Abstract

   Erythrocytes are the dominant component of a blood clot in terms of
   volume and number. However, longstanding compacted erythrocytes in
   blood clots form a physical barrier and make fibrin mesh more
   anti‐fibrinolytic, thus impeding infiltration of mesenchymal stem
   cells. The necrosis or lysis of erythrocytes that are not removed
   timely can also lead to the release of pro‐inflammatory toxic
   metabolites, interfering with bone regeneration. Proper bio‐elimination
   of erythrocytes is essential for an undisturbed bone regeneration
   process. Here, hypoxia‐mimicking is applied to enhance
   macrophage‐elimination of erythrocytes. The effect of
   macrophage‐elimination of erythrocytes on the macrophage intracellular
   reaction, bone regenerative microenvironment, and bone regeneration
   outcome is investigated. Results show that the hypoxia‐mimicking agent
   dimethyloxalylglycine successfully enhances erythrophagocytosis by
   macrophages in a dose‐dependent manner primarily by up‐regulating the
   expression of integrin α[v]β[3]. Increased phagocytosed erythrocytes
   then regulate macrophage intracellular Fe^2+‐glycolysis‐inflammation,
   creating an improved bone regenerative microenvironment characterized
   by loose fibrin meshes with down‐regulated local inflammatory responses
   in vivo, thus effectively promoting early osteogenesis and ultimate
   bone generation. Modulating macrophage‐elimination of erythrocytes can
   be a promising strategy for eradicating erythrocyte‐caused bone
   regeneration hindrance and offers a new direction for advanced
   biomaterial development focusing on the bio‐elimination of
   erythrocytes.

   Keywords: bone regeneration, erythrocyte clearance, hypoxia‐mimicking,
   inflammatory responses, macrophage phagocytosis
     __________________________________________________________________

   Longstanding compacted erythrocytes impair bone regeneration. This
   study employs the hypoxia‐mimetic agent DMOG to enhance
   macrophage‐elimination of erythrocytes by up‐regulating integrin
   α[v]β[3]. Increased phagocytosed erythrocytes regulate macrophage
   intracellular Fe^2+‐glycolysis‐inflammation, creating a favorable bone
   regenerative microenvironment characterized by loose fibrin meshes with
   down‐regulated inflammatory responses in vivo. These findings pave the
   way for advanced biomaterial development focusing on the
   bio‐elimination of erythrocytes.

   graphic file with name ADVS-11-2403921-g008.jpg

1. Introduction

   Coagulation is a natural process that forms a blood clot following
   injury to prevent excessive bleeding.^[ [42]^1 ^] The clot forms
   through the creation of a fibrin network that traps ≈95% erythrocytes,
   5% platelets, less than 1% leukocytes,^[ [43]^2 ^] and a variety of
   osteogenic and angiogenic cytokines.^[ [44]^3 ^] This clot not only
   recruits immune cells and mesenchymal stem cells (MSCs) but also acts
   as a temporary scaffold for these cells, initiating the healing of bone
   defects^[ [45]^4 ^] and playing a crucial role in determining the
   eventual outcome of the healing process.

   As the most abundant component of a blood clot, erythrocytes promote
   fibrin network forming during coagulation.^[ [46]^5 ^] However, upon
   the clot contraction, compacted erythrocytes form a physical barrier
   that impedes MSCs infiltration.^[ [47]^6 ^] In addition, longstanding
   erythrocytes interact with the fibrin network and make it denser,
   interfering with the normal fibrinolysis process,^[ [48]^7 ^] which is
   unfavorable for bone healing.^[ [49]^8 ^] Furthermore, necrosis or
   lysis of erythrocytes can release excess harmful metabolic chemicals
   including iron, which significantly compromises MSCs viability and
   triggers excessive inflammatory responses.^[ [50]^9 ^] Taken together,
   longstanding erythrocytes act as a neglected “badass” that reduces the
   osteogenic capacity of blood clots, thereby impeding natural bone
   repair. Therefore, “erythrocyte clearance” can be a promising strategy
   for enhancing bone regeneration.

   To effectively remove the overwhelming longstanding erythrocytes,
   especially in hematoma, mechanical scraping was performed to prevent
   excessive inflammation and infection, but it resulted in bone nonunion
   instead.^[ [51]^4 ^] The problem lies in the removal of the entire
   hematoma, which not only eliminates the harmful erythrocytes but also
   inadvertently removes beneficial elements, including growth factors, as
   well as osteogenic and angiogenic factors. Therefore, to properly
   handle erythrocyte clearance, precise bio‐elimination should be
   developed.

   Macrophages are major professional phagocytes that recognize and engulf
   apoptotic or necrotic cells via several pattern recognition and opsonic
   receptors expressed in their cell membranes.^[ [52]^10 ^] Moreover,
   macrophages have evolved solid mechanisms for iron ion separation,
   absorption, storage, utilization, and outflow to avoid iron toxicity,^[
   [53]^11 ^] providing them with a relatively high tolerance to
   erythrocytes. These prevent the inflammatory response of macrophages
   from being easily triggered during erythrophagocytosis, making
   macrophages the proper candidate for regulating the bio‐elimination of
   erythrocytes.

   Hypoxia has been demonstrated to enhance macrophage phagocytosis of
   apoptotic neutrophils and latex beads.^[ [54]^12 , [55]^13 ^]
   Additionally, hypoxia serves as a tissue injury repair signal as
   well,^[ [56]^14 ^] suggesting its potential use in promoting
   macrophage‐elimination of erythrocytes during bone regeneration. Oxygen
   deficiency inhibits the activity of the prolyl hydroxylase domain (PHD)
   enzymes, which prevents the degradation of the hypoxia‐inducible factor
   (HIF) and promotes its nuclear translocation, thus initiating the
   hypoxia responses.^[ [57]^15 ^] However, direct oxygen deprivation
   would not only cause harmful oxidative stress,^[ [58]^16 ^] but also
   facilitates the survival and proliferation of anaerobic bacteria,^[
   [59]^17 ^] resulting in infection and failed bone regeneration.
   Hypoxia‐mimicking can inhibit PHD activity without oxygen
   deprivation,^[ [60]^18 ^] making it a promising strategy for modulating
   macrophage‐elimination of erythrocytes. Dimethyloxalylglycine (DMOG)
   has shown higher specificity and biosafety in inhibiting PHD activity
   compared to agents like Co^2+ and desferrioxamine,^[ [61]^17 , [62]^19
   ^] making it a suitable candidate to modulate macrophage
   erythrophagocytosis.

   In this study, the dose‐dependent effect of DMOG on erythrophagocytosis
   of macrophages, and the consequent intracellular reactions and
   immunological impacts were first investigated in vitro. DMOG‐loaded
   blood clots were applied to rat calvarial defects to assess the effects
   of hypoxia‐mimicking on erythrocytes clearance, and the subsequent
   influence on inflammation and bone regeneration in vivo (Figure [63]1).
   This study highlighted the significance of the bio‐elimination of
   erythrocytes during bone regeneration, inspiring the development of
   advanced biomaterials focusing on the bio‐elimination of erythrocytes
   to promote bone regeneration. Additionally, this study confirmed that
   hypoxia‐mimicking can modulate macrophage‐elimination of erythrocytes,
   offering deeper insights into the function of hypoxia‐mimicking
   biomaterials.

Figure 1.

   Figure 1
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   The experimental flow of this study. A) The influence of DMOG on the
   expression of phagocytosis‐promoting genes was first investigated and
   then the DMOG pre‐stimulated macrophages were co‐cultured with
   erythrocytes to detect macrophage erythrophagocytosis. B) Intracellular
   reactions of macrophages after erythrophagocytosis were revealed by
   bioinformatic analysis. C) Changes in inflammatory responses were
   further explored and verified through immunofluorescence staining and
   RT‐qPCR. The possible effects on bone regeneration were analyzed
   through bioinformatic analysis. D) Blood clots loaded with DMOG were
   implanted into rat calvarial defects to verify the effect of DMOG on
   erythrocyte clearance, fibrin network degradation, and local
   inflammation in vivo through H&E staining and RT‐qPCR. E) The influence
   of changed bio‐elimination of erythrocytes on early osteogenesis and
   late new bone formation was detected through H&E staining, Micro‐CT
   scanning, and Masson's trichrome staining.

2. Results and Discussion

2.1. Macrophage Erythrophagocytosis was Enhanced by the Hypoxia‐Mimicking
Agent DMOG in a Dose‐Dependent Manner

   The safe concentration threshold of DMOG was initially assessed through
   the CCK8 assay. Results indicated that low concentrations of DMOG
   (0‐0.4 mm) had no significant impact on macrophage proliferation,
   whereas higher concentrations (0.8–1.0 mm) inhibited macrophage
   proliferation (Figure [65] 2A). These findings align with previous
   reports on DMOG's cytotoxicity toward monocytes and macrophages.^[
   [66]^20 ^] Subsequently, concentrations of 0–0.4 mm DMOG were applied
   to a rat calvarial bone defect model to evaluate in vivo systemic
   toxicity. Both macroscopic and microscopic observations revealed no
   signs of hyperemia, ischemia, necrosis, or atrophy in rat organs
   (Figure [67]S1A,B, Supporting Information), indicating no significant
   systemic toxicity. Consequently, DMOG concentrations of 0, 0.1, 0.2,
   and 0.4 mm were selected for further experiments.

Figure 2.

   Figure 2
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   Enhanced erythrophagocytosis of macrophages after stimulation with the
   hypoxia mimetic agent DMOG in gradient concentrations. A) Cell
   viability of macrophages after DMOG stimulation (n = 4). B) Gene
   expression heatmap of typical phagocytosis‐promoting molecules of
   macrophages after DMOG stimulation (n = 3). C) GO enrichment results
   and D) Interaction network analysis of the up‐regulated typical
   phagocytosis‐promoting molecules revealed that DMOG enhanced the EPC of
   macrophages by regulating recognition between integrin α[v]β[3] and
   phosphatidylserine. E) Diagram displays the mechanism by which DMOG
   pre‐stimulated macrophages phagocytose erythrocytes. F) Representative
   fluorescence images and G) Flow cytometry results of EPC in macrophages
   pre‐stimulated with DMOG demonstrated that DMOG effectively promoted
   the EPC of macrophages (n = 3). PS: phosphatidylserine; EPC:
   erythrophagocytosis; MFI: mean fluorescence intensity. Data presented
   as mean ± SD; ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p <
   0.0001, and ns means not significant by one‐way ANOVA with Tukey's post
   hoc test.

   To investigate the effect of DMOG on macrophage erythrophagocytosis
   (EPC), the gene expression of EPC‐promoting molecules, including
   receptors and bridging molecules was first examined by RT‐qPCR. Results
   showed that DMOG significantly up‐regulated the expression of tyrosine
   kinase receptor family molecules (AXL/MERTK), TIM4, integrin receptors
   (integrin α[v], integrin β[3]), and their bridging molecules (GAS‐6,
   THBS1, and MFG‐E8) (Figure [69]2B). TIM4 can directly recognize exposed
   phosphatidylserine on the erythrocyte surface,^[ [70]^21 ^] while
   AXL/MERTK and integrins can also bind phosphatidylserine through
   bridging molecules like GAS‐6 and MFG‐E8.^[ [71]^22 , [72]^23 ^] These
   upregulated genes were further subjected to Gene Ontology (GO)
   enrichment analysis to elucidate the potential mechanism by which DMOG
   enhances the phagocytic capacity of macrophages. The analysis revealed
   that these up‐regulated genes were mainly enriched in
   phosphatidylserine binding and integrin binding terms (Figure [73]2C).
   Protein‐protein interaction analysis revealed that bridging molecules
   MFG‐E8 and THBS1 are the most correlated, which primarily mediate the
   binding of integrin α[v]β[3] to the phosphatidylserine (Figure [74]2D).
   Taken together, these findings indicate that DMOG enhances macrophage
   EPC primarily through the integrin
   α[v]β[3]‐MFG‐E8/THBS1‐phosphatidylserine recognition (Figure [75]2E).

   To confirm the effect of DMOG on macrophage EPC, macrophages
   pre‐stimulated with DMOG were co‐cultured with fluorescently labeled,
   heat‐shocked erythrocytes that externalized phosphatidylserine. After
   2 h of co‐culture, erythrocyte engulfment was inconspicuous across all
   groups (Figure [76]2F,G). However, after 8 h, confocal images showed
   that DMOG increased the EPC of macrophages in a dose‐dependent manner,
   with the most significant effect observed in the 0.4 mm group. Flow
   cytometry results further indicated that DMOG promoted erythrocyte
   engulfment by increasing both the proportion of EPC‐positive
   macrophages and the phagocytic burden per individual macrophage, as
   reflected by intracellular mean fluorescence intensity
   (Figure [77]2F,G). After 16 h, although both DMOG pre‐stimulated and
   unstimulated groups showed a high proportion of EPC‐positive cells, the
   intracellular mean fluorescence intensity still exhibited a
   dose‐dependent increase (Figure [78]2F,G), indicating that DMOG
   primarily increased the phagocytic burden of macrophages during the
   later stages.

   In summary, within safe concentration thresholds, DMOG enhanced
   macrophage erythrophagocytosis by upregulating the expression of
   phagocytosis‐promoting molecules, particularly integrin α[v]β[3].

2.2. Increased Erythrocyte Engulfment Suppressed the Glycolysis Metabolism of
Macrophages

   To investigate the effect of increased erythrocyte engulfment on
   macrophage intracellular reactions, macrophages were pre‐stimulated
   with 0, 0.1, 0.2, and 0.4 mm DMOG for 24 h and then co‐cultured with
   heat‐shocked erythrocytes (designated as T0, T1, T2, and T4 groups,
   respectively). These macrophages were then collected for RNA sequencing
   (RNA‐seq) and bioinformatic analysis. The principal component analysis
   result showed that macrophages in the T1, T2, and T4 groups were
   segregated from those in the T0 group (Figure [79]S2A, Supporting
   Information), indicating that phagocytosed erythrocytes significantly
   altered the intracellular reactions of macrophages.

   Differential genes in the T1, T2, and T4 groups, compared to the T0
   group, were subjected to pathway annotation with the Kyoto Encyclopedia
   of Genes and Genomes (KEGG). Results showed that cell
   metabolism‐related pathways accounted for 3/15, 5/15, and 7/15 of the
   top 15 enriched pathways, respectively (Figure [80]S2B, Supporting
   Information), indicating that increased erythrocyte engulfment
   significantly altered macrophage metabolism. The interaction of these
   metabolism‐related genes was further investigated (Figure [81]S2C,
   Supporting Information), and the core genes were applied in GO
   enrichment analysis. Results revealed that glycolysis was the most
   significant event among the top 10 terms in the T2 and T4 groups
   (Figure [82]S2D, Supporting Information). Gene set enrichment analysis
   (GSEA) further demonstrated that the glycolysis pathway was
   significantly down‐regulated in the T1, T2, and T4 groups (Figure [83]
   3A), supported by the expression heatmap of glycolysis‐related genes
   (Figure [84]S2E, Supporting Information). RT‐qPCR results further
   verified that the expression of typical glycolytic enzyme genes was
   down‐regulated in the T1, T2, and T4 groups (Figure [85]3B), indicating
   that increased erythrocyte engulfment decreased glycolysis in
   macrophages.

Figure 3.

   Figure 3
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   Alternation of intracellular reactions of macrophages after increased
   erythrocyte engulfment. A) GSEA enrichment results of the glycolysis
   gene set demonstrated that glycolysis was significantly down‐regulated
   in T1, T2, and T4 groups (^*|NES| > 1, p <0.05, and FDR < 0.25). B) The
   gene expression of glycolysis‐related enzymes was investigated by
   RT‐qPCR (n = 3). C) GO enrichment analysis of the leading genes of
   glycolysis gene set further revealed the primarily affected processes
   of glycolysis. D) The Venn diagram analysis showed that Pfkl, Pfkm, and
   Pfkp were common genes among the selected terms, all encoding
   phosphofructokinase. E) The schematic diagram of glycolysis process
   with the primarily affected step marked with a red box. F) RT‐qPCR
   results showed that the gene expression of Pfkl, Pfkm, and Pfkp was
   significantly down‐regulated in T1, T2, and T4 groups (n = 3). G) The
   enzyme activity assay showed decreased enzyme activity of
   phosphofructokinase after enhanced erythrocyte engulfment (n = 4). H‐I)
   Representative immunofluorescence images of intracellular Fe^2+ and the
   semi‐quantitative statistical analysis results (n = 4). MFI: mean
   fluorescence intensity. Data presented as mean ± SD; ^* p < 0.05, ^**
   p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, and ns means not
   significant by one‐way ANOVA with Tukey's post hoc test.

   Glycolysis is a multi‐step biochemical reaction involving 10 enzymes
   and various intermediate metabolites.^[ [87]^24 ^] To identify which
   step of glycolysis was altered, leading genes of the glycolysis pathway
   were subjected to GO enrichment analysis. These genes were primarily
   enriched in terms related to “phosphorylation”, “fructose
   1,6‐bisphosphate metabolic process”, “fructose 6‐phosphate metabolic
   process”, and “glycolytic process through fructose 6‐phosphate”
   (Figure [88]3C). Venn diagram analysis revealed that these terms all
   included Pfkl, Pfkm, and Pfkp (Figure [89]3D), which encode
   phosphofructokinase, the key rate‐limiting enzyme that catalyzes the
   conversion of fructose 6‐phosphate into fructose 1,6‐bisphosphate,^[
   [90]^25 , [91]^26 ^] thereby controlling glycolytic flux
   (Figure [92]3E). It was found that Pfkl, Pfkm, and Pfkp were gradually
   down‐regulated in the T1, T2, and T4 groups (Figure [93]S2F, Supporting
   Information), which was confirmed by RT‐qPCR results (Figure [94]3F).
   An enzymatic activity assay further showed that phosphofructokinase
   activity was significantly decreased in the T2 and T4 groups
   (Figure [95]3G). These results indicate that macrophage glycolysis was
   reduced due to the decreased phosphofructokinase activity, limiting the
   phosphorylation of fructose 6‐phosphate following enhanced erythrocyte
   engulfment.

   Erythrocytes are a major natural reservoir of iron, which is contained
   in the heme.^[ [96]^27 , [97]^28 ^] When phagocytosed by macrophages,
   heme can be degraded into biliverdin, CO, and Fe^2+.^[ [98]^29 ^] It
   has been reported that Fe^2+ can inhibit phosphofructokinase
   activity.^[ [99]^30 , [100]^31 ^] Immunofluorescence images showed that
   intracellular Fe^2+ content increased in the T1, T2, and T4 groups
   (Figure [101]3H), which was further supported by the semi‐quantitative
   statistical result (Figure [102]3I). This increase in Fe^2+
   corresponded with the decreased activity of phosphofructokinase in the
   T1, T2, and T4 groups. Therefore, it was deduced that phagocytosed
   erythrocytes reduced phosphofructokinase activity and suppressed
   glycolysis through Fe^2+.

   In summary, the enhanced erythrocyte engulfment increased intracellular
   Fe^2+ content in macrophages, inhibiting phosphofructokinase activity
   and down‐regulating glycolysis metabolism.

2.3. Increased Erythrocyte Engulfment Down‐Regulated Macrophage Inflammatory
Responses That Modulate Bone Regenerative Events

   Glycolysis is the dominant metabolic pattern of classically activated
   macrophages (M1).^[ [103]^32 , [104]^33 ^] Additionally, it is widely
   reported that glycolysis affects macrophage inflammatory responses,^[
   [105]^34 , [106]^35 ^] and genes associated with glycolysis are closely
   correlated with those involved in inflammatory responses (Figure
   [107]S3A, Supporting Information). Therefore, it was hypothesized that
   decreased glycolysis would modulate macrophage M1 polarization and
   inflammatory responses.

   GSEA results showed that three M1 phenotype‐related terms, including
   macrophage activation, positive regulation of NO metabolic process, and
   positive regulation of ROS metabolic process, were significantly
   down‐regulated in the T1, T2, and T4 groups (Figure [108] 4A).
   Spearman's rank correlation analysis further revealed a strong positive
   correlation (R > 0.7) between glycolysis and these three M1
   phenotype‐related terms (Figure [109]4B), indicating that decreased
   glycolysis contributed to inhibited M1 polarization. Immunofluorescence
   staining and RT‐qPCR results further showed that the expression of
   typical M1 phenotype‐related markers (CCR7, TLR4, CD86) was
   significantly decreased in the T1, T2, and T4 groups
   (Figure [110]4C–E). Taken together, these results indicate that
   increased erythrocyte engulfment‐induced glycolysis suppression
   down‐regulated macrophage M1 polarization.

Figure 4.

   Figure 4
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   Down‐regulation of macrophage inflammatory responses after increased
   erythrocyte engulfment and the possible effects on bone regeneration
   events. A) GSEA enrichment results of macrophage M1 phenotype‐related
   gene sets (^*|NES| > 1, p <0.05, and FDR < 0.25). B) Spearman's rank
   correlation analysis of glycolysis and the above three M1
   phenotype‐related gene sets. C‐D) Representative immunofluorescence
   images of CCR7 and the semi‐quantitative statistical analysis results
   (n = 3). E) RT‐qPCR results of genes of typical M1‐marker TLR4 and CD86
   (n = 3). F) RT‐qPCR results of genes of typical inflammation‐related
   cytokines or chemokines (n = 3). G) Interaction between the top 30 core
   inflammatory genes and bone development‐related genes, as well as the
   GO enrichment results of the correlated genes, revealing that the top
   30 core inflammatory genes were closely associated with bone
   regeneration, especially influencing osteoblast differentiation. MFI:
   mean fluorescence intensity. Data presented as mean ± SD; ^* p < 0.05,
   ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, and ns means not
   significant by one‐way ANOVA with Tukey's post hoc test.

   Macrophage inflammatory responses after enhanced erythrophagocytosis
   were evaluated. GSEA results showed that “cytokine production involved
   in inflammatory responses”, “cytokine‐cytokine receptor interaction”,
   “chemokine production”, and “chemokine signaling pathway” were all
   significantly down‐regulated in the T1, T2, and T4 groups (Figure
   [112]S3B, Supporting Information). Spearman's rank correlation analysis
   confirmed significant positive correlations between the glycolysis gene
   set and these inflammatory response‐related terms (Figure [113]S3C,
   Supporting Information). RT‐qPCR results further showed that the
   expression of typical inflammatory cytokines (IL1β, IL6, and IL12a) and
   chemokines (CCL2, CCL3, CCL4, CCL5, and CXCL9) was significantly
   down‐regulated in the T1, T2, and T4 groups (Figure [114]4F). These
   combined results indicate that increased erythrocyte engulfment‐induced
   glycolysis suppression alleviated the inflammatory responses of
   macrophages.

   The possible effects of enhanced phagocytosed erythrocytes on immune
   responses and bone regenerative events were finally investigated. The
   expression of core genes related to inflammatory responses was first
   examined (Figure [115]S3D, Supporting Information) and their
   interactions were analyzed (Figure [116]S3E, Supporting Information).
   The top 30 core genes in the T1, T2, and T4 groups were identified
   according to the degree to perform interaction analysis with bone
   development‐related genes respectively. Results showed that these top
   30 core inflammatory genes in each group were closely correlated with
   bone development genes (Figure [117]4G). GO enrichment analysis further
   revealed that the correlated genes were enriched in the osteoblast
   differentiation term, ranking top 2 in significance for the T2 and T4
   groups, and fourth for the T1 group (Figure [118]4G). These findings
   suggest that enhanced phagocytosed erythrocytes‐induced macrophage
   immune responses could influence bone regeneration, particularly in the
   T2 and T4 groups.

   Collectively, enhanced erythrocyte engulfment‐induced glycolysis
   suppression inhibited M1 polarization, and reduced inflammatory
   responses, which could influence bone regeneration.

2.4. The Enhanced Bio‐Elimination of Erythrocytes Created Favorable Bone
Regenerative Microenvironment In Vivo

   As previously mentioned, longstanding compacted erythrocytes impair the
   infiltration and activity of MSCs.^[ [119]^6 , [120]^7 , [121]^8 ^] To
   assess whether “bio‐elimination of erythrocytes” could optimize the
   bone regenerative microenvironment in vivo, we first investigated the
   effect of DMOG on erythrocyte clearance and then characterized the
   local bone regenerative microenvironment, focusing on fibrin networks
   and inflammatory responses.

   Three days after surgery, erythrocyte clearance in blood clots was
   examined through hematoxylin and eosin (H&E) staining. In the control
   group, erythrocytes occupied the majority of the bone defect,
   indicating the dominant role of erythrocytes during bone healing
   (Figure [122] 5A‐a). Erythrocyte clearance began at the edge of the
   defect and progressed toward the center. The overall deep red area
   where erythrocytes were located in DMOG‐treated groups was
   significantly smaller compared to the control group (Figure [123]5A‐a;
   Figure [124]S4, Supporting Information). Erythrocytes were nearly
   removed at the edge of defects in DMOG‐treated groups, while they were
   scattered in the control group (Figure [125]5A‐b). At the quarter site
   of defects, the area occupied by erythrocytes was significantly smaller
   in the 0.2 and 0.4 mm groups than in the 0 and 0.1 mm groups
   (Figure [126]5A‐c). At the center of defects, tightly packed
   erythrocytes were observed in all groups (Figure [127]5A‐d).
   Erythrophagocytosis of macrophages was more apparent in the 0.2 and
   0.4 mm groups than in other groups (Figure [128]5A‐e). These results
   showed that in blood clots within rat calvarial defects, DMOG promoted
   macrophage‐mediated bio‐elimination of erythrocytes in a dose‐dependent
   manner.

Figure 5.

   Figure 5
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   The effect of enhanced bio‐elimination of erythrocytes on the bone
   regenerative microenvironment in vivo. A) H&E images of bone defects in
   different groups 3 days after surgery. (a) Overview of the bone defect.
   The general deep red area where erythrocytes lay in DMOG‐treated groups
   was significantly smaller than that of the control group, indicating
   increased bio‐elimination of erythrocytes. (b) The magnification images
   at the edge of defects with yellow arrows indicating dense matted
   deposits. (c) The magnification images at the quarter site of defects
   with the dotted line indicating the boundary of the area occupied by
   erythrocytes. (d) The magnification images at the center of defects
   displayed compacted erythrocytes. (e) Representative images of
   macrophages that have phagocytosed erythrocytes (indicated by yellow
   arrows). B) Heatmap showed that gene expression of the typical
   inflammation‐related cytokines and chemokines within defects was
   significantly down‐regulated after increased bio‐elimination of
   erythrocytes (n = 3). C) Interaction analysis showed these
   inflammation‐related cytokines and chemokines were closely correlated
   with each other. D‐E) IHC staining and semi‐quantitative analysis
   results of the expression of the universal macrophage marker CD68, the
   M1 macrophage marker iNOS, and the M2 macrophage marker CD206 (n = 9).
   Data presented as mean ± SD; ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001,
   ^**** p < 0.0001, and ns means not significant by one‐way ANOVA with
   Tukey's post hoc test.

   In addition to erythrocyte clearance, alterations in fibrin networks
   were also observed. Three days after surgery, relatively thick fibrin
   and compacted fibrin meshes with dense matted deposits, attributed to
   erythrocyte interference,^[ [130]^7 ^] were observed at the edge of
   defects in the 0 and 0.1 mm groups. In contrast, thin fibrin and loose
   fibrin meshes without dense matted deposits were observed in the 0.2
   and 0.4 mm groups (Figure [131]5A‐b). Similar results were noted at the
   quarter site of defects (Figure [132]5A‐c). However, the fibrin network
   was not visible at the center of defects due to the stacked
   erythrocytes that had not been phagocytosed (Figure [133]5A‐d). These
   results indicate that enhanced bio‐elimination of erythrocytes promotes
   the formation of a relatively loose fibrin network, which is beneficial
   for MSCs infiltration and bone regeneration.^[ [134]^36 , [135]^37 ^]

   Next, the effect of erythrocyte bio‐elimination on local inflammatory
   responses was determined. RT‐qPCR results showed a significant decrease
   in the gene expression of most inflammation‐related molecules in groups
   with increased erythrocyte bio‐elimination, particularly in the 0.2 and
   0.4 mm groups (Figure [136]5B). These molecules, including
   pro‐inflammatory cytokines (IL6, IL1β, TNF‐α, P65) and chemokines
   (CCL2, CCL3, CCL4, CCL5, and CXCL9), and M1 polarization markers (CD68,
   CD86, TLR4, CCR7, iNOS), have a close interaction (Figure [137]5C).
   Immunohistochemistry (IHC) staining further showed that compared to the
   0 mm group, the expression levels of CD68 and iNOS were significantly
   down‐regulated in the 0.1, 0.2, and 0.4 mm groups, while CD206 was
   slightly up‐regulated only in the 0.4 mm group (Figure [138]5D,E).
   These results indicate that enhanced erythrocyte bio‐elimination
   effectively limited macrophage M1 polarization and reduced local
   inflammatory responses.

   In summary, erythrocytes occupied the majority of the bone defect area,
   creating an unfavorable bone regenerative microenvironment.
   DMOG‐induced hypoxia‐mimicking promoted erythrocyte clearance within
   blood clots in rat calvarial defects through macrophage
   erythrophagocytosis. Enhanced bio‐elimination of erythrocytes created a
   loose fibrin mesh without dense matted deposits and reduced local
   inflammation, thereby optimizing the bone regenerative
   microenvironment.

2.5. The Enhanced Bio‐Elimination of Erythrocytes Promoted Early Osteogenesis
and Eventual Bone Regeneration

   Early osteogenesis was investigated 5 days post‐surgery via H&E and IHC
   staining. H&E images showed ongoing erythrocyte clearance, with some
   stacked erythrocytes accumulating in the middle of defects, and bone
   defects were filled with randomly arranged soft tissues (Figure
   [139]S5, Supporting Information). However, different events were
   observed at the edge of defects where erythrocyte clearance began
   first. In the 0 and 0.1 mm groups, fibrous‐like tissue with thin
   collagen fibers and small, long spindle‐shaped cells were observed
   (Figure [140] 6A). IHC images further revealed that there were few ALP,
   OPN, or COL1 positive cells, especially in the 0 mm group
   (Figure [141]6B,C). These results indicated that bone regeneration was
   significantly hindered due to insufficient erythrocyte clearance. In
   contrast, nascent bone matrix and thick collagen fibers were observed
   at the edge of defects in the 0.2 and 0.4 mm groups, embedded with
   various cells in spindle, oval, or oblong shapes (Figure [142]6A). IHC
   staining showed many of these cells were ALP, OPN, or COL1 positive
   (Figure [143]6B,C). Taken together, these results showed that compared
   to the 0 and 0.1 mm groups where erythrocyte clearance was inadequate,
   there were enhanced MSCs infiltration, osteoblast differentiation, and
   collagen secretion in the 0.2 and 0.4 mm groups, indicating that
   enhanced bio‐elimination of erythrocytes significantly promoted early
   osteogenesis.

Figure 6.

   Figure 6
   [144]Open in a new tab

   The effect of enhanced bio‐elimination of erythrocytes on the early
   osteogenesis and eventual bone regeneration. A) H&E images of edges of
   bone defects in different groups 5 days after surgery showed that
   DMOG‐mediated increased bio‐elimination of erythrocytes effectively
   facilitated osteogenesis. B‐C) IHC staining and semi‐quantitative
   analysis results of the expression of markers indicating osteoblast
   differentiation and osteogenesis, including ALP, OPN, and COL1 (n = 9).
   D) 3D reconstruction images of defects and E) Semi‐quantitative
   statistical analysis results of new bone volume showed apparent bone
   regeneration in 0.2 and 0.4 mm groups (n = 6). F) H&E staining (left,
   middle, and right scale bars = 1 mm, 200, and 100 µm, respectively) and
   Masson's trichrome staining (left, middle, and right scale bars = 1 mm;
   200 and 100 µm, respectively) images of the selected centric sections
   of bone defects, and G) Semi‐quantitative analysis results of
   non‐mineralized bone and mineralized bone (n = 3). Data presented as
   mean ± SD; ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001,
   and ns means not significant by one‐way ANOVA with Tukey's post hoc
   test.

   Four weeks post‐surgery, Micro‐CT 3D reconstruction images revealed
   minimal bone generation within calvarial defects in the 0 and 0.1 mm
   groups, with the defect centers nearly empty. In contrast, apparent new
   bone formation was observed in the 0.2 and 0.4 mm groups
   (Figure [145]6D), which was further supported by the semi‐quantitative
   statistical analysis of new bone volume (Figure [146]6E). H&E images
   revealed that bone defects in the 0 mm group were dominated by fibrous
   tissue, with only a few new bones present at the defect edge. Although
   bone‐like tissues slightly increased in the 0.1 mm group, the majority
   of the defect region was still occupied by fibrous tissue. In contrast,
   bone defects in the 0.2 and 0.4 mm groups were filled with confluent
   bone with larger new bone (Figure [147]6F), which aligned with the
   Micro‐CT results. Masson's trichrome staining results further showed
   significantly more mineralized bone in the 0.2 and 0.4 mm groups
   compared to the other groups (Figure [148]6F,G), indicating enhanced
   bone maturation.

   Taken together, these results showed that an optimized bone
   regenerative microenvironment created by enhanced bio‐elimination of
   erythrocytes effectively promoted early osteogenesis, late new bone
   regeneration and maturation.

2.6. Implications for Bone Regeneration‐Regulating Strategies and Advanced
Biomaterial Development Focusing on the Bio‐Elimination of Erythrocytes

   As aforementioned, erythrocytes are the dominant cells in blood clots
   and can affect the bone regeneration process, which has been ignored by
   researchers for a long time. Bio‐elimination of erythrocytes should be
   a promising strategy for improving bone regeneration outcomes. Although
   precise regulation of erythrocyte clearance is difficult, we succeeded
   in promoting erythrocyte removal through hypoxia‐mimicking‐induced
   enhanced erythrophagocytosis by macrophages. Phagocytosed erythrocytes
   then suppressed the glycolysis of macrophages by tuning
   Fe^2+‐phosphofructokinase, creating a down‐regulated inflammatory
   microenvironment with loose fibrin mesh in blood clots, and as a
   result, bone regeneration was enhanced (Figure [149] 7 ). This
   highlighted the significance of bio‐elimination of erythrocytes during
   bone regeneration and supported the feasibility of modulating
   macrophage‐elimination of erythrocytes to overcome bone regeneration
   hindrance attributed to erythrocytes, thus facilitating new bone
   formation.

Figure 7.

   Figure 7
   [150]Open in a new tab

   Schematic figure shows the effects of hypoxia‐mimicking on
   macrophage‐elimination of erythrocytes, and the subsequent
   immunomodulation and osteogenesis. Macrophage‐elimination of
   erythrocytes was enhanced by DMOG regulating integrin α[v]β[3]. The
   phagocytosed erythrocytes modulated the intracellular Fe^2+‐glycolysis
   interaction, thus reducing macrophage inflammatory responses. Enhanced
   bio‐elimination of erythrocytes in blood clots created an optimized
   bone regenerative microenvironment characterized by loose fibrin mesh
   without dense matted deposits and reduced local inflammation, which
   promoted bone regeneration. PS: phosphatidylserine. Schematics were
   created with BioRender.com.

   In this study, it was shown that the hypoxia‐mimicking agent DMOG
   promoted erythrophagocytosis of macrophages primarily by up‐regulating
   integrin α[v]β[3]. DMOG also influenced other
   erythrophagocytosis‐related receptors, including AXL, MERTK, and TIMD4.
   The underlying mechanism by which DMOG regulates the expression of
   phagocytosis‐related receptors probably involves activation of
   HIF‐1α.^[ [151]^12 , [152]^13 , [153]^38 , [154]^39 ^] Other
   hypoxia‐mimicking agents may similarly regulate these receptors and
   promote macrophage erythrophagocytosis, though their effects could vary
   due to differences in ways of HIF‐1α activation, safe concentration
   thresholds, and specificities. The elevated intracellular ferrous iron
   ion could trigger complex intracellular reactions. In addition to the
   primary down‐regulation of glycolysis and M1 polarization, other
   intracellular reactions and the consequent function changes of
   macrophages such as M2 polarization could also occur, which warrant
   further investigation. In vivo, the observed outcomes are a combination
   of multiple effects. Its promotion of osteogenic differentiation in
   mesenchymal stem cells^[ [155]^40 ^] may contribute to these outcomes.
   Additionally, DMOG‐stimulated macrophages not only phagocytize
   erythrocytes but may also engulf apoptotic neutrophils,^[ [156]^41 ^]
   which could also contribute to improved bone regeneration. However, the
   specific effects require further study.

   Erythrocyte phagocytosis involves the recognition between
   phagocytosis‐related receptors on the macrophage membrane and ligands
   on the erythrocyte surface, subsequent internalization, and
   regulation‐mediated by Piezo1‐mediated Ca^2+ influx.^[ [157]^42 ^] To
   manipulate erythrocyte phagocytosis, potential modulating strategies
   based on macrophages and erythrocytes have been proposed and summarized
   in Figure [158] 8 , which could guide the development of advanced
   biomaterials for erythrocyte clearance. By targeting key molecules in
   phagocytosis recognition, internalization, or regulation, advanced
   biomaterials that manipulate erythrocyte phagocytosis can be developed.
   For example, based on macrophages, biomaterials that modify the
   expression, conformation, or turnover of phagocytosis‐related membrane
   proteins can modulate the balance between “eat‐me” signaling and
   “do‐not‐eat‐me” signaling, thus manipulating phagocytosis. Based on
   erythrocytes, biomaterials that alter the morphology or rigidity of
   erythrocytes can tune the internalization process,^[ [159]^43 ^] thus
   modulating phagocytosis.

Figure 8.

   Figure 8
   [160]Open in a new tab

   Schematic diagram displays the strategy for manipulating
   erythrophagocytosis of macrophages to guide the development of advanced
   biomaterials focusing on the bio‐elimination of erythrocytes.
   Erythrocyte phagocytosis includes the recognition between
   phagocytosis‐related receptors on the macrophage membrane and ligands
   on erythrocyte surface and the subsequent internalization, as well as
   the regulation‐mediated by Piezo1‐mediated Ca^2+ influx. Stabilin2, TAM
   receptors (AXL, MERTK), and integrins (integrin α[V]β[3], integrin
   α[3]β[1]) can bind the externalized phosphatidylserine, and complement
   receptor 3 and Fc receptor are able to bind clustered band 3 to
   initiate the “eat‐me” signaling, while SIRPα binding CD47 can trigger
   “do not eat‐me” signaling. The internalization process involves not
   only the activity of myosin IIA of macrophages but also the shape and
   stiffness of erythrocytes. Based on macrophages, developing
   biomaterials that modify the expression, conformation, or turn‐over of
   phagocytosis‐promoting, phagocytosis‐inhibiting, and
   phagocytosis‐regulating proteins or the activation and assembly of
   myosin IIA can regulate the macrophage‐elimination of erythrocytes.
   Based on erythrocytes, developing biomaterials that modify the
   phosphatidylserine exposure and the band 3 clustering, block the CD47,
   or alter the morphology and rigidity of erythrocytes can also modulate
   the macrophage‐elimination of erythrocytes. PS: phosphatidylserine;
   TAM: Tyro3, Axl, and Mertk; CR3: complement receptor 3; SIRPα: signal
   regulatory protein alpha; Schematics were created with BioRender.com.

   Although local hypoxia develops following a bone injury, it is often
   insufficient to induce adequate erythrocyte clearance in time,
   resulting in impaired bone regeneration. The application of the
   hypoxia‐mimicking agent DMOG successfully accelerated
   macrophage‐elimination of erythrocytes, ultimately improving bone
   regeneration. These indicate that hypoxia‐mimicking biomaterials can be
   a future direction for developing advanced materials focusing on
   erythrocyte clearance to improve bone regeneration. This study
   discovered a new biological role of hypoxia‐mimicking in regulating
   macrophage‐mediated erythrocyte clearance, offering deeper insights
   into the hypoxia‐mimicking‐mediated bone regeneration, which aligns
   with the “blood‐activating and stasis‐transforming” concept in
   Traditional Chinese Medicine.^[ [161]^44 ^] Specifically,
   hypoxia‐mimicking treatment could hasten the elimination of compacted
   erythrocytes that form a physical barrier, thus promoting the entry of
   new blood and cells essential for tissue regeneration, resulting in an
   activated, favorable microenvironment instead of a stagnant one.
   Moreover, release‐controlling and sustained‐release hypoxia‐mimicking
   biomaterials can be further developed to precisely regulate erythrocyte
   phagocytosis by manipulating the hypoxia degree spatiotemporally.

3. Conclusion

   The hypoxia‐mimicking agent DMOG enhanced macrophage elimination of
   erythrocytes mainly by up‐regulating the expression of integrin
   α[v]β[3]. The phagocytosed erythrocytes decreased macrophage
   inflammatory responses by regulating intracellular
   Fe^2+‐phosphofructokinase‐glycolysis. Bio‐elimination of erythrocytes
   in vivo made a loose fibrin mesh without dense matted deposits and
   reduced local inflammation, thus creating an optimized bone
   regenerative microenvironment that effectively promoted bone
   regeneration. Modulating macrophage elimination of erythrocytes could
   be a promising and valuable strategy for overcoming erythrocyte‐caused
   bone regeneration hindrance.

4. Experimental Section

Cell Culture

   RAW 264.7 cells (referred to as RAW cells hereafter) were selected as
   the model cell to research macrophage erythrophagocytosis. RAW cells
   were cultured in a complete medium composed of Dulbecco's Modified
   Eagle's Medium (DMEM, Gibco, USA), 10% fetal bovine serum (Gibco, USA),
   and 1% penicillin/streptomycin (Gibco, USA), and culture flasks or
   plates were placed in cell incubators (37 °C, 5% CO2).

Cytotoxicity Assay of DMOG

   Cell‐Counting Kit‐8 (CCK8) (#CK04, Dojindo, Japan) was used to assess
   the cytotoxicity of DMOG on RAW cells. After the stimulation, DMOG
   solutions were removed and replaced with a culture medium containing
   10% CCK8 reagent, followed by incubation at 37 °C for another 1 h. Then
   the solution in each well was transferred to another new plate and its
   absorbance at 450 nm was measured using a microplate reader. DMOG was
   further loaded into blood clots that were implanted into rat calvarial
   bone defects to assess the in vivo systemic toxicity of DMOG.

Erythrocytes Isolation, Fluorescence Labeling, and Eryptosis Induction

   Erythrocytes were isolated from the blood of BALB/c mice. All
   experimental procedures were reviewed and approved by the Institutional
   Animal Care and Use Committee of Sun Yat‐Sen University and performed
   following the principles of animal protection, animal welfare, and
   ethics (approval NO. SYSU‐IACUC‐2023‐001132). Eight‐to‐nine‐week‐old
   BALB/c mice were anesthetized by intraperitoneal injection with 2%
   pentobarbital sodium at a ratio of 2.5 mL kg^−1. The whole blood was
   collected as previously described.^[ [162]^45 ^] Then the whole blood
   was washed with ice‐cold PBS followed by centrifugation at 500 rcf
   until the supernatant was clear. The red erythrocyte deposits were
   collected for subsequent experiments. Erythrocytes were labeled with a
   Cell Plasma Membrane Staining Kit with Dil (Beyotime, China) according
   to the manufacturer's instructions. Eryptosis was induced by 56 °C heat
   shock for 5 min in a water bath according to a previous report.^[
   [163]^46 ^]

Evaluation of the Influence of DMOG on Erythrophagocytosis In Vitro

   Within the safe concentrations, RAW cells were pre‐stimulated with DMOG
   for 24 h and then subjected to RT‐qPCR to detect the expression of
   phagocytosis‐related genes. Primers applied in this part are listed in
   Table [164]S1 (Supporting Information). Up‐regulated genes were
   submitted to DAVID Bioinformatics Resources, and GO enrichment was
   performed to analyze their main functions. Data visualization was
   performed using SangerBox 3.0
   ([165]http://vip.sangerbox.com/home.html). The interaction of these
   upregulated molecules was performed through the String database. RAW
   cells pre‐stimulated with DMOG for 24 h were co‐cultured with
   heat‐shocked erythrocytes in a new complete medium. High content cell
   real‐time analysis system (PerkinElmer, USA) and flow cytometric
   analysis were used to detect macrophage erythrophagocytosis at 2, 8,
   and 16 h after co‐culture, respectively.

Investigation of Effects of Enhanced Erythrophagocytosis on Macrophages

   After co‐culture with erythrocytes for 16 h, RAW cells were divided
   into T0, T1, T2, and T4 groups respectively, with each group having
   three biological duplicates. These RAW cells were subjected to RNA‐seq,
   RT‐qPCR, and Immunofluorescence staining. Primers applied in this part
   are listed in Table [166]S1 (Supporting Information).

RNA‐Seq

   Total RNA of RAW cells was extracted. Quality evaluation was performed
   through a fragment analyzer. According to the 150‐bp paired‐end run
   protocol provided by BGI genomics (Shenzhen, China), the cDNA library
   was constructed and sequenced on a BGISEQ‐500 platform. Low‐quality
   reads were excluded, and clean reads were mapped to the mouse reference
   genome using Bowtie2 and HISAT. Gene expression levels were expressed
   as fragments per kilobase of exon model per million mapped fragments
   (FPKM).

Bioinformatic Analysis

   Principal Component Analysis was applied to assess the general
   differences among the four groups. Differential expression analysis was
   carried out through the R package DESeq2. Differentially expressed
   genes between two different groups were used for KEGG pathway
   enrichment analysis through DAVID Bioinformatics Resources.
   Protein‐protein interactions (PPI) were obtained from the STRING
   database. The interaction network was loaded into Cytoscape, and the
   core sub‐network was sorted out using MCODE plug‐in, with the degree
   calculated. GSEA was performed to analyze the differences in target
   gene sets between two groups.^[ [167]^47 ^] Target genes were subjected
   to DAVID Bioinformatics Resources and GO enrichment was performed. Venn
   diagrams were used to identify intersected genes among several target
   terms. Gene set variance analysis (GSVA) scores of target gene sets for
   each sample were calculated using R package GSVA, and were used for
   Spearman's rank correlation analysis to assess the correlation of two
   target gene sets. All data visualization was performed with OmicStudio
   tools at [168]https://www.omicstudio.cn/tool, GraphPad Prism, TBtools,
   and Cytoscape.

Immunofluorescence Staining

   RAW cells that had engulfed erythrocytes were fixed with 4%
   paraformaldehyde (Beyotime, China) and then washed with PBS. These RAW
   cells were blocked with 1% bovine serum albumin (BSA; Amresco, USA) in
   PBS and then incubated with CCR7 (#ab32527, Abcam, UK) antibody at 4 °C
   overnight. After washing with PBS, RAW cells incubated with Alexa Fluor
   488 (#EM35141‐01, EMAR, China) fluorescent secondary antibody at room
   temperature for 1 h, followed by PBS wash. RAW cells were then stained
   with 4,6‐diamino‐2‐phenyl‐indole (DAPI; Beyotime, China) for 10 min.
   The fluorescent images were obtained through a confocal laser scanning
   microscope (Olympus FV3000, Japan). The intracellular content of Fe^2+
   was detected by a FerroOrange staining kit (#F374, Dojindo, Japan)
   according to the manufacturer's instructions. Semi‐quantitative
   analyses were performed with Image‐J.

Animal Surgery

   Eight to nine weeks old SD rats were subjected to in vivo experiments.
   All the surgical procedures were reviewed and approved by the
   Institutional Animal Care and Use Committee, Sun Yat‐Sen University,
   and met the requirements of animal protection, animal welfare and
   ethics (approval NO. SYSU‐IACUC‐ 2022‐000153). The rats were
   anesthetized by intraperitoneal injection with 2% pentobarbital sodium
   at a ratio of 2.5 ml kg^−1. Following the previously reported typical
   surgical procedures,^[ [169]^48 ^] two circle defects of 5 mm diameter
   were separately created on the left and right side of the rat calvaria
   with a drill. Blood clots with DMOG at different concentrations were
   prepared according to our previous study.^[ [170]^49 ^] Briefly, blood
   was collected from the tail vein and mixed with DMOG normal saline
   solutions (0, 1, 2, 4 mm) in a 9:1 volume ratio to obtain clots with
   DMOG at final concentrations of 0, 0.1, 0.2, and 0.4 mm. Each clot was
   made into 200 µL in a round container. After coagulation, these clots
   were separately implanted into circle calvarial bone defects and a
   layered suturing was performed to close the wound. The calvaria were
   collected as samples for histological analysis at different time
   points.

Erythrophagocytosis Assay In Vivo

   To evaluate the influence of DMOG on erythrophagocytosis in vivo, rats
   were sacrificed with excessive anesthesia 3 days after implantation.
   Calvaria samples were fixed with 4% paraformaldehyde. After
   decalcification, these samples were embedded in paraffin and 4 µm thick
   sections were generated using a Leica Biosystems RM2255 (Leica,
   Germany) for subsequent histological assay. For H&E staining, briefly,
   after dewaxing and hydration, Mayer's hematoxylin was applied to stain
   cell nuclei and eosin was applied to stain cell plasm and extracellular
   matrix on selected slides.

Characterization of The Local Inflammatory Microenvironment In Vivo

   Three days after the surgery, macrophage polarization within and near
   the defects was first investigated through immunohistochemical (IHC)
   staining of CD68, iNOS, and CD206. After dewaxing and hydration, the
   selected sections were incubated with 3% H[2]O[2] for 30 min to
   eliminate endogenous peroxidase activity, blocked with 1% BSA at room
   temperature for 1 h, and then incubated with CD68 (#ab125212, Abcam,
   UK), iNOS (#ab283655, Abcam, UK), and CD206 (#ab64693, Abcam, UK)
   antibodies at 4 °C overnight. After that, the slides were incubated
   with a goat anti‐rabbit secondary antibody (Gene Tech, China) at room
   temperature for 1 h. The antibody‐antigen complex was visualized by
   adding diaminobenzidine solution (Gene Tech, China). The slides were
   then counterstained with Mayer's hematoxylin for 2 min, dehydrated, and
   mounted. Semi‐quantitative statistics of positive cells in images were
   performed using Image‐J with an IHC profiler plug‐in. To detect the
   gene expression profile of typical inflammatory‐related cytokines or
   chemokines at the defects, blood clots within calvarial defects were
   collected 3 days after the surgery and total RNA of the clot tissue was
   extracted using TRIZOL (Beyotime, China) and the expression of
   inflammation‐related molecules was detected through RT‐qPCR. Primers
   applied in this part are listed in Table [171]S1 (Supporting
   Information).

Evaluation of Bone Regeneration

   Five days after the surgery, H&E staining was applied to observe
   osteogenesis, and IHC staining of alkaline phosphatase (ALP) (#ab65834,
   Abcam, UK), osteopontin (OPN) (#bs‐0026R, Bioss, China), and collagen I
   (COL I) (#ab34710, Abcam, UK) was applied to further verify cell types
   embedded in the new bone matrix. Four weeks after the surgery, calvaria
   were scanned through a Micro‐CT scanner (µCT50; SCANCO Medical AG,
   Switzerland) at a resolution of 10 µm, a source voltage of 70 kV, and a
   current of 114 µA. 3D reconstruction images of the bone defects were
   generated. The new bone volume was calculated using the software
   Materialise Mimics Research 19.0 (Materialise, Belgium). H&E staining
   was further used to observe the new bone formation and new bone
   mineralization. Masson's trichrome staining was performed with a
   Masson's trichrome staining kit (Solarbio, China) according to the
   manufacturer's instructions.

Statistical Analysis

   Each experiment was repeated at least three times, and the sample size
   (n) was noted in the figures. Experimental results were presented as
   mean ± standard deviations. For statistical analysis, GraphPad Prism
   was used and the built‐in One‐way ANOVA was performed followed by
   recommended Tukey's multiple comparison post hoc test in all cases. A P
   value of less than 0.05 was considered significant (without further
   explanation, ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p <
   0.0001, ns means not significant).

Conflict of Interest

   The authors declare no conflict of interest.

Supporting information

   Supporting Information
   [172]ADVS-11-2403921-s001.docx^ (13.6MB, docx)

Acknowledgements