Adult skull bone marrow is an expanding and resilient hematopoietic reservoir

doi:10.21203/rs.3.rs-3426773/v1

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Adult skull bone marrow is an expanding and resilient hematopoietic reservoir

https://doi.org/10.21203/rs.3.rs-3426773/v1

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published 13 Nov, 2024

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The bone marrow microenvironment is a critical regulator of hematopoietic stem cell self-renewal and fate. While it is appreciated that aging, chronic inflammation and other insults compromise bone marrow function and thereby negatively affect hematopoiesis, it is not known whether different bone compartments exhibit distinct microenvironmental properties and functional resilience. Here, we have employed imaging, pharmacological approaches, and mouse genetics to uncover specialized and highly surprising properties of bone marrow in adult and aging skull. Specifically, we show that the skull bone marrow undergoes lifelong expansion involving vascular growth, which results in an increasing contribution to total hematopoietic output. Furthermore, skull is largely protected against major hallmarks of aging, including upregulation of inflammatory cytokines, adipogenesis and loss of vascular integrity. Striking dynamic and rapid changes to the skull vasculature and bone marrow are induced by physiological alterations, namely pregnancy, but also pathological challenges, such as stroke and experimental chronic myeloid leukemia. These responses are highly distinct from femur, the most extensively studied bone marrow compartment. We propose that skull harbors a protected and dynamically expanding bone marrow microenvironment, which is relevant for experimental studies but, potentially, also clinical treatments in humans.

Biological sciences/Physiology/Ageing

Biological sciences/Physiology/Bone

Biological sciences/Physiology/Cardiovascular biology

bone marrow

skull

long bone

blood vessels

angiogenesis

hematopoiesis

aging

Niche microenvironments composed of multiple cell types and molecular signals regulate the self-renewal and the ultimate fate of hematopoietic stem cells (HSCs) in bone marrow^1-3. The skull, a flat bone encasing the brain, develops via intramembranous ossification and fundamentally differs in both composition and remodeling from long bones, which undergo endochondral ossification during development and sustain load-bearing mechanical stress^4-6. The thin and flat mouse calvarium, the main component of the skull roof, has been very successfully used as a platform for the imaging of HSC dynamics^7-9. Potential functional differences between bone marrow (BM) compartments, such as those in long and flat bone, however, remain little understood. It has been shown that HSCs are homogenously distributed throughout calvarium and long bone (namely diaphysis and epiphysis) under homeostatic conditions, whereas chimerism levels of mice transplanted with congenic hematopoietic stem and progenitor cells (HSPCs) are initially higher in calvarium and epiphysis relative to diaphysis¹⁰. Recently, the discovery of designated channels between the calvarial BM and the dura mater of the meninges indicates that skull BM acts as a hematopoietic reservoir for meningeal and central nervous system immunity^11-15. Aging causes an overall decline in hematopoietic function, with compromised lymphopoiesis and a bias toward dysfunctional myelopoiesis^16-19. Again, it remains unknown whether all BM compartments are equally affected by aging processes and therefore collectively contribute to hematopoietic decline. Here, we show that calvarial BM is fundamentally different from the BM in long bone. Calvarial BM continuously expands during adulthood, is surprisingly resistant to most major hallmarks of BM niche aging, and gradually increases its systemic hematopoietic contribution throughout adult life and aging. Our work also establishes that BM compartments exhibit unexpected functional differences, with implications for a range of physiological conditions, such as pregnancy and aging, but also pharmacological treatments and disease settings.

Calvarial bone marrow expands during adult life

To characterize changes in calvarial bone marrow throughout adult life, we harvested skulls from young adult (10-14 weeks), middle-aged (31-37 weeks), old (52-75 weeks) and geriatric mice (95+ weeks). Initial stereoscopic observation showed that blood-filled structures, representing blood vessels and surrounding BM, occupy only limited areas of the frontal and parietal bone in young adults, whereas interparietal bone is mostly filled by blood cells (Fig. 1a). While angiogenic growth of blood vessels is normally confined to development, regeneration or certain pathological conditions^20,21, substantial expansion of the vasculature can be seen in calvaria from middle-aged mice (Fig. 1a-d). Further increases in vascular area and vessel diameter can be observed in old and geriatric skulls (Fig. 1a-d). Skull cross-sections from young and geriatric animals show that the enlargement of calvarial BM occurs more rapidly in female mice relative to males (Fig. 1a; Extended Data Fig. 1a, b). Expansion of calvarial BM results in an increase in total skull thickness (~70% between young and geriatric mice from both sexes) without decreasing the thickness of the cortical bone tables (Extended Data Fig. 1a, b). Indicating that bone resorption by osteoclasts contributes to BM expansion, immunofluorescence (IF) analysis of skull sections shows a profound increase in V-type proton ATPase-positive activated osteoclasts lining the surface of the inner and outer cortical bone tables in skulls from old relative to young mice (Extended Data Fig. 1c, d).

To address whether skull BM expansion also occurs in aging humans, we analyzed head computerized tomography (CT) scan images obtained from 36 human patients evaluated for small cerebral aneurysms. This revealed significant increases of the area enclosed by the inner and outer cortical bone tables between younger (21-40 years) and older (61-69 years) individuals (Extended Data Fig. 2a-c). A higher difference was seen in female skulls (83.2% increase versus 24.6% in males), indicating a sexual dichotomy in both mice and humans. These results show that substantial expansion of the BM volume occurs both in murine and human skull during adulthood and aging.

Vascular growth and remodeling during skull BM expansion

Previous studies have shown that arterial and sinusoidal endothelial cells are important components of the HSC microenvironment^22,23. To gain more insight into the organization of the calvarial vasculature, we first utilized an in vivo immunofluorescence staining method with anti-CD31 antibody in Flk1-GFP knock-in mice²⁴. This approach revealed profound differences in vascular architecture seen in young adult frontal, parietal and interparietal skull bones (Extended Data Fig. 3a, c). Specifically, larger Flk1⁺ CD31⁺ sinusoidal vessels with distal sprouts dominate in frontal bone, thinner Flk1⁺ CD31⁺ sinusoidal vessel loops connect to Flk1^lo^w^/- CD31⁺ arterioles in parietal bone, and interparietal bone is completely filled by a dense network of sinusoidal vessels and arterioles (Extended Data Fig. 3a, c). Vascular leakage assays with Evans Blue corroborated local differences in vascular properties in different skull parts and revealed significantly higher leakage in frontal bone compared to the other two skull bones (Extended Data Fig. 3b, d).

To gain more insight into the organization of the calvarial vasculature and the changes during adulthood and aging, we performed in vivo IF labeling with fluorescent anti-CD31 and anti-Endomucin antibodies. While only interparietal bone is largely filled by a dense vascular network in young adults, substantial and sustained vascular expansion can be seen in frontal and parietal bones of middle-aged and old mice (Fig. 1b-d). Vascular area increased by 6.4- and 3.6-fold in frontal and parietal bones, respectively. Endomucin, which decorates calvarial sinusoidal vessels but not arterioles, has been implicated in bone maintenance and HSC function^25-27. Vascular Endomucin expression significantly increases until the geriatric stages (3.1-fold increase in frontal and 7.0-fold increase in parietal bone). We next focused on parietal bone, which shows the largest BM expansion and a transition from thin looping sinusoidal vessels in young adults to a dense network in later stages (Fig. 1b, c). Sinus diameter significantly increases 4.8-fold (Fig. 1c, d) and dorsal-ventral vertical extension of the vascular network can be observed at the old stage (Extended Data Fig. 3e). IF staining of Caveolin-1^27-29 shows a significant 2.0-fold increase of interconnecting arterioles during aging (Extended Data Fig. 3f, g). In vivo IF staining of hematopoietic cells using fluorescent anti-CD45 antibody shows that vascular growth goes along with a substantial increase in hematopoietic cells (13.1-fold increase between young and geriatric) within the expanding BM (Extended Data Fig. 4a, b). CD45⁺ cells are predominantly associated with large-caliber sinusoidal vessels, which are highly abundant in old and geriatric skull (Extended Data Fig. 4a). The link between vessel caliber and BM is further confirmed by a genetic approach labeling hematopoietic cells in the Vav-Cre Rosa26-mTmG background^30,31, which shows that hematopoietic cells expressing green fluorescent protein (GFP) surround large-caliber sinusoidal vessels (Extended Data Fig. 4c). Likewise, CD3ɛ⁺ T lymphocytes, B220⁺ B lymphocytes and CD11b⁺ cells (including monocytes/macrophages, granulocytes, and natural killer cells) are associated with large-caliber sinusoids in skull BM (Extended Data Fig. 4d). With the exception of interparietal bone, BM-associated sinusoidal vessels are comparably sparse in young adult animals and increase substantially throughout adult life and aging.

Interparietal bone, which is largely filled by vessels and BM at the young adult stage, shows minimal age-related changes in vascular area (24.3% increase between young and geriatric) and sinus diameter (9.3% decrease between young and geriatric) but a significant increase in Endomucin expression (2.0-fold increase between young and geriatric samples) (Fig. 1b, d). In stark contrast, vascular density in the metaphyseal region of the femoral BM decreases significantly by 45.2% between the middle-aged and old stages, but rebounds at geriatric stages with the emergence of very thin vessels displaying high Endomucin expression (Extended Data Fig. 5a, b), which have been recently characterized as dysfunctional sinusoidal vessels³². Electron micrographs of the luminal surface of ECs in young and old skull bone marrow show comparable patterns of endothelial fenestrations, which are a hallmark of vessels in primary lymphoid organs including BM. By contrast, ECs in old femur BM show a highly irregular pattern of fenestrae and the emergence of larger pores in the endothelial surface (Extended Data Fig. 5c). In contrast to the increasing vasculature of skull marrow, vessel density in the dura mater, which is adjacent to the skull and connected directly to the skull BM via specialized channels^11,33, decreases significantly during aging (Extended Data Fig. 5d, e). Collectively, these results show continuous expansion of the BM and vasculature in the adult and aging calvarium but not in the adjacent dura mater. Moreover, aging femoral BM displays features of compromised vascular integrity, which are not seen in aging skull BM.

Dynamic and distinct skull BM responses in pathophysiology

High demand for certain blood cells leads to strongly increased hematopoiesis in a range of physiological but also pathological conditions^1,34,35. Pregnancy, for example, induces extramedullary hematopoiesis (EMH) and the rapid expansion of maternal blood volume and red blood cells^36,37. Skull BM and associated vasculature increase substantially during pregnancy, namely 1.8-fold at 17 days postcoitum (dpc) and 2.3-fold at 2 days postpartum (dpp), which involves substantial vessel enlargement in frontal and parietal bone together with an increase in DAPI⁺ BM area (Fig. 2a, b). As reported previously³⁸, Endomucin^hightype H vessels in femur increase strongly at late-stage pregnancy and decrease postpartum, but overall vascular density in the metaphyseal region shows more limited changes (26.2% increase at 17 dpc and 8.5% decrease at 2 dpp) relative to skull (Extended Data Fig. 6a, b). Ischemic stroke in a transient mid-cerebral artery occlusion model (tMCAO) was shown to activate hematopoietic bone marrow stem cells³⁹. Consistent with this finding, tMCAO induces a striking expansion of skull BM and associated vasculature (2.1-fold increase), filling a substantial portion of parietal bone within 7 days (Fig. 2c, d). By contrast, femoral BM vascular density decreases by 26.0% in the stroke condition (Extended Data Fig. 6c, d). Hematological malignancies are known to lead to BM alterations, which, in turn, facilitate disease progression⁴⁰. A chronic myeloid leukemia (CML) model shows minimal but significant expansion of the skull BM and associated blood vessels (39.1% increase), which retain a healthy vascular morphology (Fig. 2e, f). By contrast, substantial loss of vascularity (50.4% decrease) and severe vascular rarefaction can be observed in CML femoral BM (Extended Data Fig. 6e, f). Biologically active fragments of parathyroid hormone (PTH) are not only used to treat osteoporosis⁴¹ but also lead to HSPC expansion⁴². We find that 4 weeks of daily PTH treatment results in substantial skull BM expansion (4.6-fold increase), increased large-caliber vessels and expansion of vessel-associated BM area (Fig. 2g, h), whereas minimal changes were observed in femoral BM (Extended Data Fig. 6g, h). These results demonstrate surprising dynamic plasticity of skull BM in a range of pathophysiological contexts and uncover unexpected differential responses in calvarial versus femoral BM.

Skull BM-resident HSPCs drive BM and vasculature expansion during aging

Hematopoietic stem and progenitor cells (HSPCs) residing in bone marrow change drastically during aging, both in quantity and differentiation potential^43,44. In order to address whether such cells might promote skull BM expansion, we isolated lineage-negative (Lin-) cells from the skull of young or old mice and transplanted them into lethally-irradiated young recipients (Fig. 3a). In vivo IF staining shows substantial expansion (2.2-fold) of skull BM vessels in recipient mice that have received cells from old donor skull BM compared to young donors (Fig. 3b, c). Interestingly, vascular density in femur is not significantly different in recipients of transplants from young or old donors (Extended Data Fig. 7a, b), further arguing for divergent responsiveness of different BM compartments. FACS analyses of HSPCs show that both long-term HSCs (LT-HSCs) and Lin^-Sca1⁺cKit⁺ (LSK) HSPCs increase in number in the aging skull BM (Fig. 3d, e), similar to what has been reported previously for aging long bone^45-47. To further test whether HSPCs contribute to BM expansion and vascular growth in skull, we treated mice with prostaglandin E2 (PGE2)^48-50 or G-CSF^51,52 to modulate HSPC number. PGE2 treatment increases the number of HSPCs in the skull BM by 92.3% (Extended Data Fig. 7c, d), resulting in significant expansion of calvarial vasculature and vessel-associated BM relative to control mice (Fig. 3f, g). G-CSF treatment mobilizes HSPCs into the peripheral blood by 8.2-fold (Extended Data Fig. 7e, f) and induces a significant 56.0% reduction of skull BM and vasculature relative to control mice (Fig. 3h, i). Although both PGE2 and G-CSF lead also to changes in vascular density in femoral BM, these alterations are confined to existing marrow without increase in BM area (Extended Data Fig. 7g-j). These results indicate that treatments acting on HSPCs can dynamically modulate the skull BM and its vasculature.

Signaling via VEGF-A and VEGFR-2 drives skull BM and vasculature expansion

VEGF-A is a master regulator of vascular growth^53,54. Since HSPCs can dynamically modulate the skull BM and our previous study found HSPCs isolated from long bone to express high levels of Vegfa transcripts compared to other BM cell types⁵⁵, we performed qRT-PCR analyses of various hematopoietic and stromal cell populations after flow cytometric sorting from enzymatically digested skull BM. LSK HSPCs indeed show highest Vegfa expression compared to Lin^-Sca1^-cKit⁺ cells (4.72-fold), Lin^-Sca1^-cKit^- stromal cells (5.97-fold) or Lin⁺ mature hematopoietic cells (5.92-fold) in young adult mice (Fig. 4a). Furthermore, skull BM-resident HSPCs isolated from old mice show significantly higher Vegfa expression (2.26-fold) compared to the same population from young mice. HSPCs isolated from aged skull show highest Vegfa expression relative to Lin^-Sca1^-cKit⁺ cells (8.39-fold), Lin^-Sca1^-cKit^- stromal cells (10.59-fold) and Lin+ mature hematopoietic cells (9.86-fold) (Fig. 4a). Further analyses show that expression of transcripts for the Vegfa-120 and Vegfa-164 isoforms is substantially higher in HSPCs compared to other cell types and also higher in old mice compared to young animals (Extended Data Fig. 8a). At the protein level, ELISA of whole BM lysates shows that VEGF-A is lower in skull BM relative to femoral BM but increases 2.2-fold from young to geriatric stages (Fig. 4b). Anti-VEGF-A IF staining of skull sections confirms high expression of the growth factor by resident cKit⁺ HSPCs (Fig. 4c). These results indicate that the increase in HSPC number and the concomitant increase in VEGF-A expression in these cells during aging contribute to an overall increase in VEGF-A levels, thereby promoting skull BM expansion. By contrast, VEGF-A levels in femoral BM gradually drop by 40.2% during aging (Fig. 4b). As hypoxia is a critical regulator of VEGF-A expression in angiogenesis^56-58, we administered Hypoxyprobe (Pimonidazole hydrochloride) to analyze age-related changes in the oxygenation of skull and femur (Extended Data Fig. 8b, c). Hypoxyprobe signal is significantly higher in skull BM relative to femoral BM at old (3.3-fold) and geriatric stages (4.8-fold) (Fig. 4d, e; Extended Data Fig. 8b, c), consistent with the observed increase of VEGF-A in aging skull. Next, we manipulated VEGF-A-dependent VEGFR2 signaling in vivo to address the role of this pathway in the expansion of skull vasculature and BM. For gain-of-function experiments, a cDNA encoding a bone-homing version of Vegfa (see Methods) was cloned into the vector pLIVE, which allows constitutive protein expression in liver after hydrodynamic tail vein injection⁵⁹. Vegfa overexpression induces a 2.5-fold increase of the skull BM and associated vasculature (Fig. 4f, g). Conversely, inhibition of VEGFR2 signaling with the blocking antibody DC101 inhibits BM expansion and vascular growth during aging (Fig. 4h, i). By contrast, vascular density changes within the confines of the femoral BM can be observed, but these alterations do not involve any substantial volumetric changes of the BM like in the skull (Extended Data Fig. 9a-d). These results demonstrate that VEGF-A/VEGFR2 signaling regulates skull BM expansion during aging and the increasing HSPCs in aging skull appear to be a relevant source of VEGF-A.

The skull BM microenvironment exhibits resistance to aging

The aging BM microenvironment disrupts normal self-renewal and other essential functions of HSCs⁶⁰. To assess whether hallmarks of aging affect calvarial and femoral BM equally, we characterized adipogenesis and inflammation in both compartments in young and geriatric mice. Adipocytes have previously been reported to accumulate in the aging BM and negatively regulate HSC function^61,62. Wholemount or cross-sectional staining of skull or femur, respectively, with BODIPY shows minimal changes in lipid-filled mature adipocytes in the skull, whereas a substantial 5.9-fold increase of adipocytes can be observed in the metaphyseal region of the aging femoral BM (Fig. 5a, b). Next, a bead-based multiplex array of 12 cytokines was used to assess the inflammatory milieu of lysates from skull and femoral BM isolated from young and old mice. Strikingly, only IFN-γ is significantly upregulated in old skull, whereas 7 out of 12 cytokines, including TNF-α and IL-6, are significantly elevated in old femur relative to young femur (Fig. 5c). It should also be noted that the expression of most cytokines in old femur is substantially higher than in skull, further supporting that the aging femoral BM is more inflammatory than its skull counterpart. Aging HSCs display a myeloid bias at the expense of lymphopoiesis^63,64. To compare the myeloid output from skull vs. hindlimb BM into peripheral blood, we selectively shielded these areas during lethal irradiation of young and old mice (Fig. 5d). While there was no difference in myelopoiesis between head or leg shielding in young mice, peripheral blood output from hindlimbs of old mice shows a strong myeloid bias compared to skull (Fig. 5e). Altogether, the results above show that calvarial BM is remarkably resilient against key features of aging and thereby maintains a healthy microenvironment for hematopoiesis even in old mice.

Skull BM maintains a functional hematopoietic niche during aging

It has been established that skull BM contains functional HSCs that can fully reconstitute the entire hematopoietic system in a transplantation setting¹⁰ and is the source of meningeal immune cells^12,13. The impact of aging on hematopoietic output by different BM compartments, however, has not been explored. Following lethal irradiation in a non-transplantation setting, all mice with whole body exposure died within 12 days from bone marrow failure (Fig. 5f). While vascular density (10.2-fold) and diameter (3.3-fold) are substantially increased in femoral BM at 7 days after irradiation, vessels in skull show minimal changes, suggesting higher resilience against irradiation-induced damage in experiments both with and without BM transplantation (Extended Data Fig. 10a-c). All young adult mice protected by shielding of the head or hindlimbs survived past 16 weeks without transplantation of exogenous BM, demonstrating that both body compartments contain sufficient HSCs for survival (Fig. 5f). Furthermore, shielding of the head of old mice is also fully compatible with survival. By contrast, 62.5% of old mice survived when only hindlimbs were protected (Fig. 5f), consistent with the notion that the BM microenvironment is deteriorating in aging femur but not in skull. To further assess the hematopoietic function of skull BM in a physiological context devoid of irradiation injury, we expressed the green-to-red photoconvertible fluorescent protein, Kikume Green-Red (KikGR)⁶⁵ in hematopoietic cells under control of Vav-Cre³⁰ (Fig. 5g). Following in vivo photoconversion of hematopoietic cells in calvarial BM, FACS analysis of peripheral blood revealed a striking age-dependent increase in systemic hematopoietic contribution from photoconverted BM cells (Fig. 5h). These results indicate that systemic hematopoietic output from skull increases relative to other BM compartments during aging.

In summary, our findings reveal an unexpected level of heterogeneity between two major BM compartments that are both well-established model systems in experimental studies. In contrast to almost all other organ systems and long bone, which become fully vascularized during embryonic and postnatal development^66,67, blood vessel growth in skull persists throughout adult life. Lifelong angiogenesis in skull involves the continued formation of large-caliber sinusoidal vessels, which are strongly associated with a range of hematopoietic cell populations and thereby represent a central landmark in the skull BM environment. Vascular growth and BM expansion in skull rely on VEGF-A and its receptor VEGFR2, which are key regulators of developmental and regenerative angiogenesis in many organs^54,68. Consistent with our previous work⁵⁵, skull BM-resident HSPCs express more VEGF-A than Lin⁺ differentiated hematopoietic cells, but other relevant sources of VEGF-A might exist, as has been shown for chondrocytes and osteoprogenitors in long bone^69,70. Our experiments also establish that circulation-derived VEGF-A can promote calvarial vessel growth and BM expansion, which is consistent with the finding that systemic VEGF can increase lifespan and delay aging processes in mice⁷¹. Conversely, HSPC behavior is controlled by the BM endothelium and associated reticular cells, which provide critical niche signals, such as stem cell factor (SCF/Kitl)⁷² or CXCL12 (SDF-1)⁷³. Several studies have shown that hematopoietic cell production is enhanced in response to various physiological but also pathological stimuli, such as pregnancy and ischemic stroke^74,75. Our new findings indicate that these conditions lead to the expansion of calvarial BM but induce only limited alterations in load-bearing long bone, where BM growth may not be compatible with mechanical loading and structural integrity. Furthermore, our results establish that calvarial BM is more resistant to aging-related degenerative processes, namely adipogenesis, the upregulation of inflammatory cytokines and compromised vascular integrity. This might, in part, reflect that calvarial BM is biologically “younger” due to its formation during adult life. Alternatively, unknown cellular or molecular features might differentially influence hematopoiesis in different BM compartments. Consistent with the latter, recent work has uncovered that skull BM exhibits a distinct molecular profile in health and neurological disorders⁷⁶. Remarkably, our findings also suggest that BM expansion occurs in human skull, which is consistent with previously published observations^77,78. Thus, our findings regarding the plasticity and resilience of the BM in skull may have wider relevance for aging and disease processes in humans but also pharmacological treatments targeting angiogenic signaling pathways.

Animal models

C57BL/6J mice were used for all experiments involving wild-type mice and pharmacological treatments. Mice at the age of 10-14 weeks, 31-37 weeks, 52-75 weeks and >95 weeks were chosen for young, middle-aged, old and geriatric groups, respectively. Both female and male mice of all age groups were used for initial bone marrow expansion analyses, while only female mice were used for remaining experiments. Flk1-GFP reporter mice²⁴ were used for initial blood vessel characterization. For genetic approach labeling of hematopoietic cells, Vav1-Cre animals³⁰ were interbred with ROSA26-mTmG reporter mice³¹ to generate Vav-mTmG mice. For photoconversion of hematopoietic cells, Vav1-Cre mice were interbred with ROSA26-CAG-loxP-stop-loxP-KikGR knock-in mice⁶⁵ to generate Vav-KikGR mice. For pregnancy experiments, 10-week-old C57BL/6J female mice were paired with 10~12-week-old C57BL/6J male mice and the onset of pregnancy was determined by the presence of a vaginal plug in the morning. 10-week-old mice received daily intraperitoneal injections of PTH (1-34) (Bachem, 0.1 mg kg^-1 for 28 days), PGE2 (Cayman Chemical, 2 mg kg^-1 for 7 days), G-CSF (Peprotech, 0.2 mg kg^-1 for 14 days) before sacrifice. For DC101 treatment, 10-week-old mice received intraperitoneal injections of DC101 (BioXCell, 40 mg kg^-1) every 2 days for 9 weeks.

Mice were kept in individually ventilated cages (IVC), with constant access to food and water under a 12h light and 12h dark cycle regime. Air flow, temperature (21-22°C) and humidity (55-60%) were controlled by an air management system. Animals were checked daily and maintained in specific pathogen-free (SPF) conditions. Sufficient nesting material and environmental enrichment was provided. All animal experiments were performed according to the institutional guidelines and laws, approved by local animal ethical committee and were conducted at the Max Planck Institute for Molecular Biomedicine (84-02.04.2016.A160, 81-02.04.2018.A171, 81-02.04.2020.A212, 81-02.04.2020.A416 and 81-02.04.2022.A198), Universitätsmedizin Berlin (G0220/17) and the Georg-Speyer-Haus (F123/2017) under the indicated permissions granted by the Landesamt für Natur, Umwelt und Verbraucherschutz (LANUV) of North Rhine-Westphalia, the State Office for Health and Social Affairs Berlin and the Regierungspräsidium Darmstadt, Germany. Mouse lines and other unique biological materials described in this article are either available through stock centers, commercial suppliers or, upon reasonable request, the lead author.

Human subjects, CT acquisition and data analysis

The study was approved by the local ethics committee and the institutional review board (IRB) of Asan Medical Center, and the requirement for informed consent was waived due to the retrospective nature of the study (IRB number: 2023-0658). The study population consisted of 36 patients, divided into four groups according to age (between 20 and 40 years, over 60 years) and gender (male, female), with nine patients in each group. Patients who underwent CT for evaluation of small cerebral aneurysm from April to May 2023 were eligible. Patients were excluded if they had a previous history of surgery or radiation therapy to the head and neck, vascular or bone-related medical implants, or a suspicious disease other than small cerebral aneurysm.

All human patients underwent CT examinations on the same 128-channel multidetector CT system (Somatom Definition Edge; Siemens, Erlangen, Germany). Imaging variables were as follows: 100 kV; 100 effective mAs; axial scan mode; section thickness, 0.5mm; display FOV, 20.5 cm; pitch, 1; gantry rotation time, 0.5 seconds; pixel matrix, 512 ⅹ 512. Images were obtained from the vertex to 1^st cervical spine, without an IV injection of contrast media.

The CT data were digitally transferred to a personal computer and processed with ImageJ software (http://rsb.info.nih.gov/ij/). A representative image was selected on a coronal CT image perpendicular to the outermost convex area on an axial CT image. After whole bone segmentation of the parietal bone, cortical bone and bone marrow were defined by attenuation densities on CT scan: cortical bone as over 850 Hounsfield units and bone marrow as less than 850 Hounsfield units and their areas were calculated.

Sample processing and immunostaining

Mice were sacrificed by transcardial perfusion of PBS and 4% paraformaldehyde (PFA), skulls and femur were harvested and fixed immediately in ice-cold 4% PFA for 6-8h under gentle agitation. Bones were decalcified in 0.5M EDTA for 3 days (for skulls) or 7 days (for femurs) at 4°C under gentle shaking agitation, washed 5 times in PBS in 5 min intervals, followed by overnight incubation in cryoprotectant solution (20% sucrose, 2% PVP) and embedding in bone embedding medium (8% gelatin, 20% sucrose, 2% PVP). Samples were stored overnight at -80°C. 80 μm-thick cryosections were prepared for immunofluorescence staining.

Bone sections were washed in PBS and permeabilized with 0.3% Triton-X-100 in PBS for 10 minutes at room temperature (RT). Samples were incubated in blocking solution (5% heat-inactivated donkey serum in 0.3% Triton-X-100) for 1 hr at RT. Primary antibodies (rat monoclonal anti-Endomucin (V.7C7) (Santa Cruz, Cat# sc-65495, 1:200 dilution), rabbit monoclonal anti-vATPaseB1/B2 (Abcam, Cat# 200839, 1:200 dilution), goat polyclonal anti-Osteopontin (R&D Systems, Cat# AF808, 1:200 dilution), goat polyclonal anti-CD31 (R&D, Cat# AF3628, 1:200 dilution), rabbit polyclonal anti-Caveolin 1 (Cell Signaling, Cat# 3238, 1:100), goat polyclonal anti-VEGF₁₆₄(R&D Systems, Cat# AF-493-NA, 1:200 dilution), rat monoclonal APC-conjugated anti-CD117 (c-Kit) (BD Biosciences, Cat# 553356, 1:100 dilution) were diluted in PBS with 5% donkey serum and incubated overnight at 4°C. Next, slides were washed 3-5 times in PBS in 5 min intervals. Species-specific Alexa Fluor-conjugated secondary antibodies Alexa Fluor 488 (Thermo Fischer Scientific, Cat# A21208), Alexa Fluor 594 (Thermo Fischer Scientific, Cat# A21209), Alexa Fluor 647 (Thermo Fischer Scientific, Cat# A31573 or Cat# A21447) diluted 1:500 in PBS with 5% donkey serum were added and incubated overnight at 4°C. Slides were washed 3-5 times in PBS in 5 min intervals. Nuclei were counterstained with DAPI (Sigma-Aldrich, Cat# D9542, 1:1000 dilution). Coverslips were mounted with FluoroMount-G (Southern Biotech, Cat# 0100-01).

In vivo immunostaining and Evans Blue leakage assay

Rat monoclonal anti-CD31 (BD Biosciences, Cat# 553708) was conjugated to Alexa Fluor 647 using the Alexa Fluor 647 Antibody Labeling Kit (Thermo Fischer Scientific, Cat# A20186) according to the manufacturer’s instructions. For blood vessel immunostaining, the conjugated anti-CD31 antibody and rat monoclonal PE-conjugated anti-Endomucin (V.7C7)(Santa Cruz, Cat# 65495 PE) were diluted 1:10 in 200 ml PBS and injected intravenously into the tail vein. For hematopoietic cell immunostaining, rat monoclonal FITC-conjugated anti-CD45 (eBioscience, Cat# 11-0451-82), hamster monoclonal FITC-conjugated anti-CD3e (eBioscience, Cat# 16-0031-82), rat monoclonal PE-conjugated anti-CD45R/B220 (BD Biosciences, Cat# 553090), rat monoclonal FITC-conjugated anti-CD11b (BD Biosciences, Cat# 553310) were diluted 1:10 in PBS and injected intravenously into the tail vein. Mice were sacrificed 1 hr after injection with trans-cardial perfusion with PBS and 4% PFA and bones were harvested and fixed immediately in ice-cold 4% PFA for 6-8h under gentle agitation. The dura mater was carefully removed from the skull with forceps. Bones were decalcified in 0.5M EDTA for 1 day (for skulls) or 7 days (for femurs) at 4°C under gentle shaking agitation, and washed 5 times in PBS in 5 min intervals. Skulls were counterstained with DAPI (1:500 dilution) for 1 hr and trimmed down to the calvarium before mounting with iSpacers (Sunjin Lab, Cat# IS011) in PBS. Femurs were cryosectioned, counterstained and mounted as described above.

For the Evans Blue leakage assay, mice were anaesthetized immediately prior to tail vein injection of 200 ml Evans Blue solution (Sigma-Aldrich, Cat# E2129, 1% v/w). Animals were sacrificed via transcardial perfusion 5 minutes after injection as described above. In order to distinguish vascular leakage in the dura mater from the calvarial bone marrow, dura mater tissues were separated from the calvarial bone before overnight decalcification.

Scanning electron microscopy

Skull and femur from 12-week-old and 73-week-old mice were isolated and submerged in 4% paraformaldehyde, 0.5% glutaraldehyde, 2 mM MgCl₂, 2 mM CaCl₂ in 0.1 M cacodylate buffer, pH 7.4, under agitation for 2 hours at room temperature. Samples were fixed further overnight in 2% glutaraldehyde, 2 mM MgCl₂, 2 mM CaCl₂ in 0.1 M cacodylate buffer, pH 7.4 at 4 °C. Bones were then decalcified over 12 days, changing solution every other day in 5% EDTA in 0.1 M cacodylate buffer, pH 7.4 under rotation at 4 °C. Subsequently, 150 μm sections were generated with a vibratome (VT 1200, Leica, Viennna, Austria). Sections were post-fixed in 1% OsmO₄, containing 2.5% paraformaldehyde-glutaraldehyde mixture buffered with 0.1M phosphate (pH 7.2) for 5 hours and then were placed in graded ethanol for critical point drying using E3000 (Polaron) critical point dryer. Critical point dried bones were placed on a piece of carbon tape and sputter coated with gold in a SC502 Sputter Coater (Polaron). Specimens were imaged on a Quanta 250 Field Emission Scanning Electron microscope (FEI Quanta 250 FEG, FEI, Hillsboro, OR) installed at the Korea Research Institute of Bioscience & Biotechnology.

Dura mater wholemount immunostaining

Mice were sacrificed by trans-cardial perfusion of PBS and 4% paraformaldehyde (PFA), skulls were harvested and fixed immediately in ice-cold 4% PFA for 6-8h under gentle agitation. Skulls were decalcified in 0.5M EDTA for 24 hours at 4°C under gentle shaking agitation, washed 5 times in PBS in 5 min intervals, trimmed down to the calvarium, and incubated in blocking solution (5% heat-inactivated donkey serum in 0.3% Triton-X-100) for 1 hr at RT. Goat polyclonal anti-CD31 (R&D, Cat# AF3628, 1:100 dilution) was diluted in PBS with 5% donkey serum and incubated overnight at 4°C with gentle agitation. Samples were washed 3-5 times in PBS in 10 min intervals. Alexa Fluor 647 (Thermo Fischer Scientific, Cat# A21447) diluted 1:500 in PBS with 5% donkey serum was added and incubated overnight at 4°C with gentle agitation. Samples were washed 3-5 times in PBS in 10 min intervals and mounted with iSpacers (Sunjin Lab, Cat# IS011) in PBS.

Transient Middle Cerebral Artery Occlusion (tMCAO)

16-week-old female C57BL6/J mice were used throughout the experiments. Animals were anesthetized by intraperitoneal injection of a mixture of 10 mg kg^-1 xylazine (cp-pharma) and 90 mg kg^-1 ketamine hydrochloride (cp-pharma). Throughout the whole procedure and during recovery, body temperature was maintained at 37°C via a heating pad. After ligation of the left proximal common carotid artery and external carotid artery, a 7.0-nylon monofilament (Doccol Co., NM, USA) with a 0.23mm coated tip was introduced into the distal internal carotid artery via an incision in the ligated common carotid artery. The monofilament was advanced distal to the carotid bifurcation to occlude the middle cerebral artery. Arter topical application of the local anesthetic lidocaine hydrochloride (Xylocain Spray 2%, Aspen) the neck wound was closed temporarily for a 45 min ischemic period. At reperfusion, the monofilament was withdrawn from the carotid artery and the wound was stitched with 4-0 non-resorbable sutures (Ethibond Excel, Ethicon) and the single s.c. injection of Penicillin G 20 000 U (Benzylpenicillin-Natrium, InfectoPharm) was given. The animal was returned to its cage to recover under observation.

Chronic myeloid leukemia

6-week-old female C57BL/6 mice were purchased from Charles River Laboratories (Sulzfeld, Germany) and were used as donors and recipients in all transplants. The transplantation experiments were performed as previously described⁷⁹. In brief, to induce chronic myeloid leukemia (CML)-like myeloproliferative neoplasia (MPN), donor BM cells from 5-fluorouracil (5-FU) intravenously pre-treated donor mice (200 mg/kg intravenously; 4 days prior to harvest) were pre-stimulated overnight in medium containing stem cell factor (SCF) (50 ng/ml), IL-6 (10 ng/ml) and IL-3 (6 ng/ml) and transduced on two consecutive days with MSCV IRES GFP BCR-ABL1 (CML)-expressing retrovirus or MSCV IRES GFP empty vector retrovirus. Subsequently, transduced cells were intravenously transplanted (2.5x10⁵ cells/mouse) into sublethally irradiated (900 cGy) recipient mice. Mice were sacrificed 14 days after transplantation.

Lineage depletion and transplantation

In order to transplant lineage-negative bone marrow cells, young (10-14-week-old) or old (52-75-week-old) donor mice were euthanized and skulls were harvested. The calvarium was first chopped with scissors in FACS buffer (PBS with 2% fetal calf serum), then crushed with a mortar and pestle. Cell suspension was filtered through a 40 μm mesh filter (Falcon, Cat# 352340), resuspended in RBC lysing buffer (Sigma-Aldrich, Cat# R7757) for red blood cell lysis and washed with FACS buffer. Cells were resuspended in FACS buffer and incubated with a biotinylated anti-hematopoietic lineage antibody cocktail (Miltenyi-Biotec, Cat# 130-092-613, 1:10 dilution), followed by washing with FACS buffer and incubation with R-PE-conjugated streptavidin secondary antibody (Invitrogen, Cat# S866, 1:50 dilution). DAPI (1:1000 dilution) was added to resuspended cells to distinguish live/dead cells and were FACS sorted for live lineage-negative cells on a FACSAria Fusion (BD Biosciences). Sorted cells were intravenously transplanted (5x10⁵ cells/mouse) into lethally irradiated (12 cGy, Best Theratronics, Gammacell 40 Exactor) recipient mice (12-week-old). Mice were sacrificed 14 days after transplantation.

FACS analysis of bone marrow and peripheral blood

Mice from each age group were euthanized and skull and femur were harvested. Skulls were chopped with scissors in FACS buffer before crushed with mortar and pestle; femurs were crushed without chopping. Cell suspensions were strained through a 40 μm mesh filter, resuspended in RBC lysing buffer and washed with FACS buffer. Cells were resuspended and incubated with the following primary antibodies in FACS buffer: biotinylated rat monoclonal anti-hematopoietic lineage antibody cocktail (Miltenyi-Biotgec, Cat# 130-092-613, 1:50 dilution), APC-conjugated rat monoclonal anti-CD117 (BD Biosciences, Cat# 553356, 1:100 dilution), FITC-conjugated rat monoclonal anti-Ly-6A/E (Sca-1) (eBioscience, Cat# 11-5981-85, 1:100 dilution), APC-Cy7-conjugated hamster monoclonal anti-CD48 (BD Biosciences, Cat# 561242, 1:100 dilution), and PE-conjugated rat monoclonal anti-CD150 (SLAM) (Biolegend, Cat# 115904, 1:100 dilution). Cells were washed, resuspended in FACS buffer with Alexa Fluor 405-conjugated streptavidin secondary antibody (Invitrogen, Cat# S32351, 1:100 dilution), washed again before analysis with a FACSymphony A5 Cell Analyzer (BD Biosciences).

Peripheral blood was collected from the submandibular vein with lancets (Medipoint, NY) into EDTA-coated tubes. Blood was resuspended in RBC lysing buffer and washed with FACS buffer. Cells were resuspended and incubated with the following primary antibodies in FACS buffer: biotinylated rat monoclonal anti-hematopoietic lineage antibody cocktail (Miltenyi-Biotgec, Cat# 130-092-613, 1:50 dilution), APC-conjugated rat monoclonal anti-CD117 (BD Biosciences, Cat# 553356, 1:100 dilution), FITC-conjugated rat monoclonal anti-Ly-6A/E (Sca-1) (eBioscience, Cat# 11-5981-85, 1:100 dilution), Pacific Blue-conjugated mouse monoclonal anti-CD45.2 (Biolegend, Cat# 109820, 1:100 dilution), APC-conjugated hamster monoclonal anti-CD3e (eBioscience, Cat# 17-0031, 1:100 dilution), PE-conjugated rat monoclonal anti-CD45R/B220 (BD Biosciences, Cat# 553090, 1:100 dilution), FITC-conjugated rat monoclonal anti-CD11b (BD Biosciences, Cat# 553310, 1:100 dilution). Cells were washed and resuspended in FACS buffer before analysis with a FACSymphony A5 Cell Analyzer (BD Biosciences).

RNA extraction and quantitative PCR

FACS-sorted cells from 10-week-old mouse skulls were lysed and RNA was extracted using a Monarch Total RNA Miniprep Kit (New England BioLabs, Cat# T2010S). Extracted RNA concentration was measured with a NanoDrop 8000 Spectrophotometer (Thermo Fischer Scientific) and cDNA was generated with a LuncaScript RT SuperMix Kit (New England BioLabs, Cat# E3010L). Quantitative PCR with reverse transcription was performed with a BioRad CFX96 real-time PCR system using FAM-conjugated Taqman probes for Vegfa (Mm00437306_m1) or using PowerUp SYBR Green Master Mix (Applied Biosystems, Cat# A25742) with primers designed using Pimer-BLAST or adopted from previously published studies: Vegfa₁₂₀ (5’-AACGATGAAGCCCTGGAGTG-3’; 5’-TGAGAGGTCTGGTTCCCGA-3’); Vegfa₁₆₄ (5’-AACGATGAAGCCCTGGAGTG; 5’-GACAAACAAATGCTTTCTCCG-3’); Vegfa₁₈₈ (5’- AACGATGAAGCCCTGGAGTG-3’; 5’-AACAAGGCTCACAGTGAACG-3’). Gene expression levels were normalized to the endogenous VIC-conjugated Gapdh probe (44326317E) as control.

ELISA

Hypoxia analysis

Hypoxic cells were detected with the hypoxia probe pimonidazole (Pimo, Hypoxyprobe Inc.) according to the manufacturer’s instructions. Mice were intraperitoneally injected with 60 mg kg^-1 1 hour before analysis.

VEGF-A plasmid construction and overexpression

To generate the pLIVE-VEGFA165-HA-MP-Asp_8x bone-homing protein containing VEGF165 fused to HA-tag, metalloprotease and 8x Asp peptide sequences, a cDNA fragment encoding amino acids 1-191 of human VEGFA was amplified via PCR using the following oligonucleotide primers: VEGFA-AscI-Fwd: 5'- ATGAACTTTCTGCTGTCT-3' and VEGFA-XhoI-Rev: 5'-CCGCCTCGGCTTGTCACATCTGCA-3' and annealed with the NEBuilder Assembly Cloning Kit.

10-week-old mice were used for hydrodynamic tail vein injection. Animals were injected with 0.5 μg g^-1 (plasmid/body weight) pLIVE-Vegfa plasmid suspended in TransIT-EE hydrodynamic delivery solution (Mirus, Cat# MIR5340). The appropriate amount of plasmid was suspended in an injection volume of 10% of the body weight and injected into each individual mouse via the tail vein in 5-7 s as previously reported⁸⁰.

Adipocyte analysis

To stain for neutral lipids, the entire calvarium or femur cryosections were incubated in BODIPY 493/503 (Invitrogen, Cat# D3922; 1:1000 dilution) for 1 hr at RT with gentle agitation (only calvarium). Samples were washed with PBS 3-5 times at 5 min intervals before mounting.

Analysis of inflammatory cytokines

Mice from each age group were euthanized, bones were harvested. Skulls were chopped before being crushed with a mortar and pestle in ice-cold RIPA lysis buffer; femurs were crushed without chopping. Supernatants of centrifuged lysates were further concentrated using an Ultra-0.5 Centrifugal Filter Unit with a 3 KDa cutoff (Millipore, Cat# UFC500396), resulting concentrations were measured using a Pierce BCA Protein Assay Kit (Thermo Fischer Scientific, Cat# 23225), and concentrations of inflammatory cyotokines were measured with LEGENDplex Mouse Inflammation Panel (13-plex) with V-bottom plates (Biolegend, Cat# 740446). Analysis on a FACSymphony (BD Biosciences) and quantification were performed according to the manufacturer’s protocol. Data analysis was performed using software provided by Biolegend. Manual gating was used to define beads A and B, while and automatic gating strategy was used to gate individual cytokines in the APC-PE plot.

Irradiation with partial shielding

Mice were anesthetized with ketamine (100 mg/kg) and xylazine (10 mg/kg) prior to irradiation. For partial shielding, the entire head or both legs of an animal were inserted into the opening of the cylindrical 1-inch-thick lead shield (JRT Associates, Cat# PTI-50-P) and exposed to lethal irradiation (12 cGy). The animal was returned to its cage to recover under observation.

Skull BM photoconversion

Vav1-KikGR mice were anesthetized with ketamine (100 mg/kg) and xylazine (10 mg/kg). A skin-flap was generated to expose the calvarium, as previously described⁸¹. Each exposed area of the calvarium was then exposed to UV light from a Zeiss Axio Imager (Zeiss Microscopy) for 60 seconds, confirmed for photoconversion from green to red fluorescence, before exposing another area. The skin flap was sutured back together and peripheral blood was analyzed by flow cytometry, as described above, to check for the presence of photoconverted cells, which were non-existent in the peripheral blood immediately after photoconversion. 1 week after photoconversion, peripheral blood was drawn, stained for Alexa Fluor-conjugated rat monoclonal anti-CD45, and was analyzed by flow cytometry for CD45⁺ photoconverted hematopoietic cells derived from the skull BM.

Statistical analysis

No statistical methods were used to predetermine sample size. The experiments were randomized and investigators were blinded to allocation during experiments and outcome analyses. All values are presented as mean ± standard deviation (SD). Statistical significance was determined by the two-tailed unpaired Student’s t test between two groups or the Tukey multiple comparison test (one-way ANOVA) for multiple-group comparison. Statistical analyses were performed using GraphPad Prism 9.0 (GraphPad Software). Statistical significance was set at P < 0.05.

Acknowledgments

We thank Martin Stehling for FACS technical assistance and Frank Berkenfeld for supporting animal colony maintenance. V.M. is a part of the Cells in Motion-International Max Planck Research School (CiM-IMPRS) and University of Münster, Germany. R.H.A. is supported by the Max Planck Society, the European Research Council (AdG 786672, PROVEC), the DFG (CRC 1366), and the Leducq Foundation.

Author Contributions

B.I.K. and R.H.A. designed the study and interpreted results. B.I.K. designed, conducted, and analyzed the majority of the experiments, including all pharmacological treatments, skull sample processing, immunostainings, transplantations, FACS sorting/analyses, RNA/protein analyses, and photoconversion surgeries. V.M. processed femur samples and provided technical assistance. H.-W.J. and H.P. provided technical assistance. Y.J.C. conducted and analyzed human patient head CT scans. M.N.-K. and P.V. designed and performed transient middle cerebral artery occlusion surgeries. R.S.P. and D.S.K. generated the chronic myeloid leukemia mouse model and contributed to project design. S.A. generated pLIVE plasmids. H.J.L. conducted scanning electron microscopy sample processing and imaging. M.G.B. contributed to project design and data interpretation. R.H.A. supervised the project.

Disclosures

The authors declare no competing interests.

Hoggatt, J., Kfoury, Y. & Scadden, D. T. Hematopoietic Stem Cell Niche in Health and Disease. Annual Review of Pathology: Mechanisms of Disease 11, 555-581, doi:10.1146/annurev-pathol-012615-044414 (2016).
Pinho, S. & Frenette, P. S. Haematopoietic stem cell activity and interactions with the niche. Nature Reviews Molecular Cell Biology 20, 303-320, doi:10.1038/s41580-019-0103-9 (2019).
Comazzetto, S., Shen, B. & Morrison, S. J. Niches that regulate stem cells and hematopoiesis in adult bone marrow. Developmental Cell 56, 1848-1860, doi:10.1016/j.devcel.2021.05.018 (2021).
van den Bos, T., Speijer, D., Bank, R. A., Bromme, D. & Everts, V. Differences in matrix composition between calvaria and long bone in mice suggest differences in biomechanical properties and resorption: Special emphasis on collagen. Bone 43, 459-468, doi:10.1016/j.bone.2008.05.009 (2008).
Sivaraj, K. K. & Adams, R. H. Blood vessel formation and function in bone. Development 143, 2706-2715, doi:10.1242/dev.136861 (2016).
Mills, W. A., Coburn, M. A. & Eyo, U. B. The emergence of the calvarial hematopoietic niche in health and disease. Immunological Reviews 311, 26-38, doi:10.1111/imr.13120 (2022).
Lo Celso, C. et al. Live-animal tracking of individual haematopoietic stem/progenitor cells in their niche. Nature 457, 92-96, doi:10.1038/nature07434 (2009).
Spencer, J. A. et al. Direct measurement of local oxygen concentration in the bone marrow of live animals. Nature 508, 269-273, doi:10.1038/nature13034 (2014).
Christodoulou, C. et al. Live-animal imaging of native haematopoietic stem and progenitor cells. Nature 578, 278-283, doi:10.1038/s41586-020-1971-z (2020).
Lassailly, F., Foster, K., Lopez-Onieva, L., Currie, E. & Bonnet, D. Multimodal imaging reveals structural and functional heterogeneity in different bone marrow compartments: functional implications on hematopoietic stem cells. Blood 122, 1730-1740, doi:10.1182/blood-2012-11-467498 (2013).
Herisson, F. et al. Direct vascular channels connect skull bone marrow and the brain surface enabling myeloid cell migration. Nature Neuroscience 21, 1209-1217, doi:10.1038/s41593-018-0213-2 (2018).
Brioschi, S. et al. Heterogeneity of meningeal B cells reveals a lymphopoietic niche at the CNS borders. Science 373, doi:10.1126/science.abf9277 (2021).
Cugurra, A. et al. Skull and vertebral bone marrow are myeloid cell reservoirs for the meninges and CNS parenchyma. Science 373, doi:10.1126/science.abf7844 (2021).
Mazzitelli, J. A. et al. Cerebrospinal fluid regulates skull bone marrow niches via direct access through dural channels. Nature Neuroscience 25, 555-560, doi:10.1038/s41593-022-01029-1 (2022).
Pulous, F. E. et al. Cerebrospinal fluid can exit into the skull bone marrow and instruct cranial hematopoiesis in mice with bacterial meningitis. Nature Neuroscience 25, 567-576, doi:10.1038/s41593-022-01060-2 (2022).
Young, K. et al. Progressive alterations in multipotent hematopoietic progenitors underlie lymphoid cell loss in aging. J Exp Med 213, 2259-2267, doi:10.1084/jem.20160168 (2016).
Verovskaya, E. V., Dellorusso, P. V. & Passegue, E. Losing Sense of Self and Surroundings: Hematopoietic Stem Cell Aging and Leukemic Transformation. Trends Mol Med 25, 494-515, doi:10.1016/j.molmed.2019.04.006 (2019).
Nakamura-Ishizu, A., Ito, K. & Suda, T. Hematopoietic Stem Cell Metabolism during Development and Aging. Developmental Cell 54, 239-255, doi:10.1016/j.devcel.2020.06.029 (2020).
Kovtonyuk, L. V., Fritsch, K., Feng, X., Manz, M. G. & Takizawa, H. Inflamm-Aging of Hematopoiesis, Hematopoietic Stem Cells, and the Bone Marrow Microenvironment. Front Immunol 7, 502, doi:10.3389/fimmu.2016.00502 (2016).
Potente, M., Gerhardt, H. & Carmeliet, P. Basic and therapeutic aspects of angiogenesis. Cell 146, 873-887, doi:10.1016/j.cell.2011.08.039 (2011).
Chung, A. S. & Ferrara, N. Developmental and pathological angiogenesis. Annu Rev Cell Dev Biol 27, 563-584, doi:10.1146/annurev-cellbio-092910-154002 (2011).
Ramasamy, S. K. et al. Regulation of Hematopoiesis and Osteogenesis by Blood Vessel–Derived Signals. Annual Review of Cell and Developmental Biology 32, 649-675, doi:10.1146/annurev-cellbio-111315-124936 (2016).
Ramalingam, P., Butler, J. M. & Poulos, M. G. Vascular Regulation of Hematopoietic Stem Cell Homeostasis, Regeneration, and Aging. Current Stem Cell Reports 7, 194-203, doi:10.1007/s40778-021-00198-2 (2021).
Xu, Y. et al. Neuropilin-2 mediates VEGF-C–induced lymphatic sprouting together with VEGFR3. Journal of Cell Biology 188, 115-130, doi:10.1083/jcb.200903137 (2010).
Kusumbe, A. P., Ramasamy, S. K. & Adams, R. H. Coupling of angiogenesis and osteogenesis by a specific vessel subtype in bone. Nature 507, 323-328, doi:10.1038/nature13145 (2014).
Kusumbe, A. P. et al. Age-dependent modulation of vascular niches for haematopoietic stem cells. Nature 532, 380-384, doi:10.1038/nature17638 (2016).
Langen, U. H. et al. Cell–matrix signals specify bone endothelial cells during developmental osteogenesis. Nature Cell Biology 19, 189-201, doi:10.1038/ncb3476 (2017).
Yu, J. Direct evidence for the role of caveolin-1 and caveolae in mechanotransduction and remodeling of blood vessels. Journal of Clinical Investigation 116, 1284-1291, doi:10.1172/jci27100 (2006).
Heil, J. et al. Bone marrow sinusoidal endothelium controls terminal erythroid differentiation and reticulocyte maturation. Nature Communications 12, doi:10.1038/s41467-021-27161-3 (2021).
de Boer, J. et al. Transgenic mice with hematopoietic and lymphoid specific expression of Cre. Eur J Immunol 33, 314-325, doi:10.1002/immu.200310005 (2003).
Muzumdar, M. D., Tasic, B., Miyamichi, K., Li, L. & Luo, L. A global double-fluorescent Cre reporter mouse. Genesis 45, 593-605, doi:10.1002/dvg.20335 (2007).
Mitchell, C. A. et al. Stromal niche inflammation mediated by IL-1 signalling is a targetable driver of haematopoietic ageing. Nature Cell Biology 25, 30-41, doi:10.1038/s41556-022-01053-0 (2023).
Cai, R. et al. Panoptic imaging of transparent mice reveals whole-body neuronal projections and skull–meninges connections. Nature Neuroscience 22, 317-327, doi:10.1038/s41593-018-0301-3 (2019).
Olson, O. C., Kang, Y. A. & Passegue, E. Normal Hematopoiesis Is a Balancing Act of Self-Renewal and Regeneration. Cold Spring Harb Perspect Med 10, doi:10.1101/cshperspect.a035519 (2020).
Janssen, H., Koekkoek, L. L. & Swirski, F. K. Effects of lifestyle factors on leukocytes in cardiovascular health and disease. Nat Rev Cardiol, doi:10.1038/s41569-023-00931-w (2023).
Nakada, D. et al. Oestrogen increases haematopoietic stem-cell self-renewal in females and during pregnancy. Nature 505, 555-558, doi:10.1038/nature12932 (2014).
Oguro, H. et al. 27-Hydroxycholesterol induces hematopoietic stem cell mobilization and extramedullary hematopoiesis during pregnancy. J Clin Invest 127, 3392-3401, doi:10.1172/JCI94027 (2017).
Rodrigues, J. et al. Estrogen enforces the integrity of blood vessels in the bone during pregnancy and menopause. Nature Cardiovascular Research 1, 918-932, doi:10.1038/s44161-022-00139-0 (2022).
Courties, G. et al. Ischemic stroke activates hematopoietic bone marrow stem cells. Circ Res 116, 407-417, doi:10.1161/CIRCRESAHA.116.305207 (2015).
Mendez-Ferrer, S. et al. Bone marrow niches in haematological malignancies. Nat Rev Cancer 20, 285-298, doi:10.1038/s41568-020-0245-2 (2020).
Reid, I. R. & Billington, E. O. Drug therapy for osteoporosis in older adults. Lancet 399, 1080-1092, doi:10.1016/S0140-6736(21)02646-5 (2022).
Calvi, L. M. et al. Osteoblastic cells regulate the haematopoietic stem cell niche. Nature 425, 841-846, doi:10.1038/nature02040 (2003).
Beerman, I., Maloney, W. J., Weissmann, I. L. & Rossi, D. J. Stem cells and the aging hematopoietic system. Current Opinion in Immunology 22, 500-506, doi:10.1016/j.coi.2010.06.007 (2010).
Akunuru, S. & Geiger, H. Aging, Clonality, and Rejuvenation of Hematopoietic Stem Cells. Trends in Molecular Medicine 22, 701-712, doi:10.1016/j.molmed.2016.06.003 (2016).
Morrison, S. J., Wandycz, A. M., Akashi, K., Globerson, A. & Weissman, I. L. The aging of hematopoietic stem cells. Nat Med 2, 1011-1016, doi:10.1038/nm0996-1011 (1996).
Rossi, D. J. et al. Cell intrinsic alterations underlie hematopoietic stem cell aging. Proc Natl Acad Sci U S A 102, 9194-9199, doi:10.1073/pnas.0503280102 (2005).
Colom Diaz, P. A., Mistry, J. J. & Trowbridge, J. J. Hematopoietic stem cell aging and leukemia transformation. Blood 142, 533-542, doi:10.1182/blood.2022017933 (2023).
North, T. E. et al. Prostaglandin E2 regulates vertebrate haematopoietic stem cell homeostasis. Nature 447, 1007-1011, doi:10.1038/nature05883 (2007).
Hoggatt, J., Singh, P., Sampath, J. & Pelus, L. M. Prostaglandin E2 enhances hematopoietic stem cell homing, survival, and proliferation. Blood 113, 5444-5455, doi:10.1182/blood-2009-01-201335 (2009).
Porter, R. L. et al. Prostaglandin E2 Increases Hematopoietic Stem Cell Survival and Accelerates Hematopoietic Recovery After Radiation Injury. Stem Cells 31, 372-383, doi:10.1002/stem.1286 (2013).
Petit, I. et al. G-CSF induces stem cell mobilization by decreasing bone marrow SDF-1 and up-regulating CXCR4. Nature Immunology 3, 687-694, doi:10.1038/ni813 (2002).
Bendall, L. J. & Bradstock, K. F. G-CSF: From granulopoietic stimulant to bone marrow stem cell mobilizing agent. Cytokine Growth Factor Rev 25, 355-367, doi:10.1016/j.cytogfr.2014.07.011 (2014).
Simons, M., Gordon, E. & Claesson-Welsh, L. Mechanisms and regulation of endothelial VEGF receptor signalling. Nature Reviews Molecular Cell Biology 17, 611-625, doi:10.1038/nrm.2016.87 (2016).
Apte, R. S., Chen, D. S. & Ferrara, N. VEGF in Signaling and Disease: Beyond Discovery and Development. Cell 176, 1248-1264, doi:10.1016/j.cell.2019.01.021 (2019).
Chen, Q. et al. Apelin(+) Endothelial Niche Cells Control Hematopoiesis and Mediate Vascular Regeneration after Myeloablative Injury. Cell Stem Cell 25, 768-783 e766, doi:10.1016/j.stem.2019.10.006 (2019).
Schipani, E., Wu, C., Rankin, E. B. & Giaccia, A. J. Regulation of Bone Marrow Angiogenesis by Osteoblasts during Bone Development and Homeostasis. Front Endocrinol (Lausanne) 4, 85, doi:10.3389/fendo.2013.00085 (2013).
Wong, B. W., Marsch, E., Treps, L., Baes, M. & Carmeliet, P. Endothelial cell metabolism in health and disease: impact of hypoxia. The EMBO Journal 36, 2187-2203, doi:10.15252/embj.201696150 (2017).
Wicks, E. E. & Semenza, G. L. Hypoxia-inducible factors: cancer progression and clinical translation. Journal of Clinical Investigation 132, doi:10.1172/jci159839 (2022).
Suda, T. & Liu, D. Hydrodynamic gene delivery: its principles and applications. Mol Ther 15, 2063-2069, doi:10.1038/sj.mt.6300314 (2007).
Ho, Y.-H. & Méndez-Ferrer, S. Microenvironmental contributions to hematopoietic stem cell aging. Haematologica 105, 38-46, doi:10.3324/haematol.2018.211334 (2020).
Ambrosi, T. H. et al. Adipocyte Accumulation in the Bone Marrow during Obesity and Aging Impairs Stem Cell-Based Hematopoietic and Bone Regeneration. Cell Stem Cell 20, 771-784.e776, doi:10.1016/j.stem.2017.02.009 (2017).
Zioni, N. et al. Inflammatory signals from fatty bone marrow support DNMT3A driven clonal hematopoiesis. Nature Communications 14, doi:10.1038/s41467-023-36906-1 (2023).
Pang, W. W. et al. Human bone marrow hematopoietic stem cells are increased in frequency and myeloid-biased with age. Proceedings of the National Academy of Sciences 108, 20012-20017, doi:10.1073/pnas.1116110108 (2011).
Ho, Y. H. et al. Remodeling of Bone Marrow Hematopoietic Stem Cell Niches Promotes Myeloid Cell Expansion during Premature or Physiological Aging. Cell Stem Cell 25, 407-418 e406, doi:10.1016/j.stem.2019.06.007 (2019).
Tomura, M. et al. Tracking and quantification of dendritic cell migration and antigen trafficking between the skin and lymph nodes. Sci Rep 4, 6030, doi:10.1038/srep06030 (2014).
Blumer, M. J., Longato, S. & Fritsch, H. Structure, formation and role of cartilage canals in the developing bone. Ann Anat 190, 305-315, doi:10.1016/j.aanat.2008.02.004 (2008).
Maes, C. Role and regulation of vascularization processes in endochondral bones. Calcif Tissue Int 92, 307-323, doi:10.1007/s00223-012-9689-z (2013).
Carmeliet, P. & Jain, R. K. Molecular mechanisms and clinical applications of angiogenesis. Nature 473, 298-307, doi:10.1038/nature10144 (2011).
Hu, K. & Olsen, B. R. Osteoblast-derived VEGF regulates osteoblast differentiation and bone formation during bone repair. J Clin Invest 126, 509-526, doi:10.1172/JCI82585 (2016).
MacLean, H. E. & Kronenberg, H. M. Expression of Stra13 during mouse endochondral bone development. Gene Expr Patterns 4, 633-636, doi:10.1016/j.modgep.2004.04.011 (2004).
Grunewald, M. et al. Counteracting age-related VEGF signaling insufficiency promotes healthy aging and extends life span. Science 373, doi:10.1126/science.abc8479 (2021).
Ding, L., Saunders, T. L., Enikolopov, G. & Morrison, S. J. Endothelial and perivascular cells maintain haematopoietic stem cells. Nature 481, 457-462, doi:10.1038/nature10783 (2012).
Greenbaum, A. et al. CXCL12 in early mesenchymal progenitors is required for haematopoietic stem-cell maintenance. Nature 495, 227-230, doi:10.1038/nature11926 (2013).
Heidt, T. et al. Chronic variable stress activates hematopoietic stem cells. Nat Med 20, 754-758, doi:10.1038/nm.3589 (2014).
Letsky, E. A. Erythropoiesis in pregnancy. J Perinat Med 23, 39-45, doi:10.1515/jpme.1995.23.1-2.39 (1995).
Kolabas, Z. I. et al. Distinct molecular profiles of skull bone marrow in health and neurological disorders. Cell 186, 3706-3725 e3729, doi:10.1016/j.cell.2023.07.009 (2023).
Sabanciogullari, V. et al. Diploe thickness and cranial dimensions in males and females in mid-Anatolian population: an MRI study. Forensic Sci Int 219, 289 e281-287, doi:10.1016/j.forsciint.2011.11.033 (2012).
Lillie, E. M., Urban, J. E., Lynch, S. K., Weaver, A. A. & Stitzel, J. D. Evaluation of Skull Cortical Thickness Changes With Age and Sex From Computed Tomography Scans. J Bone Miner Res 31, 299-307, doi:10.1002/jbmr.2613 (2016).
Krause, D. S. et al. Differential regulation of myeloid leukemias by the bone marrow microenvironment. Nat Med 19, 1513-1517, doi:10.1038/nm.3364 (2013).
Liu, F., Song, Y. & Liu, D. Hydrodynamics-based transfection in animals by systemic administration of plasmid DNA. Gene Ther 6, 1258-1266, doi:10.1038/sj.gt.3300947 (1999).
Koh, B. I. et al. VEGFR2 signaling drives meningeal vascular regeneration upon head injury. Nat Commun 11, 3866, doi:10.1038/s41467-020-17545-2 (2020).

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Adult skull bone marrow is an expanding and resilient hematopoietic reservoir

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