limited in many aspects. One of them is the development of the vertebral column. For many years it was widely believed that the column ossifies anteroposteriorly. However, later studies indicated [...] Read more.
Despite many decades of studies, our knowledge of skeletal development in birds is limited in many aspects. One of them is the development of the vertebral column. For many years it was widely believed that the column ossifies anteroposteriorly. However, later studies indicated that such a pattern is not universal in birds and in many groups the ossification starts in the thoracic rather than cervical region. Recent analyses suggest that two loci, located in the cervical and thoracic vertebrae, were ancestrally present in birds. However, the data on skeletal development are very scarce in the Neoaves, a clade that includes approximately 95% of extant species. We review the available information about the vertebral column development in birds and describe the ossification pattern in three neoavians, the domestic pigeon (Columba livia domestica), the great crested grebe (Podiceps cristatus) and the red-necked grebe (Podiceps grisegena). In P. cristatus, the vertebral column starts ossifying in the thoracic region. The second locus is present in the cervical vertebrae. In the pigeon, the cervical vertebrae ossify before the thoracics, but both the thoracic and cervical loci are present. Our ancestral state reconstructions confirm that both these loci were ancestrally present in birds, but the thoracic locus was later lost in psittacopasserans and at least some galloanserans. Full article
(This article belongs to the Section Developmental Biology)
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<p>Phylogenetic relationships of the species discussed in the text. The topology of the dendrogram follows the ‘consensus phylogeny of birds’ from Braun and Kimball [<a href="#B27-biology-11-00180" class="html-bibr">27</a>], while the interrelationships of the main clades follow primarily Prum et al. [<a href="#B28-biology-11-00180" class="html-bibr">28</a>] and Kuhl et al. [<a href="#B29-biology-11-00180" class="html-bibr">29</a>]. Codes: 1—Neornithes, 2—Palaeognathae, 3—Neognathae, 4—Galloanserae, 5—Galliformes, 6—Anseriformes, 7—Neoaves, 8—Charadriiformes, 9—Psittacopasserae.</p>
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<p>A schematic summary of the ossification patterns of the vertebral bodies in birds (<b>A</b>,<b>B</b>). Blue indicates cervical loci while orange indicates thoracic loci. The bird silhouette was taken from PhyloPic and is available in public domain.</p>
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<p>A schematic summary of the developmental patterns of the vertebral arches in birds (<b>A</b>–<b>D</b>). Ossification (appearance) of the vertebral arches. Blue indicates cervical loci, orange—thoracic loci, red—synsacral loci, purple—caudal loci.</p>
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<p>A schematic summary of developmental patterns in the vertebral arch fusion (<b>A</b>–<b>C</b>). Blue indicates cervical loci, orange—thoracic loci.</p>
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<p>Ossification pattern of the vertebral column in <span class="html-italic">Podiceps cristatus</span>. (<b>A</b>) A schematic drawing of a single vertebra, showing the body and the vertebral arch. (<b>B</b>–<b>I</b>) Ossifications present in <span class="html-italic">P. cristatus</span> specimens, from state 1 (<b>A</b>) to state 8 (<b>I</b>). Grey colour indicates the presence of bone, while white indicates cartilage.</p>
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<p>Ossification of the vertebral column in a state 3 specimen of <span class="html-italic">Podiceps cristatus</span>. The thoracic locus (with the ossification spreading into cervical and lumbosacral (synsacral) vertebrae) is seen on the photograph. Scale bar = 5 mm. tv—thoracic vertebrae.</p>
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<p>Ossification of the vertebral column in a state 3 specimen of <span class="html-italic">Podiceps cristatus</span>. The cervical locus is seen on the photograph. Scale bar = 5 mm. cra—cranium, cv—cervical vertebrae.</p>
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<p>The neck and thorax of a neonatal <span class="html-italic">Podiceps grisegena</span>. The vertebral arches are fused in cervical vertebrae but still separate in the thoracic vertebrae. Scale bar = 5 mm. cra—cranium, cv—cervical vertebrae, fu—furcula.</p>
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<p>Ossification pattern of the vertebral column in <span class="html-italic">Columba livia</span>. (<b>A</b>–<b>C</b>) Ossifications present in studied specimens, from specimens at the time of hatching (<b>A</b>) to 3 days old (<b>C</b>). Dark grey indicates the presence of bone, while white indicates cartilage. The vertebrae that are missing in the specimen are highlighted by light grey. See <a href="#biology-11-00180-f005" class="html-fig">Figure 5</a>A for the legend.</p>
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<p>The state of ossification in a hatchling <span class="html-italic">Columba livia domestica</span>. (<b>A</b>) Ventral view. (<b>B</b>) Dorsal view. Scale bar = 1 cm. cv—cervical vertebrae.</p>
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<p>The state of ossification in a 1-day-old <span class="html-italic">Columba livia domestica</span> in lateral view. Scale bar = 1 cm. va—vertebral arches, vb—vertebral bodies.</p>
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<p>The state of ossification in a 2-day-old <span class="html-italic">Columba livia domestica</span>. (<b>A</b>) Lateral view. (<b>B</b>) Dorsal view. Scale bar = 1 cm. cv—cervical vertebrae, tv—thoracic vertebrae, lsv—lumbosacral (synsacral) vertebrae.</p>
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<p>Parsimony-based ancestral state reconstructions of the ossification patterns of the vertebral body ossification. Black indicates the presence of a thoracic locus, while white indicates the lack thereof.</p>
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<p>Maximum likelihood-based ancestral state reconstructions of the ossification patterns of the vertebral body ossification. Black indicates the presence of a thoracic locus, while white indicates the lack thereof. (<b>A</b>) Reconstructions based on the tree with equal branch length (=1). (<b>B</b>) Reconstructions based on the temporally calibrated tree. See <a href="#app1-biology-11-00180" class="html-app">Table S2</a> for the exact proportional likelihood values.</p>
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13 pages, 2473 KiB  
Review
Retinoid Agonists in the Targeting of Heterotopic Ossification
by Robert J. Pignolo and Maurizio Pacifici
Cells 2021, 10(11), 3245; https://doi.org/10.3390/cells10113245 - 19 Nov 2021
Cited by 9 | Viewed by 3781
Abstract
Retinoids are metabolic derivatives of vitamin A and regulate the function of many tissues and organs both prenatally and postnatally. Active retinoids, such as all trans-retinoic acid, are produced in the cytoplasm and then interact with nuclear retinoic acid receptors (RARs) to [...] Read more.
Retinoids are metabolic derivatives of vitamin A and regulate the function of many tissues and organs both prenatally and postnatally. Active retinoids, such as all trans-retinoic acid, are produced in the cytoplasm and then interact with nuclear retinoic acid receptors (RARs) to up-regulate the transcription of target genes. The RARs can also interact with target gene response elements in the absence of retinoids and exert a transcriptional repression function. Studies from several labs, including ours, showed that chondrogenic cell differentiation and cartilage maturation require (i) the absence of retinoid signaling and (ii) the repression function by unliganded RARs. These and related insights led to the proposition that synthetic retinoid agonists could thus represent pharmacological agents to inhibit heterotopic ossification (HO), a process that recapitulates developmental skeletogenesis and involves chondrogenesis, cartilage maturation, and endochondral ossification. One form of HO is acquired and is caused by injury, and another severe and often fatal form of it is genetic and occurs in patients with fibrodysplasia ossificans progressiva (FOP). Mouse models of FOP bearing mutant ACVR1R206H, characteristic of most FOP patients, were used to test the ability of the retinoid agonists selective for RARα and RARγ against spontaneous and injury-induced HO. The RARγ agonists were found to be most effective, and one such compound, palovarotene, was selected for testing in FOP patients. The safety and effectiveness data from recent and ongoing phase II and phase III clinical trials support the notion that palovarotene may represent a disease-modifying treatment for patients with FOP. The post hoc analyses showed substantial efficacy but also revealed side effects and complications, including premature growth plate closure in some patients. Skeletally immature patients will need to be carefully weighed in any future regulatory indications of palovarotene as an important therapeutic option in FOP. Full article
(This article belongs to the Special Issue Retinoic Acid and Retinoid X Receptors)
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<p>Schematic depicting the major developmental steps during endochondral bone formation. Following progenitor cell commitment and condensation (left panels), chondrogenic differentiation, cartilage maturation, hypertrophy, and vascular invasion require the concerted and stage-dependent up-regulation of indicated pathways and transcription factors (middle panels, green arrows). This leads to replacement of hypertrophic cartilage with bone (upright panel). Importantly, chondrogenesis and cartilage maturation require down-regulation of antagonistic pathways, such as <b>retinoid</b> and Wnt signaling (red arrow), as well as action by unliganded RNA receptors exerting transcriptional repression (<b>ul-RAR</b>). TGFβ, transforming growth factor β; BMP, bone morphogenetic protein; IHH, Indian hedgehog; VEGF, vascular endothelial growth factor; Osx, osterix.</p>
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<p>Translation of retinoid signaling roles in chondrogenic differentiation into clinical studies on inhibition of endochondral heterotopic ossification (HO). Retinoids, especially RARγ agonists, have inhibitory effects on chondrogenesis in micromass cultures (left); preclinical mouse models of HO (middle); and HO formation in fibrodysplasia ossificans progressiva (right).</p>
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13 pages, 2458 KiB  
Article
Deubiquitinating Enzyme USP8 Is Essential for Skeletogenesis by Regulating Wnt Signaling
by Sachin Chaugule, Jung-Min Kim, Yeon-Suk Yang, Klaus-Peter Knobeloch, Xi He and Jae-Hyuck Shim
Int. J. Mol. Sci. 2021, 22(19), 10289; https://doi.org/10.3390/ijms221910289 - 24 Sep 2021
Cited by 5 | Viewed by 2654
Abstract
Disturbance in a differentiation program of skeletal stem cells leads to indecorous skeletogenesis. Growing evidence suggests that a fine-tuning of ubiquitin-mediated protein degradation is crucial for skeletal stem cells to maintain their stemness and osteogenic potential. Here, we demonstrate that the deubiquitinating enzyme [...] Read more.
Disturbance in a differentiation program of skeletal stem cells leads to indecorous skeletogenesis. Growing evidence suggests that a fine-tuning of ubiquitin-mediated protein degradation is crucial for skeletal stem cells to maintain their stemness and osteogenic potential. Here, we demonstrate that the deubiquitinating enzyme (DUB) ubiquitin-specific protease 8 (USP8) stabilizes the Wnt receptor frizzled 5 (FZD5) by preventing its lysosomal degradation. This pathway is essential for Wnt/β-catenin signaling and the differentiation of osteoprogenitors to mature osteoblasts. Accordingly, deletion of USP8 in osteoprogenitors (Usp8Osx) resulted in a near-complete blockade in skeletal mineralization, similar to that seen in mice with defective Wnt/β-catenin signaling. Likewise, transplanting USP8-deficient osteoprogenitors under the renal capsule in wild-type secondary hosts did not to induce bone formation. Collectively, this study unveils an essential role for the DUB USP8 in Wnt/β-catenin signaling in osteoprogenitors and osteogenesis during skeletal development. Full article
(This article belongs to the Section Molecular Biophysics)
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<p>USP8 controls Wnt/β–catenin signaling in osteoblasts. (<b>A</b>) Immunohistochemistry (IHC) for USP8 in mouse femurs at postnatal day 0 (<span class="html-italic">n</span> = 3/group). Left, IgG control; right, IHC for USP8 in the cortical bone. CB, cortical bone; PO, periosteum. Arrows indicate bone-residing osteoblasts and osteocytes. Scale bar, 200 μm. (<b>B</b>,<b>C</b>) Kinetics of USP8 expression during osteoblast differentiation. Skeletal stem cells (SSCs, CD45<sup>−</sup>Ter119<sup>−</sup>Tie2<sup>−</sup>αVInt<sup>+</sup>Thy1<sup>−</sup>CD105<sup>−</sup>CD200<sup>+</sup>) isolated from E18.5 wild-type embryonic limbs were cultured under osteogenic condition, and USP8 expression was assessed by immunoblotting analysis. Protein levels of USP8 were displayed by the ratio of USP8 to GAPDH loading control (<span class="html-italic">n</span> = 4). (<b>D</b>) Primary osteoblast precursors (COBs) isolated from the calvaria of P0 <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> neonates (<span class="html-italic">n</span> = 5/group) were cultured under undifferentiated or osteogenic conditions for 6 days, and lysates were immunoblotted with the indicated antibodies. GAPDH was used as a loading control. (<b>E</b>) Immunohistochemistry for β–catenin in the primary ossification center of E18.5 <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> femurs (<span class="html-italic">n</span> = 3/group). Scale bar, 100 µm. Data are shown as a box-and-whisker plot (with median and interquartile ranges) from min to max, with all data points shown. Statistical analysis was performed using an unpaired Student’s t–test within Prism.</p>
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<p>USP8 controls Wnt/FZD5/β–catenin signaling in osteoblasts. (<b>A</b>) Wild–type COBs were lysed, immunoprecipitated with anti–IgG control or anti–FZD5 antibody and protein G-conjugated agarose beads and immunoblotted with the indicated antibodies. WCL; whole cell lysate. (<b>B</b>) <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> COBs (<span class="html-italic">n</span> = 5/group) were cultured under undifferentiated or osteogenic conditions for 6 days, and lysates were immunoblotted with the indicated antibodies. GAPDH was used as a loading control. (<b>C</b>) <span class="html-italic">Usp8<sup>fl/fl</sup></span> COBs were transduced with lentivirus expressing vector (WT) or CRE recombinase (ΔUSP8). Two days after puromycin–selection, cells were treated with vehicle or Bafilomycin A1 (Baf A1, 10 nM) overnight and immunoblotted with indicated antibodies. (<b>D</b>) HA–FZD5–expressing WT and ΔUSP8 COBs were treated with 10 nM of Baf A1 overnight, immunoprecipitated with anti–HA conjugated agarose, and immunoblotted with anti–ubiquitin antibody. (<b>E</b>) WT or ΔUSP8 COBs were transfected with a TopFlash luciferase reporter gene along with vector, USP8C, USP8N, or FZD5. Cells were cultured under osteogenic condition for 3 days, and β–catenin activity was assessed by measuring luciferase activity. (<b>F</b>,<b>G</b>) WT or ΔUSP8 COBs expressing vector, USP8C, USP8N, or FZD5 were immunoblotted with indicated antibodies. Protein levels of β–catenin were displayed by the ratio of β–catenin to GAPDH loading control (<span class="html-italic">n</span> = 3). Data are shown as a box-and-whisker plot (with median and interquartile ranges) from min to max, with all data points shown. Statistical analysis was performed using an unpaired Student’s t–test within Prism.</p>
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<p><span class="html-italic">Usp8<sup>Osx</sup></span> mice display impaired ossification during skeletogenesis. (<b>A</b>) Alizarin red/alcian blue staining of skeletal preparations of whole body, forelimbs, hindlimbs, skull, and clavicle of E18.5 <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> embryos (<span class="html-italic">n</span> = 3/group). Scale bar, 1 mm. (<b>B</b>,<b>C</b>) Alizarin red/alcian blue staining of skeletal preparations of hindlimbs of P0 neonates (B, <span class="html-italic">n</span> = 3/group) and E.15.5 embryos (C, <span class="html-italic">n</span> = 3/group). Scale bar, 1 mm. (<b>D</b>) Safranin O–stained longitudinal sections of P0 <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> femurs (<span class="html-italic">n</span> = 3/group). Scale bar, 250 μm (left) and 100 μm (right, enlarged insert image). (<b>E</b>) Immunohistochemistry (IHC) for Bglap in the trabecular bone of P0 <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> femurs (<span class="html-italic">n</span> = 3/group). Upper: IgG control, middle and down: IHC for Bglap. Scale bar, 100 μm.</p>
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<p>USP8 is essential for osteogenic differentiation of skeletal progenitors. (<b>A</b>) Quantification of flow cytometry on subpopulations of skeletal progenitors isolated from E18.5 <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> embryonic limbs (<span class="html-italic">n</span> = 3/group). SSC (CD45<sup>−</sup>Ter119<sup>−</sup>Tie2<sup>−</sup>αVInt<sup>+</sup>Thy<sup>−</sup>CD105<sup>−</sup>CD200<sup>+</sup>); pre-BCSP (CD45<sup>−</sup>Ter119<sup>−</sup>Tie2<sup>−</sup>αVInt<sup>+</sup>Thy<sup>−</sup>CD105<sup>−</sup>CD200<sup>−</sup>); BCSP (CD45<sup>−</sup>Ter<sup>−</sup>119<sup>−</sup>Tie2<sup>−</sup>αVInt<sup>+</sup>Thy<sup>+</sup>CD105<sup>+</sup>). (<b>B</b>) <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> SSCs (<span class="html-italic">n</span> = 5/group) were cultured under undifferentiated or osteogenic conditions for 6 days, and protein levels of FZD5, β–catenin, and USP8 were assessed by immunoblotting. (<b>C</b>–<b>E</b>) <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> SSCs (<span class="html-italic">n</span> = 5/group) were cultured under osteogenic conditions for 6 days, and cell proliferation (C), alkaline phosphatase activity (ALP, D) and mRNA levels of <span class="html-italic">Opn</span> (E) were assessed. (<b>F</b>,<b>G</b>) <span class="html-italic">Usp8<sup>fl/fl</sup></span> and <span class="html-italic">Usp8<sup>Osx</sup></span> SSCs (<span class="html-italic">n</span> = 5/group) were transplanted beneath the kidney capsule of 8–week-old male mice (<span class="html-italic">n</span> = 4/group), and 8 weeks later bone formation was analyzed using X–radiography (left, red box) and microCT analysis. Representative 3D–reconstruction (F, left) and relative quantification (F, right) of bone mass in the kidney capsule were displayed. Alternatively, longitudinal sections of the kidney capsule were stained with Weigert Van Gieson (G). (<b>H</b>) A schematic diagram showing molecular actions of the DUB USP8 in the regulation of the Wnt/FZD/β–catenin pathway. USP8 deubiquitinates ubiquitinylated FZD5 through the C–terminal DUB domain (USP8C) in osteoblast precursors, important for Wnt/β–catenin signaling, the differentiation of osteoprogenitors to osteoblasts. Scale bars, 200 μm. Data are shown as box–and–whisker plot (with median and interquartile ranges) from min to max, with all data points shown. Analyses were performed using the unpaired Student’s <span class="html-italic">t</span>–test within Prism. NS, not significant.</p>
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29 pages, 7620 KiB  
Review
Comparative Approaches in Vertebrate Cartilage Histogenesis and Regulation: Insights from Lampreys and Hagfishes
by Zachary D. Root, Claire Gould, Margaux Brewer, David Jandzik and Daniel M. Medeiros
Diversity 2021, 13(9), 435; https://doi.org/10.3390/d13090435 - 10 Sep 2021
Cited by 3 | Viewed by 5404
Abstract
Jawed vertebrates (gnathostomes) have been the dominant lineage of deuterostomes for nearly three hundred fifty million years. Only a few lineages of jawless vertebrates remain in comparison. Composed of lampreys and hagfishes (cyclostomes), these jawless survivors are important systems for understanding the evolution [...] Read more.
Jawed vertebrates (gnathostomes) have been the dominant lineage of deuterostomes for nearly three hundred fifty million years. Only a few lineages of jawless vertebrates remain in comparison. Composed of lampreys and hagfishes (cyclostomes), these jawless survivors are important systems for understanding the evolution of vertebrates. One focus of cyclostome research has been head skeleton development, as its evolution has been a driver of vertebrate morphological diversification. Recent work has identified hyaline-like cartilage in the oral cirri of the invertebrate chordate amphioxus, making cyclostomes critical for understanding the stepwise acquisition of vertebrate chondroid tissues. Our knowledge of cyclostome skeletogenesis, however, has lagged behind gnathostomes due to the difficulty of manipulating lamprey and hagfish embryos. In this review, we discuss and compare the regulation and histogenesis of cyclostome and gnathostome skeletal tissues. We also survey differences in skeletal morphology that we see amongst cyclostomes, as few elements can be confidently homologized between them. A recurring theme is the heterogeneity of skeletal morphology amongst living vertebrates, despite conserved genetic regulation. Based on these comparisons, we suggest a model through which these mesenchymal connective tissues acquired distinct histologies and that histological flexibility in cartilage existed in the last common ancestor of modern vertebrates. Full article
(This article belongs to the Special Issue Evolution, Development, and Diversification of Vertebrates)
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<p>Overview of cyclostome cartilaginous head skeletons. (<b>A</b>) Larval lamprey. (<b>B</b>) Adult lamprey. (<b>C</b>) Adult hagfish. Colors correspond to suggested tissue contributions. Red: Pre-mandibular neural crest. Green: Mandibular/Post-mandibular neural crest. Blue: Mesoderm. Textures correspond to distinctions in cartilage type. Solid color: hard cartilage. Color with black stripes: soft cartilage. White with colored spots: cartilage-like tissues. Cartilage distinctions for larval and adult lampreys are adapted from Parker (1888) and Johnel (1948) illustrations and descriptions. Cartilage distinctions for hagfishes are adapted from Cole (1905) illustrations and descriptions. Keywords: BA: branchial arches; Hy: hyoid; LA: lingual apparatus; NA: neural arches; NC: nasal cartilage(s); Otc: otic capsule; PHP: post-hypophyseal processes; Tr: trabecula; VB: velar bar.</p>
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<p>Variants of cartilage in cyclostomes. (<b>A</b>) Lamprey ammocoete (top) and adult lamprey (bottom). (<b>B</b>) Adult hagfish. Letters correspond with their respective panels. (<b>C</b>) Hard cartilage from adult lamprey annular cartilage. (<b>D</b>) Hard cartilage from adult hagfish medial basal plate. Both hard cartilages are surrounded by abundant ECM. (<b>E</b>) Soft cartilage from lamprey ammocoete branchial arch. (<b>F</b>) Soft cartilage from adult hagfish lateral tentacular cartilages. Both soft cartilages are compact and have sparse ECM. (<b>G</b>) Mucocartilage from lamprey ammocoete ventral pharynx. (<b>H</b>) Pseudo-cartilage from adult hagfish posterior basal plate. Both cartilage-like tissues have fibroblast-like morphology with cartilage features. Panels D, F, and H were reproduced via Miyashita (2012) with permission from Tetsuto Miyashita.</p>
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<p>Differentiation of cartilage along the joint interface in osteichthyans. BMP signaling and FGF signaling contribute to initial mesenchymal condensation while genetic factors proportionally contribute to either articular and hypertrophic cartilage development. Increased TGFβ and hedgehog (HH)signaling along with <span class="html-italic">grem</span>, <span class="html-italic">gdf5</span>, and <span class="html-italic">sox9</span> activity drive prechondrogenic mesenchyme towards the articular cartilage fate (blue), with increased expression of archetypal cartilage extracellular matrix (ECM) genes like <span class="html-italic">acan</span> and <span class="html-italic">col2a1</span>. Conversely, increased WNT and BMP signaling drive these cells towards hypertrophy (red) and expression of bone-related ECM genes like <span class="html-italic">vcan</span>, <span class="html-italic">col1a1</span>, <span class="html-italic">col1a2</span>.</p>
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<p>Common variants of cartilage in tetrapods. (<b>A</b>) Alcian stain of hyaline cartilage from human trachea. Chondrocytes are embedded in a deep extracellular matrix rich in acidic proteoglycans. (<b>B</b>) Orcein stain of elastic cartilage from human epiglottis. Chondrocytes are similar to hyaline, but elastic fibers are more prevalent. Arrows indicate elastic fibers. (<b>C</b>) H&amp;E stain of fibrocartilage from human intervertebral disc. Chondrocytes are more interspersed, and collagenous fibers, indicated by arrows, run mostly parallel to one another. All three photographs depict tissue sections from the histological collection of the Department of Zoology, Comenius University in Bratislava.</p>
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<p>Molecular heterogeneity in larval lamprey cartilages during development. (<b>A</b>) Diagram of the larval lamprey skeleton at Tahara [<a href="#B131-diversity-13-00435" class="html-bibr">131</a>] stage T28–30. (<b>B</b>) Expression of <span class="html-italic">lecA</span> at stage T28. (<b>C</b>) Expression of <span class="html-italic">lecC</span> at T28. (<b>D</b>) Expression of <span class="html-italic">col9a1a</span> at stage T28. Certain ECM proteins are restricted to different regions of the larval skeleton. Scale bar approximately 250 μm. Keywords: BA: branchial arches; EB: epibranchial bar; EC: endostylic cartilages (also known as ventromedial lateral bars); HB: hypobranchial bar; LB: lateral bar; LL: lower lip; NC: nasal capsule; OH: oral hood; Otc: otic capsule; Tr: trabeculae; VB: velar bar; VMLB: ventromedial longitudinal bar; VLP: ventrolateral plate; VP: ventral pharynx mucocartilage.</p>
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<p>Histological differences in larval lamprey cartilages. (<b>A</b>) Left lateral view of stage T30 lamprey. Scale bar approximately 100 μm. (<b>B</b>,<b>D</b>,<b>F</b>) Toluidine blue staining of T30 sagittal sections of the anterior head skeleton. (<b>C</b>,<b>E</b>,<b>G</b>) Toluidine blue staining of T30 sagittal sections of the posterior pharyngeal skeleton. Lamprey cartilages can be divided between mucocartilage (pale purple matrix) and hyaline-like tissues (discoidal morphology, thin blue matrix), but there are further differences in morphology and histology within these groups. Keywords: BA: branchial arches; EB: epibranchial bar; HB: hypobranchial bar; LB: lateral bar; LL: lower lip; OH: oral hood; Tr: trabeculae; VMLB: ventromedial longitudinal bar; VP: ventral pharynx mucocartilage.</p>
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<p>Proposed spectrum of vertebrate cartilage types and their relatedness with one another as well as cell types like tendon. From a common mesenchymal precursor, cells receive continuous input from chondrogenic genes like <span class="html-italic">sox9</span> or tenogenic genes like <span class="html-italic">scx</span> or <span class="html-italic">mkx</span> and are driven towards an archetypal cell fate. These fates share expression of SLRP genes like <span class="html-italic">lumican</span>, <span class="html-italic">biglycan</span>, and <span class="html-italic">decorin</span>, and are also similarly affected by TGFβ and FGF signaling. Even after cells begin displaying tendon-like features, they may transdifferentiate to a cartilage-like fate through mechanotransduction. As a spectrum, there would be quantifiable differences in gene expression between individual cartilage types, both regulatory transcription factors and signaling ligands and receptors as well as ECM proteins.</p>
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34 pages, 2244 KiB  
Review
The Mandibular and Hyoid Arches—From Molecular Patterning to Shaping Bone and Cartilage
by Jaroslav Fabik, Viktorie Psutkova and Ondrej Machon
Int. J. Mol. Sci. 2021, 22(14), 7529; https://doi.org/10.3390/ijms22147529 - 14 Jul 2021
Cited by 9 | Viewed by 5221
Abstract
The mandibular and hyoid arches collectively make up the facial skeleton, also known as the viscerocranium. Although all three germ layers come together to assemble the pharyngeal arches, the majority of tissue within viscerocranial skeletal components differentiates from the neural crest. Since nearly [...] Read more.
The mandibular and hyoid arches collectively make up the facial skeleton, also known as the viscerocranium. Although all three germ layers come together to assemble the pharyngeal arches, the majority of tissue within viscerocranial skeletal components differentiates from the neural crest. Since nearly one third of all birth defects in humans affect the craniofacial region, it is important to understand how signalling pathways and transcription factors govern the embryogenesis and skeletogenesis of the viscerocranium. This review focuses on mouse and zebrafish models of craniofacial development. We highlight gene regulatory networks directing the patterning and osteochondrogenesis of the mandibular and hyoid arches that are actually conserved among all gnathostomes. The first part of this review describes the anatomy and development of mandibular and hyoid arches in both species. The second part analyses cell signalling and transcription factors that ensure the specificity of individual structures along the anatomical axes. The third part discusses the genes and molecules that control the formation of bone and cartilage within mandibular and hyoid arches and how dysregulation of molecular signalling influences the development of skeletal components of the viscerocranium. In conclusion, we notice that mandibular malformations in humans and mice often co-occur with hyoid malformations and pinpoint the similar molecular machinery controlling the development of mandibular and hyoid arches. Full article
(This article belongs to the Special Issue Neural Crest Development in Health and Disease)
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<p>Schematics of a frontal section through developing oral cavity of a mouse embryo at E12. <span class="html-italic">Dlx5/Dlx6</span> are expressed in ectomesenchyme of the mandibular process and the hyoid arch. <span class="html-italic">Hand1/Hand2</span> and <span class="html-italic">Meis2</span> are expressed in ectomesenchyme in the medial region of the mandibular process and the hyoid arch, which also includes the lingual ectomesenchyme. <span class="html-italic">Msx1</span> and <span class="html-italic">Runx2</span> are expressed within the primordium of prospective dentary bone. <span class="html-italic">Pax3</span> is expressed in the ectomesenchyme around the alveolingual sulcus, which represents an anatomical boundary between the dentary bone and tongue. <span class="html-italic">Shh</span> is expressed in the lingual epithelium, whereas <span class="html-italic">Foxf1/Foxf2</span> are expressed in the lingual ectomesenchyme. Abbreviations: E, embryonic day; MC, Meckel’s cartilage. Expression domains of genes written in black text are not highlighted in the figure.</p>
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<p>Schematic model of PA patterning along the dorsal–ventral axis in zebrafish at 30–36 hpf (hours post-fertilization). Lateral view, head to the left, pharyngeal endoderm in dark yellow, pharyngeal ectoderm in dark grey. The dorsal region of PAs is established by co-operative action of <span class="html-italic">jag1b</span> and <span class="html-italic">grem2</span> and characterized by the expression of <span class="html-italic">dlx2a</span>. The intermediate region of PAs is controlled by <span class="html-italic">dlx3b/4b/5a</span> and <span class="html-italic">msxe,</span> which are downregulated by <span class="html-italic">edn1</span> in the pharyngeal ectoderm. The ventral region of PAs is dominated by the expression of <span class="html-italic">bmp4-hand2</span>.</p>
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<p>Schematics of a mid-sagittal section through the mandibular and hyoid arches of a mouse embryo at E12. <span class="html-italic">Bmp4</span> is expressed in the distal mandibular epithelium, at the site of presumptive incisors, while <span class="html-italic">Msx1</span> is expressed in the distal mandibular ectomesenchyme, surrounding the incisor primordia. <span class="html-italic">Shh</span> is expressed in the vestibular lamina and dental epithelium, as well as in the lingual epithelium, while <span class="html-italic">Foxf1/Foxf2</span> are expressed in ectomesenchyme in the medial region of the mandibular process, surrounding the incisor primordium and inside the nascent tongue. <span class="html-italic">Pax3</span> is expressed in the ectomesenchyme of distal tip of the mandibular process and nascent tongue. <span class="html-italic">Lhx6/Lhx8</span> are expressed in ectomesenchyme on the rostral side, while <span class="html-italic">Gsc</span> is expressed on the caudal side of the mandibular process. <span class="html-italic">Meis2</span> is expressed in the medial proximal region of the mandibular process and in the medial region of the hyoid arch. Abbreviations: E, embryonic day; MC, Meckel’s cartilage; HB, cartilage primordium of the hyoid bone. Expression domains of genes written in black text are not highlighted in the figure.</p>
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<p>Schematics of the mandibular and hyoid regions of a mouse embryo at E12, superior view at the nascent tongue from inside of the alimentary canal. <span class="html-italic">Bmp4</span> and <span class="html-italic">Edn1</span> are expressed in the distal mandibular epithelium, while <span class="html-italic">Bmp4</span> also marks the site of presumptive incisors. <span class="html-italic">Shh</span> is expressed in lingual epithelium of the tongue primordium. <span class="html-italic">Fgf8</span> is expressed in the proximal mandibular epithelium, at the site of presumptive molars, while <span class="html-italic">Barx1</span> is expressed in the proximal mandibular ectomesenchyme, surrounding the molar primordia, and in ectomesenchyme of the hyoid arch. <span class="html-italic">Dlx5/Dlx6</span> are expressed in ectomesenchyme of the mandibular process and the hyoid arch. <span class="html-italic">Lhx6/Lhx8</span> are expressed in the ectomesenchyme on the rostral side, while <span class="html-italic">Gsc</span> is expressed on the caudal side of the mandibular process. Abbreviations: E, embryonic day; T, tongue primordium; HE, hypobranchial eminence (second PA); FC, foramen caecum. Expression domains of genes written in black text are not highlighted in the figure.</p>
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21 pages, 1464 KiB  
Review
Regulation and Role of Transcription Factors in Osteogenesis
by Wilson Cheuk Wing Chan, Zhijia Tan, Michael Kai Tsun To and Danny Chan
Int. J. Mol. Sci. 2021, 22(11), 5445; https://doi.org/10.3390/ijms22115445 - 21 May 2021
Cited by 121 | Viewed by 9676
Abstract
Bone is a dynamic tissue constantly responding to environmental changes such as nutritional and mechanical stress. Bone homeostasis in adult life is maintained through bone remodeling, a controlled and balanced process between bone-resorbing osteoclasts and bone-forming osteoblasts. Osteoblasts secrete matrix, with some being [...] Read more.
Bone is a dynamic tissue constantly responding to environmental changes such as nutritional and mechanical stress. Bone homeostasis in adult life is maintained through bone remodeling, a controlled and balanced process between bone-resorbing osteoclasts and bone-forming osteoblasts. Osteoblasts secrete matrix, with some being buried within the newly formed bone, and differentiate to osteocytes. During embryogenesis, bones are formed through intramembraneous or endochondral ossification. The former involves a direct differentiation of mesenchymal progenitor to osteoblasts, and the latter is through a cartilage template that is subsequently converted to bone. Advances in lineage tracing, cell sorting, and single-cell transcriptome studies have enabled new discoveries of gene regulation, and new populations of skeletal stem cells in multiple niches, including the cartilage growth plate, chondro-osseous junction, bone, and bone marrow, in embryonic development and postnatal life. Osteoblast differentiation is regulated by a master transcription factor RUNX2 and other factors such as OSX/SP7 and ATF4. Developmental and environmental cues affect the transcriptional activities of osteoblasts from lineage commitment to differentiation at multiple levels, fine-tuned with the involvement of co-factors, microRNAs, epigenetics, systemic factors, circadian rhythm, and the microenvironments. In this review, we will discuss these topics in relation to transcriptional controls in osteogenesis. Full article
(This article belongs to the Special Issue Osteoblast Differentiation and Activity in Skeletal Diseases)
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Graphical abstract

Graphical abstract
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<p>Current understanding of osteogenesis and bone remodeling. (<b>A</b>) Flat bones are formed through intramembranous ossification, cells are originated from the cranial neural crest (CNC). Limb bones are formed through endochondral ossification (details shown in (<b>C</b>)), cells are originated from the mesoderm-derived limb bud (LB) mesenchyme. (<b>B</b>) Two major routes for osteoblast differentiation. Mesoderm cells give rise to mesenchymal osteochondroprogenitors (OCPs) which can diverge into chondrocytic and osteoblastic lineages. Chondrocytes undergo hypertrophy and a portion of them differentiate into osteoblasts at the chondro-osseous junction. Neural crest-derived mesenchymal progenitors can differentiate directly to osteoblasts during intramembraneous ossification. (<b>C</b>) Endochondral ossification is a process of converting cartilage to bone and is essential for bone elongation. Cartilage anlagen of a future bone forming in the limb bud during embryogenesis. Chondrocyte hypertrophy (HC) initiates in the center of the anlagen where blood vessels (BVs) invade, bringing in osteoprogenitors and bone marrow cells. The primary spongiosa (PS) separates the cartilage into proximal and distal growth plates (GPs). From childhood to adolescence, there is an active proliferation of chondrocytes prior to hypertrophy, and the mineralizing cartilage is replaced by bone at the chondro-osseous junction (COJ). Thickening of cortical bone continues from birth to puberty when the GPs become inactive. (<b>D</b>) Bone remodeling maintains the integrity and homeostasis of bone in adulthood. Osteoclasts are bone resorptive cells originated from hematopoietic stem cells (HSCs). They remove microfractured segments of bone and mobilize osteoblasts to form new bone. Osteomorphs are a novel cell type generated through fission of osteoclasts. Subsequent fusion of osteomorphs can reform active osteoclasts. Multiple sources of skeletal stem cells (SSCs) and OCPs have been identified as the source of osteoblasts for bone formation. Some of the mature osteoblasts are embedded into the osteoid and further differentiate into osteocytes which have a critical role in bone remodeling coordination.</p>
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<p>Transcriptional regulation of osteoblast differentiation. SOX9 and RUNX2 are major fate determinants of mesenchymal progenitors to chondrogenesis and osteoblastogenesis, respectively. Cells can “detour” to chondrogenesis or commit to an osteoblast lineage. RUNX2 is the master transcription factor that regulates multiple steps in osteoblast commitment and differentiation. Its transcriptional activity is controlled at multiple levels such as transcriptional co-factors, inhibitors, osteo-enhancing and -suppressing miRNAs, and environmental cues such as light–dark cycle.</p>
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<p>Circadian clock in bone. (<b>A</b>) Suprachiasmatic nucleus (SCN) in the hypothalamus receives the 24 h light–dark signals and conveys them in the form of nerve or hormonal signals. The rhythmic level of hormone controls the peripheral clock in bone, hence leading to rhythmic expression of osteoblastic (OB) genes. (<b>B</b>) The molecular clock involves the positive regulators CLOCK and BMAL1 which bind to the E-box elements and activate expression of circadian negative regulators PER and CRY. PER and CRY inhibit activities of CLOCK and BMAL1 to form a feedback loop that occurs within a period of 24 h. CLOCK/BMAL1 can bind to the E-box region and activate expression of P300 which subsequently promotes the acetylation of histone 3 and facilitates the formation of a transcriptional complex with RUNX2 to drive expression of osteoblastic genes. Sirt1 has dual roles in the circadian clock and osteogenesis. It binds CLOCK/BMAL1 in a circadian manner and promotes the deacetylation and degradation of PER and is a positive regulator of RUNX2.</p>
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