Hypoxia-inducible factor 1 a (Hif1α) plays a crucial role in brain development. To study the function of Hif1α in early brain development, we generated neuroepithelial cell-specific Hif1α-knockout mice. Hif1α-knockout mice died soon after birth; these mice exhibited an abnormal head shape, indicating the presence of brain defects. Morphological analysis revealed that Hif1α ablation reduced the overall size of the brain, especially affecting the telencephalon. Neuronal apoptosis predominantly occurred in deep-layer neurons, consequently the alignment of cortical layers was severely disorganized in Hif1α knockout mice. Furthermore, we demonstrated that Vegf signaling contributes to the survival of deep-layer neurons as a downstream effector of Hif1α-dependent hypoxia signaling. Taken together, our findings demonstrate that Hif1α plays a critical role in the early stages of telencephalon development.
Research
Hif1α-Dependent Hypoxia Signaling Contributes to the Survival of Deep-Layer Neurons and Cortex Formation in A Mouse Model
https://doi.org/10.21203/rs.3.rs-942004/v1
This work is licensed under a CC BY 4.0 License
Journal Publication
published 31 Mar, 2022
You are reading this latest preprint version
In placental mammals, the embryo encounters drastic changes in the environmental oxygen concentration during development. While the oxygen concentration is relatively high in the oviduct (approximately 8%), it is lower in the uterus (approximately 3–5%). Post-implantation, embryos encounter more anaerobic conditions while they are embedded into the endometrium, and this constraint persists until the onset of oxygen supply from maternal blood [1–4]. Even after the supply of oxygen, embryonic tissues, such as the neural tube, heart primordia, and intersomitic mesenchyme, continue to experience hypoxia [5, 6]. Proper development of mammalian embryos requires their adaptation to drastic changes in oxygen concentration. Therefore, mammalian embryos employ unique molecular pathways to enable gastrulation and organogenesis within a transient hypoxic environment. Indeed, it has been reported that culturing rodent embryos exo utero under hyperoxic conditions leads to abnormal embryo morphology [7].
Hypoxia-inducible factor 1 α (Hif1α), a key transcription factor that forms a complex with Hif1β, is involved in the cellular response to anaerobic conditions [8–11]. The stability of the Hif1α–Hif1β complex is regulated by the enzyme prolyl hydroxylase 1–3 (PHD1-3) in an oxygen-dependent manner. Under anaerobic conditions, the Hif1α–Hif1β complex is not hydroxylated, and therefore, is not targeted for degradation. The stabilized Hif1α–Hif1β complex binds to the hypoxia response element (HRE) to activate the expression of genes involved in energy metabolism, erythropoiesis, and angiogenesis, thereby protecting cells and tissues from hypoxic stress [12, 13]. In mammalian embryos, a Hif1α-dependent hypoxic response is required not only for the prevention of damage caused by hypoxic stress but also for proper progression of embryonic development. Hif1α-deficient mouse embryos exhibit defects in cardiovascular formation, somitogenesis, and neural tube closure, resulting in developmental arrest and lethality by embryonic day (E) 11 [14–17]. Furthermore, studies have shown that Hif1α is required for the formation of the heart, cartilage, and limbs, using conditional Hif1α−knockout mice [18–20]. Regarding the central nervous system (CNS), Christian et al. demonstrated that the ablation of Hif1α using the Emx1-Cre driver causes apoptosis of neurons and cortical hypoplasia in the cerebral cortex [21]. However, the function of Hif1α in cortical development has not been examined in detail. Hif1α-knockout mice have been generated using the Nestin-Cre driver to observe a more broad-spectrum effect on neural development. Studies using Nestin-Cre driver have revealed that the ablation of Hif1α disrupts angiogenesis and consequently, expands a hypoxic region in the telencephalon at E16 [22]. Neural cell-specific ablation did not affect brain development until E15, but caused massive neuronal apoptosis in the telencephalon at E19, resulting in hydrocephalus at postnatal day (P) 70 [23]. Although Nestin-Cre mice express Cre mRNA throughout the CNS at E11.5, it is uncertain whether Cre is expressed before E11.5 [24]. In addition, Cre recombination activity is not detected in the ventricular zone (VZ) and subventricular zone (SVZ) at E12.5 and E14.5 in the commercially available Nestin-Cre line. Therefore, the Nestin-Cre line may be inefficient as a Cre-driver for recombination in embryonic neural progenitor cells [25]. Therefore, novel functions of Hif1α in early brain development could be discovered using other Cre-driver mice that express Cre within embryonic neural progenitor cells before E11.5. Herein, we investigated the function of Hif1α by neuroepithelial cell-specific ablation of Hif1α using Sox1-Cre [26]. In the Sox1-Cre driver, Cre activity was observed in neuroepithelial cells at least from E8.5; thus, the crossing of Hif1α-floxed mice with Sox1-Cre mice allow us to analyze the function of Hif1α in early neural development. In the present study, we demonstrated that Hif1α is required for neuronal survival, thereby facilitating the formation of cortical layers during telencephalon development.
Generation of neuroepithelial cell-specific Hif1α-knockout mice
To address the biological significance of hypoxia in early brain development, we focused on deleting Hif1α, a master regulator of cellular response to hypoxia [8–11]. We crossed Hif1α flox/flox mice [16] with Sox1-Cre+/− mice [26] to selectively ablate Hif1α in early neural progenitor cells, thereby affecting all CNS cells. The resulting Hif1α gene carrying an exon 2 deletion encoded a malfunctioning Hif1α protein that lost the ability to induce the expression of genes containing HRE [16]. Thus, we investigated the function of Hif1α-dependent hypoxic response in neural progenitor cells and their progeny during brain development. Heterozygotes (Hif1α flox/wt; Sox1-Cre+/−) were viable and fertile, indicating that ablation of a single Hif1α allele or the expression of Cre did not affect CNS development. In contrast, homozygotes (Hif1α flox/flox; Sox1-Cre+/−) conditional mutants died within several hours after birth. Mice homozygous for the floxed Hif1α allele with a Sox1-Cre allele (Hif1α flox/flox; Sox1-Cre+/−) were used to generate conditional knockout mice, which are hereafter referred to as “KO” mice. Mice homozygous for the floxed Hif1α allele without a Sox1-Cre allele (Hif1α flox/flox; Sox1-Cre−/−) were used as controls and are hereafter referred to as “WT” mice.
We confirmed the deletion of Hif1α using in situ hybridization with an antisense probe against exon 2. At E8.5, Hif1α mRNA was predominantly expressed in neuroepithelial cells and mesenchymal cells of WT embryos. The expression patterns and levels of Hif1α mRNA in KO embryos were comparable with those in WT embryos (Fig. 1A and B). At E9.5, Hif1α continued to be expressed in neuroepithelial cells and mesenchymal cells in WT embryos (Fig. 1C). In contrast, Hif1α expression diminished in the neural tube of KO embryos, although it remained unchanged in mesenchymal cells at E9.5 (Fig. 1D). To further validate the ablation of Hif1α, the expression of Hif1α and its target genes, Ldha, Aldoa (encoding a glycolytic enzyme) and Epo (encoding erythropoietin), was analyzed using RT-qPCR. Hif1α expression was slightly decreased by loss of Hif1α, while Hif1α target genes were expressed in KO embryos at levels similar to those in WT embryos at E8.5 (Fig. 1E). However, Hif1α expression decreased by approximately 60% in KO embryos relative to that in WT embryos at E9.5 (Fig. 1F), which was consistent with the results of in situ hybridization showing the near-complete loss of Hif1α transcripts in the neuroepithelium (Fig. 1D). Accordingly, the expression of Hif1α target genes was downregulated in E9.5 KO embryos (Fig. 1F; Ldha, reduced by 62%; Aldoa, by 77%; and Epo, by 59%). We isolated total RNA from whole mouse embryos and then prepared cDNA for RT-qPCR. Although it is preferred that total RNA is isolated exclusively from the neuroepithelium, this is technically challenging in early-stage embryos. Therefore, the reduced expression of Hif1α and Hif1α target genes would have been more severe if cDNA prepared from isolated neuroepithelium was used. As controls, we evaluated the expression of EpoR (encoding erythropoietin receptor) and Nqo1 (encoding electron respiratory chain enzyme), which are not dependent on Hif1α. The results showed that their expression was not altered in KO embryos at E9.5 (Fig. 1F); consequently, it was inferred that the decrease in Ldha, Aldoa, and Epo expression levels was not due to a deficit in respiratory metabolism and erythropoiesis, but instead reflected the loss of Hif1α-dependent gene regulation in the early neuroepithelium. Taken together, these results indicate that Hif1α was specifically ablated in neuroepithelial cells as early as E9.5 in KO embryos.
Hif1α ablation causes severe brain defects
Next, we examined the gross morphology of embryos and pups. Although KO embryos were indistinguishable from WT embryos in appearance until E12.5, they exhibited an abnormal-shaped head at E14.5. The length of the head along the sagittal axis was substantially reduced in KO embryos compared to that in WT embryos at E14.5 (Fig. 2A and B, curved two direction arrow). At E16.5, the parietal region of the head was flattened in KO embryos (Fig. 2D, arrowhead), which became more evident by P0 (Fig. 2F, arrowhead). Head abnormalities typically indicate brain hypoplasia; thus, we next examined the morphology of the brain. The brain of KO embryos was smaller than that of their WT counterparts, particularly the telencephalon at E14.5 and E16.5 (Fig. 2G-J). The brain of P0 KO pups exhibited considerable hypoplasia compared to that of WTs. Notably, the cerebrum of KO pups was extremely small compared to that of WT littermates (Fig. 2K and L). Collectively, these results demonstrate the requirement of Hif1α for normal development of the brain, especially that of the cerebrum.
Neurons undergo massive apoptosis in Hif1α-ablated embryos
Because the telencephalon was most acutely affected by the loss of Hif1α, we investigated the role of Hif1α-dependent hypoxia signaling in telencephalic development. The ablation of Hif1α has been reported to induce apoptosis of neuronal cells in the telencephalon [21, 23]. Thus, we examined apoptosis using immunofluorescent labeling of cleaved caspase 3, a marker of apoptotic cells. Some apoptotic cells were detected in the dorsal telencephalon of KO embryos at E12.5 (Fig. 3B). By E14.5, KO embryos displayed enhanced levels of apoptotic cells in the pallium (dorsal telencephalon) and the subpallium (ventral telencephalon) (Fig. 3D). The number of apoptotic cells in KO embryos was further increased in both the pallium and subpallium by E16.5 and E18.5 (Fig. 3F and H).
Cerebral cortex layers are disorganized by the loss of Hif1α
We further analyzed apoptosis in the KO cortex at P0. A large proportion of apoptotic cells resided in the Tuj1+ neuronal layer of the cortex, and few apoptotic cells were detected in the intermediate, subventricular, and VZ (Fig. 4B). This observation suggests that Hif1α loss led to the preferential elimination of postmitotic neurons, and consequently led to the reduction of the size of the cerebrum. We counted the number of Satb2+ and Ctip2+ cortical neurons in four arbitrarily defined regions (100 µm-wide) in the P0 cortex. Satb2 is intensely expressed in upper-layer neurons (layers II–IV), whereas Ctip2 is mainly expressed in deep-layer neurons (layers V and VI) in the cortex. The number of Satb2+ and Ctip2+ cells in the P0 cortex was reduced by approximately 87% and 53%, respectively, compared to WT controls (Table 1). This indicated that deep-layer neurons are more sensitive to Hif1α ablation than upper-layer neurons. Remarkably, apoptotic cells formed aggregates in the cortex (Fig. 4B), implying a disruption of the cortical layer formation in the KO cortex. The cortex consists of molecularly distinct Satb2+ upper layer and Ctip2+ deep layer projection neurons in the cerebrum of WT embryos (Fig. 4C). In contrast, in the KO cortex each cortical layer was obscured, and Satb2+ and Ctip2+ cells were broadly scattered throughout the cortex (Fig. 4D).The disorganized cortical layers was reminiscent of the phenotype of Reelin-deficient mice, reeler [27]. Cajal-Retzius (C-R) cells, which occur in the most superficial marginal zone layer of the cerebral cortex, regulate cortical layer formation through the secretion of Reelin [28, 29]. Therefore, the Sox1-Cre-mediated deletion of Hif1αduring early telencephalic development may have affected the survival of C-R cells. However, C-R cells were detected in the most superficial layer (layer I) of the cortex, underneath the pia, in both WT and KO telencephalons (Fig. 4E and F), suggesting that the Reelin-mediated guidance of postmitotic neurons is intact in the Hif1αKO cortex.
Neuronal migration is impaired in Hif1α-ablated cortex
To explore the mechanism inducing neuronal apoptosis by loss of Hif1α, we examined whether upper- or deep-layer neurons undergo apoptosis via immunofluorescent staining using anti-cleaved caspase3 and anti-Satb2 or anti-Ctip2 antibodies (Fig. 5A-D). We observed that many Ctip2+ cells were co-labeled with cleaved caspase3 (Fig. 5D, inset). The number of Satb2+/cleaved caspase3+ double-labelled cells was lower than that of Ctip2+/cleaved caspase3+ cells in the E18.5 KO cortex (Fig. 5B, inset). Specifically, approximately 8.4% and 35.1% of total cleaved caspase3+ cells were co-labeled with Satb2 and Ctip2, respectively (Fig. 5E). This demonstrated that deep-layer neurons preferentially undergo apoptosis in the absence of Hif1α, which was consistent with data showing a predominant reduction of deep-layer neurons in the KO cortex (Table 1). Approximately 56% of all cells were cleaved caspase3+ cells, thereby explaining the severe loss of layer marker expression.
Many Satb2+ upper-layer neurons were detected around apoptotic cell aggregates, especially on the apical side of the KO cortex (Fig. 5A, yellow arrowheads). Consistent with this, Satb2+ cells were sparsely distributed on the pial surface side of apoptotic cell aggregates. Conversely, Ctip2+ neurons were abnormally detected near the pial surface side where the upper layers were formed in the normal cortex (Fig. 5C, white arrowheads). The cerebral cortex consists of six neuronal layers that develop in an inside-out manner, i.e., early-born neurons settle in the deep layers, whereas late-born neurons migrate through the deep-layer neurons and form more superficial layers (upper layers) [30]. Thus, we hypothesized that the migration of the upper-layer neurons from the VZ to the pial surface was impeded by apoptosis of deep-layer neurons, resulting in a partial reversal of the cortical layer in the KO cortex. To test this hypothesis, we performed a BrdU birthdating assay. To label the upper-layer neurons, BrdU was injected into pregnant mice at E14.5, when upper layer neurons are being born, and then the position of BrdU-labeled neurons were compared between KO and WT cortices at E18.5. A large proportion of BrdU+ neurons were located in the upper layer of the WT cortex (corresponding to bins 1 and 2; Fig. 6A and C). In contrast, many BrdU+ neurons were detected in the deeper layers of the KO cortex. Importantly, most BrdU+ neurons were negative for apoptosis markers (Fig. 6B and D), which is consistent with the results shown in Fig. 5B and E. The distribution of BrdU+ neurons born at E14.5 was quantified in five arbitrarily defined regions (500 µm-wide) in the cortex. In the WT cortex, approximately 72% of BrdU+ neurons were present in the upper layers (bins 1 and 2). In contrast, the percentage of BrdU+ neurons in the presumptive area of the upper layers (bins 1 and 2) was reduced to approximately 36% in the KO cortex (Fig. 6G). Approximately 42% of BrdU+ neurons populated the middle area (bins 3 and 4) in the KO cortex, where a large number of apoptotic cells were observed. In contrast, approximately 21% of BrdU+ neurons were detected in the same region of the WT cortex (Fig. 6G). Notably, the total number of BrdU+ neurons in the KO cortex was comparable with that of BrdU+ neurons in the WT cortex, suggesting that the proliferation of neural progenitors of upper-layer neurons was not altered by the loss of Hif1α (Fig. 6H). Taken together, these results strongly support our hypothesis that the migration of upper-layer neurons was impeded by massive apoptosis of deep-layer neurons in the KO cortex.
Possible mechanism underlying the role of Hif1α in cortical development
Previous studies demonstrated that vascular endothelial growth factor (encoded by Vegf), a downstream target of Hif1α, functions not only as an angiogenic growth factor but also as a neurotrophic factor [31, 32]. It has also been demonstrated that neuronal cell-specific Vegf homozygous knockout mice die within 24 h after birth due to the dysmorphogenesis of the cortex caused by massive neuronal apoptosis [33, 34]. Moreover, as apoptotic cells seem to accumulate in the middle area of the cortex [33], the Vegf homozygous knockout phenotype resembles that of our Hif1α homozygous knockout mice. We therefore investigated if the reduced Vegf expression was associated with reduced survival of deep-layer neurons upon Hif1α loss. Vegf is expressed in the ventral layer of the brain at E14.5 [35]. First, we analyzed Vegf expression levels in WT and KO telencephalons at E13.5, when deep-layer neurons are born. To analyze this, the dorsal part of the telencephalon, including the cortex, was surgically isolated, and RNA was prepared for RT-qPCR. We confirmed a significant reduction in Vegf expression in the cortex of KO mice (Fig. 7A). Next, we examined whether Vegf signaling activity is required for the survival of deep-layer neurons. Although the suppression of Vegf receptor function is the ideal method to inhibit the input of Vegf signaling, known receptors for Vegf, such as Flk1, Flt1, Nrp1, and Nrp2, do not appear to be involved in cortical development and thus unlikely targets to manipulate Vegf signaling in the developing brain [34, 36–38]. Therefore, we took advantage of the ligand-binding domain of the Vegf receptor, sFlt1, to interfere with Vegf signaling [39]. It has been reported that the overexpression of sFlt1 inhibit Vegf signaling in vivo [40]. We hypothesized that secreted sFlt1 functions as a decoy Vegf receptor and inhibits paracrine and/or autocrine Vegf signaling inputs to neural progenitors and/or neurons of deep layers. The expression construct of mouse sFlt1 was introduced into neural progenitor cells of E13.5 WT embryos using in utero electroporation along with the EGFP expression construct. At E18.5, numerous EGFP+ sFlt1-overexpressing cells were co-labeled with apoptotic markers, while no apoptosis was observed in the control cortex (Fig. 7B and D). However, we could not detect apoptotic cell aggregates, which may be due to the variation in transfection efficiency of sFlt1 (Fig. 7D and E). In contrast, apoptosis was not induced when the sFlt1-expressing construct was electroporated into neural progenitor cells at E15.5 (Supplementary Fig. 1), suggesting that Vegf signaling is not required for the survival of upper-layer neurons. Next, we investigated cortical layer formation in the sFlt1-transfected cortex. Satb2+ neurons were dispersed in the apical side of the cortex where deep-layer neurons normally settle in the control cortex (Fig. 7F, G, J, K). The Ctip2+ neuron layer was considerably shifted to the pial surface in the transfected region of sFlt1, consistent with the accumulation of upper-layer neurons in the deep layers of the KO cortex (Fig. 7H, I, L, M). The overexpression of sFlt1 in neural progenitors and neurons of deep layers mimics cortical development defects seen in KO embryos.
In the present study, we elucidated the role of Hif1α in early-stage telencephalic development. We observed that neuroepithelial cell-specific ablation of Hif1α causes massive apoptosis of neurons in the telencephalon and cerebral cortex, which manifests as reduced brain size. We demonstrated that deep-layer neurons predominantly undergo apoptosis by Hif1α-ablation, and their loss affects the radial migration of upper-layer neurons and subsequent cortical layer formation. Furthermore, we showed that Vegf acts as a neurotrophic factor likely acting downstream of Hif1α to promote the survival of deep-layer neurons (Fig. 8).
Neural cell-specific knockout of Hif1α using Nestin-Cre driver causes massive apoptosis in the cerebrum at E19, resulting in hydrocephalus at P70, while no gross morphological differences can be observed in E15 embryos [23]. In our KO embryos, apoptosis occurred as early as E12.5, and gross morphological abnormalities of the head were apparent at E14.5 (Fig. 2). These phenotypic differences are presumably the result of the timing of Cre-mediated recombination in the Nestin-Cre vs. Sox1-Cre driver alleles. In the Nestin-Cre mouse line, Cre mRNA expression is detected by E11.5; however, it is unclear if Cre is expressed at earlier stages [24]. In contrast, Cre-mediated recombination was evident as early as E9.5 in our Sox1-Cre-mediated Hif1α KO embryos (Fig. 1D and F). Liang et al. indicated that Cre-mediated recombination occurs at sufficient levels in postmitotic neuronal cells, and not only in neural progenitors in Nestin-Cre embryos [25]. Thus, by restricting the recombination to early neuroepithelial cells using the Sox1-Cre driver, we produced a more complete neural-specific ablation of Hif1α and revealed a specific function for the gene in early telencephalic development. Sox1 mRNA expression begins at the anterior neuroepithelium of wild-type mouse embryos at E8.0 [41]. Consistent with this, Cre recombination activity has been detected in the neuroepithelium of Sox1-Cre; ROSA26R-EYFP, at least E8.5 onward [26]. However, Cre-mediated recombination was not detected at E8.5 in Sox1-Cre-mediated Hif1α KO embryos using in situ hybridization. The delay in Cre activity may be related to the difference in recombination efficiency of the ROSA26R allele compared to that of the Hif1α flox/flox allele.
Our findings strongly suggest that massive apoptosis of deep-layer neurons is the major cause of cortical layer formation deficiency in the KO cortex. However, the mechanism by which apoptotic elimination disturbs the migration of upper-layer neurons is still elusive. One possibility is the destruction of the chemoattractant gradient. For example, Semaphorin-3A (Sema3A) mRNA is highly expressed in the outer cortical layers of the rat cortex from E14 (nearly equal to E12 mouse cortex), and Sema3A protein is expressed across the rat cortex in a graded manner from E18 (equivalent to the E16 mouse cortex). The endogenous gradient of Sema3A functions as a guidance factor and regulates the migration of upper-layer neurons [42]. Apoptotic elimination of deep-layer neurons may destroy the Sema3A gradient in the cortex; consequently, upper-layer neurons may not be able to pass through the deep layers. Another possibility is that apoptotic cell aggregates may physically block the migration of upper-layer neurons in the KO brains. The molecular mechanism of neuronal migration defects caused by the loss of Hif1α remains to be elucidated, and further detailed experiments are needed in this regard.
As we postulated that neuronal apoptosis is due to the decrease in Vegf levels in the KO cortex, it is important to identify the source of Vegf in the telencephalon. Unfortunately, we could not detect the expression of Vegf mRNA and Vegf protein in the telencephalon using in situ hybridization and immunofluorescence (data not shown). In Vegf-LacZ knock-in mice, Vegf expression was detected in the ventral layer of the brain at E14.5 [35]. Based on this finding, we assumed that Vegf is primarily secreted from neural progenitor cells in the ventricular zone and that it probably acts as a paracrine neurotrophic factor to postmitotic or mature deep-layer neurons (Fig. 8). We do not have data to exclude the possibility of the autocrine function of Vegf. Vegf mRNA is highly expressed in cortical neurons at P7 [36], suggesting that Vegf may be secreted from mature deep-layer neurons. Based on our findings, we propose a model to explain the function of Vegf in neuronal survival (Fig. 8). Although certain unresolved points remain; it is unknown how Vegf signaling is activated specifically in deep-layer neurons and which receptor(s) mediate Vegf signaling. Considerable efforts are required to resolve these issues.
There lacks a consensus of whether Vegf directly regulates neuronal survival as a neurotrophic factor or indirectly maintains neurons through vascular formation. The telencephalon of KO mice exhibited abnormal vascular development (Supplementary Fig. 2), thus it is difficult at present to define the cellular mechanism of Vegf signaling in deep-layer neuron survival. However, our overexpression experiment showed that vascular networks normally develop whereas cortical layer formation is disturbed in the sFlt1-transfected cortex (Supplementary Fig. 3). These data suggest the primary role of Vegf as a neurotrophic factor rather than an endothelial cell growth factor during cortical development. In support of our findings, Lange et al. demonstrated that Gpr124-LacZ knock-in null embryos do not exhibit any neuronal apoptosis, even though the vasculature is largely diminished in the cortex [21]. Taken together, our findings emphasize that the neurotrophic function of Vegf is essential for cortical layer formation. However, we can not exclude the possibility that Vegf regulates neural migration as well. In future studies it will be important to confirm this possibility.
In conclusion, our findings provide insights into the molecular mechanisms underlying the development of murine embryonic telencephalon, which is regulated by Hif1α-dependent hypoxia signaling.
Mice
All mice were maintained on a C57BL/6 background. Hif1α flox/flox mice were obtained from The Jackson Laboratory (Stock No. 007561) and Sox1-Cre+/− mice from the RIKEN Bioresource Research Center (Accession No. CDB0525K, http://www2.clst.riken.jp/arg/mutant%20 mice%20list.html) [26]. Hif1α flox/flox mice and heterozygotes (Hif1α flox/wt; Sox1-Cre+/−) were crossed to obtain homozygotes (Hif1α flox/flox; Sox1-Cre+/−). For embryonic staging, the morning on which the vaginal plug was observed was designated as E0.5.
RT-qPCR
Total RNA was extracted from E8.5 and E9.5 WT and KO embryos, or the cortex of E13.5 WT and KO brains using the RNeasy Mini Kit (Qiagen) according to the manufacturer’s instructions. Isolated RNA was then reverse-transcribed to cDNA using PrimeScript 1st strand cDNA Synthesis Kit (Takara) according to the manufacturer’s instructions. qPCR using SYBR green was performed on a LightCycler Nano System (Roche). Gene expression was normalized to Gapdh expression levels. Each sample was analyzed at least in triplicate. The relative fold change was calculated using the 2-ΔΔCt method. For detection of Hif1α, Ldha, Aldoa, Epo, EpoR, Nqo1, Vegfa, and Gapdh expression levels, the following primers were used: Hif1α FW: TGAGCTTGCTCATCAGTTGC, Hif1α RV: CATAACAGAAGCTTTATCAAGATGTGA; Ldha FW: GGCACTGACGCAGACAAG, Ldha RV: TGATCACCTCGTAGGCACTG; Aldoa FW: TGGGAAGAAGGAGAACCTGA, Aldoa RV: GACAAGCGAGGCTGTTGG; Epo FW: TCTGCGACAGTCGAGTTCTG, Epo RV: CTTCtGCACAACCCATCGT; EpoR FW: GTCCTCATCTCGCTGTTGCT, EpoR RV: ATGCCAGGCCAGATCTTCT; Nqo1 FW: AGCGTTCGGTATTACGATCC, Nqo1 RV: AGTACAATCAGGGCTCTTCTCG; Vegfa FW: CTTGTTCAGAGCGGAGAAAGC, Vegfa RV: ACATCTGCAAGTACGTTCGTT; Gapdh FW: CATGTTCCAGTATGACTCCACTC, Gapdh RV: GGCCTCACCCCATTTGATGT.
In situ hybridization
A part of the mouse Hif1α sequence was PCR-amplified using the following primer set: FW-Hif1α: AAAGAGTCTGAAGTTTTTTATGAG and RV-Hif1α: CTTCTTAAGCTTATCAAAAAGGCA. The PCR products were cloned into the pGEM-T Easy vector (Promega) to synthesize the cRNA probe.
Whole-mount and cryosection in situ hybridization was performed as described previously [43, 44].
Immunofluorescence
Mouse brains were fixed with 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) for 3 h at 4o C and cryopreserved in 30% sucrose in PBS. Brains were embedded in optimal cutting temperature compound (OCT) and stored at − 80o C until further use. The cryostat sections were cut at 10 µm and adhered onto glass slides. The sections were washed with PBS and then incubated with 10 mM citric acid (pH 6.0) for 30 min at 80o C for antigen retrieval. After washing with PBS briefly, sections were incubated with 0.5% Triton X-100 in PBS for 15 min at room temperature and then with blocking buffer (3% BSA in TBST) for 30 min at room temperature. Further, the sections were incubated at 4oC overnight with primary antibodies against cleaved caspase 3 (9661, Cell Signaling; 1/200), Reelin (MAB5364, Chemicon; 1/1000), β-tubulin class III (Tuj1, PRB-435P, Covance; 1/1000), Satb2 (ab51502, Abcam; 1/500), and Ctip2 (ab18465, Abcam; 1/500). To visualize the vasculature, IB4-FITC (Sigma, 1/100) was used. Sections were washed thrice with TBST for 10 min and then incubated with the appropriate secondary antibodies conjugated with Alexa 488 or 546 (A11001, A21208, Invitrogen; 1/300) for 1 h at room temperature. Nuclei were stained with DAPI. Fluorescence microscopy was performed on a BX51 microscope equipped with a DP30BW CCD camera (Olympus) using 10× and 20× objective lenses. Images were acquired using the DP controller software (Olympus).
BrdU birthdating assay
Pregnant mice (E14.5) were injected intraperitoneally with BrdU at 0.1 mg/kg of body weight and euthanized at E18.5. Brains were fixed with 4% PFA solution in PBS for 3 h at 4o C and cryopreserved in 30% sucrose in PBS. Brains were embedded in OCT and stored at − 80o C until further use. The cryostat sections were cut at 12 µm and adhered onto glass slides. Then, the sections were washed with PBS and incubated with 10 mM citric acid (pH 6.0) for 30 min at 80oC for antigen retrieval. The sections were incubated with 0.1 N HCl for 1 h at 37oC. After incubation and brief washing with PBS, the sections were incubated with 0.5% Triton X-100 in PBS for 15 min at room temperature, and then incubated with blocking buffer (10% horse serum in TBST) for 30 min at room temperature. Further, the sections were incubated at 4oC overnight with primary antibodies against BrdU (347580, BD Bioscience; 1/100) and cleaved caspase 3 (9661, Cell Signaling; 1/200), washed twice with TBST for 10 min, and incubated with the appropriate secondary antibodies conjugated with Alexa 488 or 546 (A11001, A21208, Invitrogen; 1/300) for 1 h at room temperature. Fluorescence microscopy was performed on a BX51 microscope equipped with a DP30BW CCD camera (Olympus) using 10× and 20× objective lenses. Images were processed using the DP controller software (Olympus).
In utero electroporation
In utero electroporation was performed as previously described, with slight modifications [45]. Briefly, pregnant mice at E13.5 or E15.5 were anesthetized by intraperitoneal (IP) injection of saline containing butorphanol (5 mg/kg body weight), midazolam (4 mg/kg body weight), and medetomidine (0.3 mg per kg body weight), and then the uterine horns were exposed. Approximately 1 µL of plasmid DNA mixture with 0.01% Fast Green solution was injected into the lateral ventricle of embryos using a fine-tip glass micropipette (G-1.0, Narishige). The final concentration of plasmid DNA was as follows: pEGFP-CAGS (0.5 µg/µL), m-sFlt1-pCI (1 µg/µL), and pCI (1 µg/µL). Electric pulses (50 V, 1 pulse and 35 V, 4 pulses; 50 ms on, 950 ms off) were delivered to the dorsomedial part of the cortex using a forceps-type electrode (5 mm diameter, BEX) through CUY21 EDIT II (BEX). After wound closure, anesthesia was terminated by IP injection of atipamezole (3 mg/kg body weight). The embryos were then allowed to develop to E18.5.
Statistics
For statistical analysis, two-tailed Student's t-test was used to determine p-values, and values with p less than 0.05 were considered statistically significant.
Ethics approval: All animal experiments were performed in accordance with the Guidelines for the Care and Use of Laboratory Animals of Kanazawa Medical University and Doshisha University. A minimum sample size of five individuals was used in each assay, unless otherwise stated.
Consent for publication: Not applicable.
Availability of data and materials: The datasets used and /or analyzed during the current study are available from the corresponding author on reasonable request.
Competing interests: The authors declare that they have no competing interests.
Funding: This study was supported by MEXT KAKENHI, grant numbers 19K06680 and 17H05965 to DS and Assist KAKEN from Kanazawa Medical University, grant number K2019-3 to DS.
Authors’ contributions: DS and HS designed the study; DS, TS, TK, and YM performed experiments; MT and HM analyzed the RT-qPCR data; HS-H, TH, and HS interpreted the data and edited the manuscript; DS wrote the paper. All authors read and approved the final version of the manuscript.
Acknowledgements: We thank Dr. Angelo Iulianella (Dalhousie University) for critical reading of the manuscript. We also thank Dr. Noriko Osumi and Dr. Yoshio Wakamatsu (Tohoku University) for their valuable help and advice. m-sFlt1-pCI was kindly gifted by Dr. Takayuki Ikeda (Kanazawa Medical University). The Sox1-Cre+/- mouse strain (RBRC05065) was provided by RIKEN through the National Bio-Resource Project of the MEXT, Japan. We would like to thank Editage (www.editage.com) for English language editing.
- Dunwoodie SL. The role of hypoxia in development of the Mammalian embryo. Dev Cell. 2009;17:755–73.
- Fischer B, Bavister BD. Oxygen tension in the oviduct and uterus of rhesus monkeys, hamsters and rabbits. J Reprod Fertil. 1993;99:673–9.
- Leese HJ. Metabolic control during preimplantation mammalian development. Hum Reprod Update. 1995;1:63–72.
- Ufer C, Wang CC. The Roles of Glutathione Peroxidases during Embryo Development. Front Mol Neurosci. 2011;4:12.
- Lee YM, Jeong CH, Koo SY, Son MJ, Song HS, Bae SK, et al. Determination of hypoxic region by hypoxia marker in developing mouse embryos in vivo: a possible signal for vessel development. Dev Dyn. 2001;220:175–86.
- Ream M, Ray AM, Chandra R, Chikaraishi DM. Early fetal hypoxia leads to growth restriction and myocardial thinning. Am J Physiol Regul Integr Comp Physiol. 2008;295:R583-95.
- Morriss GM, New DA. Effect of oxygen concentration on morphogenesis of cranial neural folds and neural crest in cultured rat embryos. J Embryol Exp Morphol. 1979;54:17–35.
- Semenza GL, Wang GL. A nuclear factor induced by hypoxia via de novo protein synthesis binds to the human erythropoietin gene enhancer at a site required for transcriptional activation. Mol Cell Biol. 1992;12:5447–54.
- Semenza GL. Hypoxia-inducible factor 1: master regulator of O2 homeostasis. Curr Opin Genet Dev. 1998;8:588–94.
- Wang GL, Jiang BH, Rue EA, Semenza GL. Hypoxia-inducible factor 1 is a basic-helix-loop-helix-PAS heterodimer regulated by cellular O2 tension. Proc Natl Acad Sci U S A. 1995;92:5510–4.
- Wang GL, Semenza GL. Purification and characterization of hypoxia-inducible factor 1. J Biol Chem. 1995;270:1230–7.
- Kaelin WG Jr, Ratcliffe PJ. Oxygen sensing by metazoans: the central role of the HIF hydroxylase pathway. Mol Cell. 2008;30:393–402.
- Ruas JL, Poellinger L. Hypoxia-dependent activation of HIF into a transcriptional regulator. Semin Cell Dev Biol. 2005;16:514–22.
- Iyer NV, Kotch LE, Agani F, Leung SW, Laughner E, Wenger RH, et al. Cellular and developmental control of O2 homeostasis by hypoxia-inducible factor 1 alpha. Genes Dev. 1998;12:149–62.
- Kotch LE, Iyer NV, Laughner E, Semenza GL. Defective vascularization of HIF-1alpha-null embryos is not associated with VEGF deficiency but with mesenchymal cell death. Dev Biol. 1999;209:254–67.
- Ryan HE, Lo J, Johnson RS. HIF-1 alpha is required for solid tumor formation and embryonic vascularization. EMBO J. 1998;17:3005–15.
- Compernolle V, Brusselmans K, Franco D, Moorman A, Dewerchin M, Collen D, et al. Cardia bifida, defective heart development and abnormal neural crest migration in embryos lacking hypoxia-inducible factor-1alpha. Cardiovasc Res. 2003;60:569–79.
- Amarilio R, Viukov SV, Sharir A, Eshkar-Oren I, Johnson RS, Zelzer E. HIF1alpha regulation of Sox9 is necessary to maintain differentiation of hypoxic prechondrogenic cells during early skeletogenesis. Development. 2007;134:3917–28.
- Krishnan J, Ahuja P, Bodenmann S, Knapik D, Perriard E, Krek W, et al. Essential role of developmentally activated hypoxia-inducible factor 1alpha for cardiac morphogenesis and function. Circ Res. 2008;103:1139–46.
- Provot S, Zinyk D, Gunes Y, Kathri R, Le Q, Kronenberg HM, et al. Hif-1alpha regulates differentiation of limb bud mesenchyme and joint development. J Cell Biol. 2007;177:451–64.
- Lange C, Turrero Garcia M, Decimo I, Bifari F, Eelen G, Quaegebeur A, et al. Relief of hypoxia by angiogenesis promotes neural stem cell differentiation by targeting glycolysis. EMBO J. 2016;35:924–41.
- Wagenfuhr L, Meyer AK, Braunschweig L, Marrone L, Storch A. Brain oxygen tension controls the expansion of outer subventricular zone-like basal progenitors in the developing mouse brain. Development. 2015;142:2904–15.
- Tomita S, Ueno M, Sakamoto M, Kitahama Y, Ueki M, Maekawa N, et al. Defective brain development in mice lacking the Hif-1alpha gene in neural cells. Mol Cell Biol. 2003;23:6739–49.
- Isaka F, Ishibashi M, Taki W, Hashimoto N, Nakanishi S, Kageyama R. Ectopic expression of the bHLH gene Math1 disturbs neural development. Eur J Neurosci. 1999;11:2582–8.
- Liang H, Hippenmeyer S, Ghashghaei HT. A Nestin-cre transgenic mouse is insufficient for recombination in early embryonic neural progenitors. Biol Open. 2012;1:1200–3.
- Takashima Y, Era T, Nakao K, Kondo S, Kasuga M, Smith AG, et al. Neuroepithelial cells supply an initial transient wave of MSC differentiation. Cell. 2007;129:1377–88.
- Dekimoto H, Terashima T, Katsuyama Y. Dispersion of the neurons expressing layer specific markers in the reeler brain. Dev Growth Differ. 2010;52:181–93.
- Sekine K, Kubo K, Nakajima K. How does Reelin control neuronal migration and layer formation in the developing mammalian neocortex? Neurosci Res. 2014;86:50–8.
- Hevner RF, Neogi T, Englund C, Daza RA, Fink A. Cajal-Retzius cells in the mouse: transcription factors, neurotransmitters, and birthdays suggest a pallial origin. Brain Res Dev Brain Res. 2003;141:39–53.
- Gotz M, Huttner WB. The cell biology of neurogenesis. Nat Rev Mol Cell Biol. 2005;6:777–88.
- Rosenstein JM, Mani N, Khaibullina A, Krum JM. Neurotrophic effects of vascular endothelial growth factor on organotypic cortical explants and primary cortical neurons. J Neurosci. 2003;23:11036–44.
- Silverman WF, Krum JM, Mani N, Rosenstein JM. Vascular, glial and neuronal effects of vascular endothelial growth factor in mesencephalic explant cultures. Neuroscience. 1999;90:1529–41.
- Raab S, Beck H, Gaumann A, Yuce A, Gerber HP, Plate K, et al. Impaired brain angiogenesis and neuronal apoptosis induced by conditional homozygous inactivation of vascular endothelial growth factor. Thromb Haemost. 2004;91:595–605.
- Haigh JJ, Morelli PI, Gerhardt H, Haigh K, Tsien J, Damert A, et al. Cortical and retinal defects caused by dosage-dependent reductions in VEGF-A paracrine signaling. Dev Biol. 2003;262:225–41.
- Miquerol L, Gertsenstein M, Harpal K, Rossant J, Nagy A. Multiple developmental roles of VEGF suggested by a LacZ-tagged allele. Dev Biol. 1999;212:307–22.
- Okabe K, Fukada H, Tai-Nagara I, Ando T, Honda T, Nakajima K, et al. Neuron-derived VEGF contributes to cortical and hippocampal development independently of VEGFR1/2-mediated neurotrophism. Dev Biol. 2020;459:65–71.
- Fantin A, Schwarz Q, Davidson K, Normando EM, Denti L, Ruhrberg C. The cytoplasmic domain of neuropilin 1 is dispensable for angiogenesis, but promotes the spatial separation of retinal arteries and veins. Development. 2011;138:4185–91.
- Giger RJ, Cloutier JF, Sahay A, Prinjha RK, Levengood DV, Moore SE, et al. Neuropilin-2 is required in vivo for selective axon guidance responses to secreted semaphorins. Neuron. 2000;25:29–41.
- Kendall RL, Thomas KA. Inhibition of vascular endothelial cell growth factor activity by an endogenously encoded soluble receptor. Proc Natl Acad Sci U S A. 1993;90:10705–9.
- Takahashi T, Takase Y, Yoshino T, Saito D, Tadokoro R, Takahashi Y. Angiogenesis in the developing spinal cord: blood vessel exclusion from neural progenitor region is mediated by VEGF and its antagonists. PLoS One. 2015;10:e0116119.
- Wood HB, Episkopou V. Comparative expression of the mouse Sox1, Sox2 and Sox3 genes from pre-gastrulation to early somite stages. Mech Dev. 1999;86:197–201.
- Chen G, Sima J, Jin M, Wang KY, Xue XJ, Zheng W, et al. Semaphorin-3A guides radial migration of cortical neurons during development. Nat Neurosci. 2008;11:36–44.
- Sakai D, Dixon J, Dixon MJ, Trainor PA. Mammalian neurogenesis requires Treacle-Plk1 for precise control of spindle orientation, mitotic progression, and maintenance of neural progenitor cells. PLoS Genet. 2012;8:e1002566.
- Ogoh H, Yamagata K, Nakao T, Sandell LL, Yamamoto A, Yamashita A, et al. Mllt10 knockout mouse model reveals critical role of Af10-dependent H3K79 methylation in midfacial development. Sci Rep. 2017;7:11922.
- Kawauchi T, Chihama K, Nabeshima Y, Hoshino M. The in vivo roles of STEF/Tiam1, Rac1 and JNK in cortical neuronal migration. EMBO J. 2003;22:4190–201.