Ablation of SAMD1 in Mice Causes Failure of Embryonic Blood Vessel Maturation and Embryonic Lethality

doi:10.21203/rs.3.rs-2117607/v1

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Ablation of SAMD1 in Mice Causes Failure of Embryonic Blood Vessel Maturation and Embryonic Lethality

https://doi.org/10.21203/rs.3.rs-2117607/v1

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published 21 Feb, 2023

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SAM domain-containing protein 1 (SAMD1) has been implicated in atherosclerosis, as well as in chromatin and transcriptional regulation, suggesting a versatile and complex biological function. However, its role at an organismal level is currently unknown. Here, we generated SAMD1−/− and SAMD1+/− mice to explore the role of SAMD1 during mouse embryogenesis. Homozygous loss of SAMD1 was embryonic lethal with no living animals seen after embryonic day 18.5. At embryonic day 14.5, organs were degrading and/or incompletely developed and no functional blood vessels were observed. Sparse red blood cells were scattered and pooled, primarily near the embryo surface. Some embryos had malformed heads and brains at embryonic day 15.5. In vitro, SAMD1 absence impaired neuronal differentiation processes. Heterozygous SAMD1 knockout mice were born alive, but postnatal genotyping showed a reduced ability of these mice to thrive, possibly due to altered steroidogenesis. In summary, the characterization of SAMD1 knockout mice suggests a critical role of SAMD1 during developmental processes in multiple organs and tissues.

Biological sciences/Developmental biology

Biological sciences/Molecular biology/Chromatin

SAMD1

Embryogenesis

Arteriogenesis

Angiogenesis

Development

Exencephaly

Neuronal Differentiation

Human SAMD1 (SAM domain-containing protein 1) is a 538 amino acid protein, which has orthologues in most vertebrates, including zebrafish. Originally, SAMD1 was identified as a protein involved in atherosclerotic LDL binding in humans¹ and mice². SAMD1 may participate in low-density lipoprotein (LDL) retention and uptake, and its knockdown suppresses vascular smooth muscle cell (VSMC) differentiation and proliferation, which affects foam cell development³. However, SAMD1 is widely expressed in most organs and tissues, both in mice (EMBL-EBI) and humans (ProteinAtlas, GTEx). The extensiveness of expression suggests roles in adult homeostasis, but the molecular function of SAMD1 was minimally studied so far. Recently, SAMD1 was found to directly associate with CpG islands at the chromatin, acting as a transcriptional regulator, and be required for proper embryonic stem (ES) cell differentiation⁴. Earlier reports further support epigenetic functions for SAMD1. SAMD1 was enriched at nucleosomes and H3K4me3-possessing chromatin regions, as revealed by mass spectrometry^5,6. It was also among the highest 5% of proteins that associate with both H3K4me3-modified and bivalently H3K4me3/H3K27me3-modified chromatin ⁷.

The SAMD1 protein has two globular domains. At the C-terminus, it possesses a SAM domain that can interact with other SAM domain-containing proteins, such as L3MBTL3, and allows multimerization⁴. The N-terminal winged helix domain binds to unmethylated CpG motifs. SAMD1 associates with chromatin regulatory complexes that contain the histone demethylase KDM1A and several other SAM domain proteins including L3MBTL3 and SFMBT1^4,8−10. It is often upregulated in cancer tissues and correlates with worse prognosis in some cancer types, such as adenoid cystic carcinoma (ACC) and liver cancer^11,12. A CRISPR screen in K562 cells suggested that SAMD1 is required for the efficient proliferation of these cells¹³. Further work demonstrated that SAMD1 deletion in HepG2 hepatocellular carcinoma cells leads to a global readjustment of the active H3K4me2 chromatin mark and a more favorable gene signature, supporting an oncogenic role in this context¹². SAMD1 has also been described to play a role in antiphospholipid syndrome¹⁴ and muscle adaptation¹⁵, implying a rather versatile biological function. Taken together, these findings suggest that SAMD1 likely has a function in multiple biological processes and diseases, but the biological role of SAMD1 at an organismal level has yet to be investigated.

Here, we report the first knockout of SAMD1 in mice. Fewer homozygous (SAMD1−/−) embryos were observed at embryonic day (E) 14.5, and none were observed after E18.5. The development appeared grossly normal at E12.5, but at E14.5 the investigated embryos exhibited numerous defects, including degrading endothelial cell (EC) tubes and an almost complete absence of red blood cells (RBCs). Approximately 70% of heterozygotic (SAMD1+/−) mice survived past week 3 postnatally but exhibited reduced weight gain and hormonal changes.

SAMD1 knockout mice are embryonic lethal

The SAMD1 gene was deleted from C57BL/6J mice by recombineering (Fig. 1a), which led to the removal of 2396 bps of the SAMD1 gene. Successfully obtained heterozygous mice were mated. The ratios between the wild-type (WT, aka SAMD1+/+), heterozygous (HET, aka SAMD1+/−), and knockout (KO, aka SAMD1−/−) mice were close to the expected Mendelian ratios through E12.5. The first dead SAMD1 KO embryos, and abnormal phenotypes were observed at E14.5 (Fig. 1b) and manifested at later time points (Supplementary Table S1) (p = 0.049). Dying pups were not seen, so the few KO mice that had not been resorbed by E18.5 died just prior to birth (Supplementary Table S1). This suggests there was variable expression and penetrance, and thus multiple embryonic lethal SAMD1 KO phenotypes. Crossing SAMD1+/− mice with SAMD1+/− mice and phenotyping 3 weeks after birth (P21) also showed no KO’s (p = 5.1 x 10^− 13) (Supplementary Table S2), confirming the lethality of the SAMD1 KO.

In contrast, the E14.5 heterozygous (HET) embryos were not noticeably different from that of the WT embryo, and HET mice were born alive. However, genotyping at P21 showed that the number of observed HETs was significantly lower than the expected number, when SAMD1+/− mice were crossed with SAMD1+/− mice (p = 0.003) (Supplementary Table S2), or when SAMD1+/+ mice were crossed with SAMD1+/- mice (p = 0.003) (Supplementary Table S3).

Embryonic fibroblasts (MEFs) raised from WT, HET, and KO mice were investigated for SAMD1 expression via RT-qPCR. SAMD1 mRNA was absent in the homozygous KO cells and slightly but not significantly reduced in the heterozygous cells (Fig. 1c).

Defects of the SAMD1 KO embryos are visually obvious

Macrolevel photography of the E14.5 WT embryos within the yolk sac showed normal development, noting branching blood vessels in the yolk sac (Fig. 1b,d). In contrast, the yolk sac of the KO embryos lacked obvious blood vessels, it had only a few broken thin red lines, and the amniotic fluid was pink as if blood had leaked into it (Fig. 1b, 1d). In the KO the yolk sac did not seem to be normally attached to the placental disk.

When the yolk sac was removed, the KO embryo within the amnion appeared developmentally delayed and visually smaller than the WT (Fig. 1e). Bloody fluid was also pooled around the midsection within the amnion (Fig. 1e). The embryo surface was very pale, and across the back the edema could be seen separating the skin from the tissue beneath. Red blood cells were not obvious in the skin, other than on the head and back, where a few small hemorrhages and a few broken red lines that were probably failed blood vessels were recognizable.

The KO placenta was pale compared to the WT (Fig. 1e), probably because the labyrinth contained fewer RBCs and the vasculature seemed to be pink instead of red. The metrial gland lacked obvious blood vessels and the decidua appeared to be thinner. Maternal blood vessels could be seen in a few locations immediately beneath the decidua. The umbilical cord was avascular (Fig. 1e).

At E15.5 (Fig. 1f), a SAMD1 KO embryo lacked a skull vault, had obvious exencephaly, and had a hypertrophic brain. Clear edema fluid separated the skin from most of the embryo surface. Some areas of the embryo surface beneath the skin, particularly above the brain, had spots and lines of RBCs that may have once been contained in vessels, and small hemorrhages. RBCs could not have arrived at these locations without a functional circulatory system. Since the E14.5 KO had a skull vault, the investigated E15.5 KO embryo is likely an example of a different SAMD1 KO phenotype, but the surface RBC patterns appear similar to the E14.5 KO. Failed skull vault development may be caused by failure of neural tube closure. Fluid leakage due to failing capillaries likely produced the edema (Fig. 1f), although failing lymphatic vessels, which are also constructed from ECs and SMCs, may have added to the edema fluids.

Given the strong defects observed in SAMD1 KO embryos, we used public gene transcription data to assess embryonic SAMD1 expression^16,17. SAMD1 mRNA was detected in very early developmental stages (Fig. 1g) as well as in all investigated organs during all embryonic stages (Fig. 1h). Similar results were also obtained using RT-qPCR experiments from isolated embryonic organs (Supplementary Figure S1). This observation suggests that SAMD1 has a biological function during all embryonic stages and in most embryonic tissues.

SAMD1 KO embryos have multiple organ defects

To further assess the consequence of the SAMD1 KO on specific organs, we first used hematoxylin and eosin (H&E) staining on sagittal slices of an E14.5 embryo (Fig. 2). Hematoxylin reveals not only nuclei, but also glycosaminoglycans, for example in cell walls, and thus allows visualization of fragmenting cells, while eosin stains cytoplasm and most connective tissue pink, orange, and/or red¹⁸. A comparison of embryonic tissues and organs in WT and KO mice revealed several abnormalities (Fig. 2). Although cardiac muscle cells stained red and had associated nuclei, the heart appeared to be breaking up, and the atria, ventricles, and pulmonary trunk had partially collapsed (Fig. 2a). Endocardium was separating from the chamber walls. Red-stained RBCs were absent, and the ventricles appeared to contain some lymphocytes/immune cells instead of RBCs (Fig. 2a). This suggests RBCs were trapped due to ejection failure and became necrotic.

The KO lung appeared also to be fragmenting, and RBCs were not apparent, in contrast to the WT lung where RBCs filled large vessels (Fig. 2b). Bronchioles had formed but were smaller than those in the WT. They appeared to be degrading and may not have formed correctly. Faint pink staining of connective tissue in the lung and diaphragm may indicate dying cells.

The KO liver had lobes and was large enough to have developed at least until E12.5 (Fig. 2c). Similar to the heart, the liver had no RBCs, again in contrast to the WT, where RBCs filled large vessels. The tissue was broken up in patterns that suggested degradation rather than a malformation. The fact that organs were substantially developed means that a functional circulatory system necessarily existed prior to E14.5.

SAMD1 deletion led to disorganization and reduced levels of RBCs in the skin

The most obvious difference between the appearances of WT and KO embryos was the paleness of the latter, suggesting impaired blood circulation. The E12.5 KOs appeared normal, and E14.5 has been established as an optimal time point for mouse developmental disorder phenotype analyses¹⁹. Thus, we tested an E14.5 embryo for the presence of endothelial cells (ECs), which line all blood vessels, from primitive endothelial tubes to fully mature arteries and veins, as well as lymph vessels. We used H&E, CD31 or VEGFR2 staining (with hematoxylin counterstain) to mark ECs and provide morphological identification of other cells. Cluster of Differentiation 31 (CD31, also known as PECAM1) is widely used as a marker for ECs, staining the cell surface. Vascular endothelial growth factor 2 (VEGFR2, also known as FLK1), stains the cytoplasm of ECs. Using these different markers, we investigated the organs and tissues of the SAMD1 KO embryo.

Looking first at the embryo surface, H&E staining of the WT embryo showed groups of RBCs in the skin and RBCs in a larger blood vessel beneath the subcutaneous tissue. Here, CD31 stained ECs of capillaries and larger vessels (Fig. 3a-c), as expected. In contrast, KO skin, except around the limbs and skull, appeared to have been lost during fixation (Fig. 3a), likely due to edema separating the skin from the tissue beneath. In locations beneath the lost skin, the KO embryonic surface appeared to consist of degrading and fragmenting skeletal muscle (Fig. 3a, d). This tissue included individual RBCs and a few scattered clusters of RBCs that, lacking typical organizational patterns, did not appear to be contained in vessels (Fig. 3a-c). Coagulation in the clusters could not be determined, but gradations in red staining implied the recent onset of RBC degradation. These RBCs could be the result of fresh microhemorrhages, or RBCs recently stranded as vessel development failure halted circulation. Consistent with this second hypothesis, intact vessels were not obvious, but RBCs could not have arrived at their present locations unless at least functional endothelial tubes had once been present. CD31 stained numerous misshapen subcutaneous ECs in broken brown lines (Fig. 3e). These patterns suggested that the ECs were part of capillaries or larger vessels that had ceased providing circulatory function. Cytoplasmic instead of cell surface CD31 brown staining patterns suggest necrosis, and the association of CD31 staining with rounded nuclei suggests possible phagocytosis. Several broken small circles of stained ECs also suggested failed vessels (Fig. 3d, e).

Edema did not separate the skin from the KO forepaws, which appeared to have developed normally except for being smaller, pale, and lacking intact blood vessels (Fig. 3f). H&E staining of the forepaw showed a small group of faintly stained RBCs, where faint blurred hematoxylin stain suggested impending necrosis. CD31 showed a few nuclei partially surrounded by brown-stained EC material, suggesting ingestion. The patterns of broken lines suggest the presence of failed small vessels in the forepaw (Fig. 3f, arrow). Interestingly, red-stained RBCs were noted primarily at the embryo surface, but not in the heart or other organs. Surface perfusion may have been sufficient to delay RBC necrosis compared to locations deeper in the embryo.

SAMD1 KO embryos have degraded internal organs

To determine the impact of SAMD1 deletion on internal organs, we stained lung, liver (Fig. 4), and heart samples (Fig. 5) from E14.5 WT and KO mice with H&E, CD31, or VEGFR2. In all WT samples, H&E staining showed normal organ development and large blood vessels filled with RBCs (Fig. 4, 5). Larger lung and liver vessels, identified by sizeable areas of RBCs, were encircled by stained ECs (Fig. 4b,4d). Capillaries were also made evident by the presence of EC markers and RBCs. Pulmonary vascular ECs normally line the surfaces of the lung vasculature, which stained strongly for CD31 and VEGFR2, and epithelial cells in bronchioles stained faintly for VEGFR2 (Fig. 4c). Liver sinusoidal ECs also stained for CD31 and VEGFR2 (Fig. 4e).

In contrast, in the lung and liver of the SAMD1 KO mice, CD31 and VEGFR2 staining were almost absent. In KO organs, no RBCs or intact ECs were visible (Fig. 4a, b, d, e). The limited CD31 and VEGFR2 staining in the lung and liver appeared to be cell fragments of necrotic vessel wall ECs, and the thin lines of associated faint hematoxylin stain likely mark dermatan sulfate and heparan sulfate from these ECs. Examples of probable phagocytosis were indicated by uneven brown staining around strongly blue-stained round nuclei (Fig. 4b, c, e, red arrow, Supplementary Figure S2). This implies that ECs had been present, became necrotic, and were being removed. Bronchioles had formed in the KO lungs but were much smaller than those in the WT lungs (Fig. 4c). Based on the available data, it is not possible to judge whether the smaller bronchioles in the KO were due to incomplete formation during development or whether they had degraded (Fig. 4c).

In the WT hearts, normal CD31 and VEGFR2 staining patterns were noted in ECs of capillaries and larger vessels (Fig. 5a, b, c). H&E staining of the WT pulmonary trunk showed numerous RBCs contained in the vessel, demonstrating normal heart function. VEGFR staining also identified ECs at the lumen and several layers of circumferentially oriented spindle-shaped vascular smooth muscle cells (VSMCs) (Fig. 5d). As expected, epithelial cells in the endocardium stained for CD31 and VEGFR2 (Fig. 5e).

In the KO heart, H&E staining uncovered signs of epithelial, cardiac muscle cell, and EC degradation (Fig. 2a, Fig. 5b). Hematoxylin counterstain in the CD31 and VEGFR2 slides revealed fragmenting cardiac muscle cells (Fig. 5b). In a few places, round hematoxylin-blue nuclei were loosely associated with faint CD31 or VEGFR2-brown patterns demonstrating the presence of necrotic ECs and epithelial cells (Fig. 5c). Phagocytosis was indicated by uneven and foamy brown staining around strongly blue-stained round nuclei (Supplementary Figure S2). CD31 and VEGFR2 stained broken circles and lines of misshapen ECs and EC fragments. These patterns suggested incomplete and/or failed vessels (Fig. 5b, c). This was seen deep in the heart muscle where capillaries should have been (Fig. 5b), and more frequently at the heart’s surface, where larger stained circles suggested failed coronary vessels (Fig. 5c). Spindle-shaped VSMCs (blue) that were noted next to coronary artery ECs in the WT, were absent from the KO (Fig. 5e). The KO pulmonary trunk wall consisted only of the myocardium, with a few disconnected ECs above the lumen, and lacked the VSMCs that were seen in the WT (Fig. 5a, d). The lumen of the KO pulmonary trunk contained no RBCs and only a few rounded probable immune cells (Fig. 5d). The heart chambers and large vessels were partially collapsed, possibly due to serum fluid being insufficient to maintain shape (Fig. 2a, 5a-c). The ventricles contained dark blue stained rounded probable immune cells and cell fragments that may have been RBCs, but trabeculation was not observed. Epithelial and endothelial cells are closely linked - during embryonic development, the epithelial layer is the source of endothelial cells for the heart’s blood vessels²⁰. CD31- and VEGFR2-stained epithelial cell fragments were detectable in the KO endocardium, including detached cells and cells undergoing probable necrosis and phagocytosis (Fig. 5e). These above findings suggest that in the investigated SAMD1 KO mouse the development of the heart may have stopped before VSMC differentiation.

SAMD1 KO ribs showed premature ossification

Next, we investigated ribs and associated skeletal muscles. Structural organization in the E14.5 WT and KO mice were roughly similar (Fig. 6a), but H&E staining highlighted substantial differences. The WT rib consisted of cartilage primordium, while in the KO mouse, the rib was ossifying, as seen by the bubbly appearance of hypertrophic chondrocytes, some of which lack nuclei (Fig. 6b, arrow). Bone development is tightly linked to hypoxia²¹, which is required for the transition from proliferating cartilage to endochondral ossification. Hypoxia interior to the developing bone causes chondrocytes to become hypertrophic and delays chondrocyte apoptosis/necrosis. Subsequent apoptosis/necrosis is required for ossification²¹. Thus, we hypothesize that in the SAMD1 KO mice, the hypertrophic chondrocytes appeared to have begun necrosis before E14.5, instead of the normal condition at approximately E18, suggesting an early hypoxic condition, possibly due to the lack of RBC delivery.

Skeletal muscle cells in the WT mouse stained strongly with H&E and had associated blue nuclei (Fig. 6c). Strands of cells are separated by lines of unstained connective tissue (Fig. 6c). In the KO, there are several signs of degradation. The KO has fewer eosin-stained red strands, which are less strongly stained red and are more widely separated from each other than in the WT (Fig. 6c). The strands are also interspersed with many nuclei. Some of these nuclei may be immune cells that are attracted to necrotizing muscle cells. Hypoxia from a lack of delivery of RBCs is the likely cause of muscle cell necrosis.

RBCs (bright orange) in a blood vessel above a WT rib do not have nuclei, and few if any lymphocytes are seen (Fig. 6d). RBCs are absent from the KO, other than a few scattered pools near the embryo surface and near ribs (Fig. 6a). A small blood pool above a KO rib contains some RBCs with eccentric nuclei and many lymphocytes/immune cells (rounded, blue) (Fig. 6d). The initiation of heartbeat around E8.5 marks the onset of embryo-vitelline circulation, as yolk sac-derived hematopoietic cells are spread through the developing embryo²². These primitive erythroid (EryP) cells enucleate between E12.5 and E16.5²³. RBCs could not have arrived at these locations unless a functional circulatory system existed prior to E14.5. EryP enucleation takes several days after entering circulation, suggesting that circulation continued until at least E12, and subsequent circulation failure trapped EryPs in a nonfunctioning vessel. Some KO RBCs appeared faded (Fig. 6d), implying a recent loss of O₂ from hemoglobin, and faint pink staining between RBCs and lymphocytes/immune cells suggests phagocytosis of RBCs.

SAMD1 is required for proper neuronal differentiation in vitro

To better understand the molecular source of the many defects upon SAMD1 deletion, we investigated our previously published RNA-Seq data upon undirected ES cell differentiation⁴. We found that multiple pathways are affected upon SAMD1 KO, implicating a rather pleiotropic role of SAMD1 in the differentiation process, consistent with the multifaceted mouse phenotype. A closer investigation of the data using GSEA (gene set enrichment analysis) suggests that pathways related to angiogenesis are mostly downregulated (Supplementary Figure S3a). Intriguingly, vascular endothelial growth factor A (VEGFA), PECAM1 (CD31), and THY1, which are critical factors for angiogenesis and arteriogenesis^24,25, were downregulated in SAMD1 KO cells, both in differentiated and undifferentiated cells (Supplementary Figure S3b). Furthermore, we found that pathways related to cardiac chamber development were significantly dysregulated in differentiated SAMD1 KO cells (Supplementary Figure S3c). This includes key transcription factors such as GATA4, GATA6, and MEF2C²⁶ (Supplementary Figure S3d). In addition to defects in blood vessel maturation, we also observed malformation of the brain and skull vault (Fig. 1f). Several genes related to brain development, such as CBLN1, NTRK2, and PLXNA4, are directly bound by SAMD1 and become derepressed upon SAMD1 deletion during differentiation (Supplementary Figure S3f). Consistently, pathways that are linked to brain development, such as synapse assembly (Supplementary Figure S3e), become predominantly upregulated in SAMD1 KO cells upon differentiation, suggesting that SAMD1 may enhance neuronal differentiation processes.

Mouse ES cells can be differentiated into various lineages using specific protocols²⁷. Since we observed severe exencephaly at E15.5 (Fig. 1f), implicating defects during neuronal developmental processes, and since neuronal differentiation belongs to the most commonly performed differentiation procedure²⁷, we addressed the role of SAMD1 during neuronal differentiation (Fig. 7a). We used our previously established SAMD1 KO mouse ES cells (Fig. 7b)⁴ and differentiated them into neuronal cells. First, we created embryoid bodies, which already showed dysregulation of multiple genes (Fig. 7c). The embryoid bodies were differentiated into neuronal progenitor cells (NPCs) and subsequently into neuronal cells (Fig. 7a). We observed no significant differences in the cell growth of the NPCs, although SAMD1 KO NPCs trended to grow slightly faster than wild-type NPCs (Fig. 7d). Upon differentiation of NPCs into neuronal cells, we observed that several marker genes, such as glial fibrillary acidic protein (GFAP) and Nestin (NES), were dysregulated upon SAMD1 KO (Fig. 7e). Immunofluorescence of the differentiated cells on day 6 showed an enhanced level of Tuj1 (Tubb3)-stained neurons in the KO cells (Fig. 7f), indicating that SAMD1 KO enhances the differentiation preferentially towards neurons. Furthermore, we observed higher levels of H3K4me2 in the differentiated cells (Fig. 7f, g). This observation supports that SAMD1 deletion may impair the activity of the KDM1A histone demethylase complex during neuronal differentiation, in line with our previous observation in undifferentiated ES cells⁴. However, other indirect effects could also be the source of this observation. Combined, these data support the hypothesis that the absence of SAMD1 leads to aberrant neuronal differentiation processes.

SAMD1 heterozygous mice fail to strive

Finally, we assessed the consequence of a heterozygous deletion of SAMD1. H&E and CD31 staining in an E14.5 heterozygous (HET) embryo were not noticeably different from that in the WT embryo (Supplementary Fig. 4), and HET mice were born alive, but approximately 30% of the HETs failed to survive past P21 (Supplementary Table S2, S3)

Longitudinal body weight analysis on regular chow revealed that male and female HET mice weighed less than WT mice (Fig. 8a). High-fat diet (HFD) is commonly used to induce metabolic changes in mice. HFD (60% fat by kcal, vegetable shortening) was started at 4 weeks and continued for 15 weeks. We found that after HFD, HETs weighed less than control WT (Fig. 8b), and fat depots from HET mice weighed approximately half as much as those from the WT mice (Fig. 8c). Consistent with what appeared to be reduced adiposity in the SAMD1 HET mice on the HFD, glucose disposal was increased in the HET mice during the oral glucose tolerance test (OGTT) (Fig. 8d) and baseline (time 0) fasted glucose levels were significantly lower in the HET mice following the HFD challenge (Fig. 8e). Insulin measurements from the blood samples collected during the OGTT found that insulin levels in the HET mice did not increase following the HFD (Fig. 8f) and were significantly lower than the corresponding levels in the WT mice (Fig. 8g).

Serum VLDL (very low-density lipoprotein) and LDL (low-density lipoprotein) were higher in the HETs, while HDL (high-density lipoprotein) was lower in the HETs (Fig. 8h). Proper steroidogenesis is essential for correct lipid metabolism²⁸. Indeed, the steroid hormone levels appeared to be altered in adult HETs. Corticosterone, aldosterone, and angiotensin II levels were significantly increased, and testosterone levels trended higher (Fig. 8i). Steroidogenesis and transformation of cholesterol to oxysterols occurs in many cell types²⁹. It is possible that the approximately 10–15% reduction in SAMD1 expression in SAMD1 HET mice (Fig. 1c) caused or required a compensatory change in steroid expression. The lower weights and adiposity may also be related to steroidal differences. Notably, we measured large differences in the above markers between individual adult HETs. It is likely that the HETs dying prior to postnatal week 3 expressed a more severe phenotype, resulting in a failure to thrive.

In this study, we investigated the effects of ablation of the SAMD1 gene in mice. SAMD1 is an epigenetic regulator that plays a gene regulatory role at unmethylated CpG islands^4,12. Upon mouse ES cell differentiation, the absence of SAMD1 enhances the expression of genes involved in neuronal pathways, while factors important for metabolism and angiogenesis are downregulated (Supplementary Figure S3)⁴. In addition to gene regulatory functions SAMD1 may also have a role in atherosclerosis^1,3 and muscle adaptation ¹⁵.

Looking at the whole organism, we noted that without SAMD1 there was an absence of functional blood vessels leading to a pale appearance of the embryo by E14.5 (Fig. 1e). Nonetheless, SAMD1 KO mice, including the extra-embryonic tissues, necessarily had developed a vascular plexus, at least to the point of EC tubes. This is shown by the extent of organ development and the presence of EC fragments in locations where vasculature would normally have been. Additionally, the observed RBCs and RBC clusters could not have arrived at their current locations without a circulatory system to transport RBCs. The few nucleated and enucleated RBCs in the KO demonstrated that early hematopoiesis, functional heartbeat, and vasculogenesis had occurred (Fig. 1f, 3a, 6)^30,31. These findings suggests that in the SAMD1 KO mice at least partially functional ECs differentiated, proliferated, connected, and organized into a functional vascular plexus during the early stage of embryogenesis. However, the E14.5 KO no longer has functional vasculature. At this stage, staining for EC markers appeared to mark only necrotic cells (Figs. 3,4,5), and RBCs were seen only in small clusters and scattered broken lines suggesting that functional blood vessels had degraded. Similarly, RBCs, and the lack thereof, in the E14.5 KO yolk sac, placenta, and umbilical cord seem to mark failed vessels resembling those seen in the embryo. This should not be surprising, since vessel development is similar in embryonic and extra-embryonic tissue^32–34. Given that that by approximately E12.5 the definitive placenta is mature³⁵ and the E12.5 embryos and extra-embryonic tissue appeared grossly normal, we conclude that embryonic lethality later than E12.5 is unlikely to have placental defects as a primary cause. We hypothesize therefore that lack of SAMD1 causes roughly simultaneous changes in the embryo, the yolk sac and the placental vasculature. Resorption is very rapid after cessation of heartbeat³⁶, implying that death in the E14.5 mouse occurred at approximately E13.5.

The presence of SAMD1 mRNA in the entire WT embryo and yolk sack (Fig. 1g, h, Supplementary Fig. 1) suggests that SAMD1 is relevant beyond vessel development. Thus, some SAMD1 KO features may not be related to failure of vascular development. The high expression of SAMD1 in the head, as well the consequences of SAMD1 deletion on neuronal differentiation processes (Fig. 7), supports a role of SAMD1 during head and neuronal development (Fig. 1f). Given that undirected ES cell differentiation in the absence of SAMD1 influences multiple distinct pathways⁴, it is likely that SAMD1 is involved in additional molecular processes, which await further clarification. Thus, experiments that address the role of SAMD1 during other differentiation processes will be important to clarify the relevance of SAMD1 for distinct lineages. Generating conditional SAMD1 knockout mice may be a suitable strategy to assess the role of SAMD1 during the development of specific tissues and organs. Useful data might also be gathered from studies of SAMD1 KO mice that were embryonic lethal later in development.

Mutations that result in absence of a gene often do not have uniform effects across a population. The mechanisms can be environmental, epigenetic, synergistic, and multigenic³⁷. Variable expression and penetrance are likely explanations for the survival until as late as E18, of the few KOs that had heartbeats past E14.5 (Supplementary Table S1), and for the approximately 30% postnatal HET mortality by 3 weeks of age (Supplementary Table S2 and S3). It is possible that compensating proteins, which typically have domain similarities with a knocked-out protein, allowed at least partial rescue of the developing embryo³⁸. The E15.5 KO shown in Fig. 1f may be a somewhat different phenotype from the E14.5. The lack of blood vessels appears similar, but the absence of skull vault and the exencephaly in the E15.5 KO is different, and might have been caused by an “open” neural tube defect³⁹.

Heterozygous SAMD1 KO mice had substantially increased aldosterone, corticosterone, angiotensin II, and testosterone levels (Fig. 8i). Steroidogenesis and transformation of cholesterol to oxysterols occurs in many cell types²⁹. It is currently unclear whether SAMD1 directly regulates hormone production pathways in cells or whether these changes are caused by compensatory effects, thus requiring further investigation. The HETs also showed reduced body weight and adiposity (Fig. 8a-c) that may be related to steroidal changes

This investigation aimed to present the first description of the effect of SAMD1 ablation on mouse embryonic development. Based on the E14.5 and E15.5 phenotypes, we focused on features related to vasculogenesis, angiogenesis and neuronal differentiation. The study is therefore limited to a restricted time window of embryonic development and does not provide deeper insights into the role of SAMD1 during earlier or later timepoints. Given the use of a limited set of markers and probes, only differentiated ECs and epithelial cells were unambiguously identified. Therefore, it is not determined whether failure of the early vascular plexus is due to failure of migration, proliferation, differentiation, communication, or connection of necessary progenitor cells or of an unknown mechanism. Furthermore, we investigated only a restricted subset of animals and organs for the KO mice and only selected measures for the heterozygous mice. Thus, it is likely that other phenotypes or defects occur in the absence of SAMD1. Additionally, this study does not address the cellular and molecular mechanisms by which the absence of SAMD1 leads to the observed phenotypes. Given that SAMD1 is expressed in all tissues and organs (Fig. 1g, h), more research will be necessary to elucidate the role of SAMD1 in specific cellular contexts.

This is the first report on ablation of SAMD1 in mice. We observed embryonic lethality, which may be secondary to the myriad of effects stemming from the lack of appropriate vascular development. Given the wide expression of SAMD1 in most organs, numerous other processes may also be affected, potentially explaining the complex phenotype, both in the homozygous and the heterozygous mice.

Animal handling

Mice were bred, studied, and maintained in the Longwood Medical Research Center facility in accordance with guidelines of the Committee on Animals of the Harvard Medical School and those prepared by the Committee on Care and Use of Laboratory Animals of the Institute of Laboratory Resources, National Research Council [DHEW publication No. (NIH) FS-23]. The Pfizer Institutional Animal Care and Use Committee (IACUC) approved all of the animal procedures and protocols according to the criteria stated by the National Academy of Sciences National Research Council (NRC) publication 86 − 23, 1985. All methods were performed in accordance with the relevant guidelines and regulations, as well as in accordance with the ARRIVE guidelines.

SAMD1 gene inactivation

SAMD1 knockout mice were genetically engineered by Pfizer (Groton, CT). To generate the SAMD1 targeting vector, recombineering⁴⁰ was used to replace 2396 bp of the mouse SAMD1 (Accession ID D3YXK1) gene encompassing the 3’ 615 bp of exon 1, exons 2–4 and all of exon 5 except for the 3’ 83 bp with a neomycin phosphotransferase cassette in a 9433 bp genomic subclone obtained from a C57BL/6J BAC (RP23-128H6; Invitrogen). The vector was introduced into Bruce4 (mixed C57Bl/6N:C57BL/6J) mouse embryonic stem cells⁴¹ using a standard homologous recombination technique. Southern blot analysis was used to identify correctly targeted mouse embryonic stem cell clones. Chimeric mice were generated by injection of the targeted ES cells into Balb/c blastocysts. Chimeric mice were bred with C57Bl/6J mice to produce F1 heterozygotes. Germline transmission was confirmed by PCR analysis (SAMD1g1: 5’-CCAAACCCCTCTTCAGTTCA-3’; SAMD1g2: 5’-GCCGTAGCTATTCTGCCTCA-3’; dNEO2: 5’-ACATAGCGTTGGCTACCCGTGATA-3’). F1 heterozygous males and females were mated to produce F2 mice. The colony was maintained by intercrossing under specific pathogen-free conditions with unrestricted access to food and water.

RNA and cDNA measurements for mouse experiments

RNA and cDNA measurements were performed at the Brigham & Women’s Hospital (Boston, MA). Synthesis of cDNA (ThermoScript RT-PCR System (Invitrogen Cat 11146-016)). RT-PCR was performed in a MyiQ single-color real-time PCR system (Bio-Rad Laboratories, Inc., Hercules, CA). Total ribonucleic acid (RNA) from 250,000 cells was reverse transcribed by Superscript II (Invitrogen) according to the manufacturer's instructions. Quantitative PCR was performed with SYBR green PCR mix (primers for SAMD1 from QIAGEN N.V.) and analysis performed with StepOne Software (ThermoFisher, Applied Biosystems). Levels of mRNA were normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) mRNA levels. cDNAs were synthesized using a High-Capacity RNA-to-cDNA™ Kit (Thermo Fisher, Catalog No. 4387406). Total 2 µg RNA/100 µl cDNA synthesis reaction and 1 µl cDNA per TaqMan reaction. GAPDH was used as an endogenous control (Applied Biosystem 4352339E. Catalogue No. 43-523-39E).

Isolation of mouse embryonic fibroblasts

Mouse embryonic fibroblasts (MEFs) were isolated as described⁴². In short, embryos from WT, SAMD1+/−, and SAMD1−/− mice were isolated between E12.5 and E18.5. Internal organs, heads, tails, and limbs were removed from embryos prior to treatment with trypsin to dissociate the cells. Cells were seeded into T-75 cell culture dishes in 15 mL of MEF media: KO DMEM (Life Technologies, 10829-018), 15% FBS (Life Technologies), nonessential amino acids (Life Technologies 111140-050), 2 mM L-glutamine (Life Technologies, 25030-081), 0.1 mM β-mercaptoethanol (Sigma M7522), and 0.25 mg/ml gentamicin (Life Technologies, 15710-072).

Phenotypic studies

All behavioral, physical, hormonal, and response phenotype comparisons of SAMD1+/+ to SAMD1+/− were performed by Caliper Life Sciences (Xenogen Biosciences, Cranbury, NJ) based on the Phenotype Pfinder platform⁴³.

Immunohistochemistry

Antibodies against CD31 and VEGFR2 were purchased from Abcam (Cambridge, UK). Immunohistochemical staining procedures were performed by Pfizer (Groton, CT) and at the Brigham & Women’s Hospital (Boston, MA). E14.5 has been established as the optimal time point for mouse developmental disorder phenotype analyses¹⁹. In short, tissues were collected immediately after E14.5 embryo collection, fixed in 10% buffered formalin and embedded in paraffin. Four micrometer serial sections were cut and mounted on glass slides. One section from each lesion was stained with hematoxylin and eosin (H&E). Other slices were stained for CD31 (ab28364) or VEGFR2 (ab2349) with hematoxylin counterstaining following Abcam’s protocols.

Cell culture

E14 mESCs (E14TG2a) were provided from the lab of Jacqueline Mermoud (University of Marburg, Germany). SAMD1 KO E14 mESCs were established previously⁴. The WT and SAMD1 KO mESCs were cultured in Dulbecco’s modified Eagle’s medium (DMEM) and GlutaMAX (Gibco; 61965-026), 15% fetal calf serum (FCS) (Biochrom; S0115, Lot: 1247B), 1× nonessential amino acids (Gibco; 11140-035), 1× sodium pyruvate (Gibco; 11160-039), 1× penicillin/streptomycin (Gibco; 15140- 122), 10 µM β-mercaptoethanol (Gibco; 31350-010), and LIF (1000 U/ml; Millipore; ESG1107, lot: 3060038) on 0.2% gelatin-coated plates.

Neuronal differentiation of mouse ES cells – Embryoid body system

mESCs were plated at a density of 1x 10⁶ on 0.1% gelatin-coated and dried 100 mm cell culture dishes for 3 days in mESC medium, and the medium was changed every 24 h. To start neurodifferentiation, 1x10⁶ dissociated ES cells were incubated on nonadhesive bacterial dishes (100 mm) for 6–8 days in mESC medium without LIF, and the medium was changed every 2 d. Embroid bodies (EBs) were collected and redistributed to 4 wells of a 6-well plate containing 2 ml differentiation medium containing DMEM (Gibco; 11960-044), 20% FCS, 2 mM L-glutamine (Gibco; 25030-081), 1x penicillin/streptomycin, 1x nonessential amino acids, 50 µM β-mercaptoethanol, and 0.5 µM trans-retinoic acid (Millipore; 554720) and incubated for 2 days. EBs from two wells each were collected and redistributed to a 100 mm cell culture dish, coated with 20 µg/mL poly-L-ornithine hydrobromide in PBS (Sigma-Aldrich; P3655; 4 h to overnight at room temperature, rinsed twice with water) and incubated for 7 days in neurobasal medium (Gibco; 21103-049) with 1x L-glutamine, 1x penicillin/streptomycin, and 1x B-27 supplement (Gibco; 17504-044)⁴⁴.

The established neural precursor cells (NPCs) were cultured in Euromed-N medium (EuroClone; ECM0883L) supplemented with 1x N-2 (Gibco; 17502-048), 1x L-glutamine, 1x penicillin/streptomycin, 50 µg/ml BSA (Sigma-Aldrich; A9418), 20 µg/ml insulin (Merck; 11376497001), 10 ng/ml EGF (Peprotech; 315-09), and 10 ng/ml FGF (Peprotech; 100-18B) on 0.1% gelatin-coated and subsequently 5 µg/ml laminin-coated (Sigma-Aldrich; L2020) six-well tissue culture plates and passaged with Accutase (Sigma-Aldrich; A6964) (adapted from ⁴⁵).

To promote the neurogenic capacity of NPCs and allow the formation of mature neurons, WT and SAMD1 KO NPCs were seeded at high density (6x10⁵) into gelatinized (0.1% in H2O) and poly-L-ornithine hydrobromide-coated (20 µg/ml in PBS) six-well tissue culture plates for differentiation with N2B27 medium, consisting of DMEM/F12 (Gibco; 11320-033) and neurobasal medium (1:1), with 1x N-2, 1x B-27, 100 µM β-mercaptoethanol and the addition of 10 ng/ml FGF (Peprotech; 100-18B) (adapted from ⁴⁶). Analysis of the cells was performed with immunofluorescence staining and RT-qPCR at chosen time points.

RT-qPCR experiments

For RNA isolation, cells were cultivated on 6-well plates up to 80–100% confluency. RNA was prepared using the RNeasy Mini Kit (Qiagen; 74004) according to the manufacturer’s manual, including an on-column DNA digest (Qiagen; 79254). The PrimeScript RT Reagent Kit (TaKaRa; RR037A) was used to transcribe mRNA into cDNA according to the manufacturer’s manual. Samples were incubated for 30 min at 37°C followed by 5 min at 85°C to inactivate PrimeScript RT enzymes. Subsequently, cDNA was diluted 1:20 to be used in RT-qPCR. For analysis by real-time quantitative PCR, MyTaq Mix (Bioline; BIO-25041) was used. For gene expression analysis, values were normalized to mActb and mGapdh expression. The qPCR primers used are presented in Supplementary Table S4.

Antibodies for neuronal studies

The following commercial antibodies were used: H3K4me2 (Diagenode; C15410035), TUJ1/TUBB3 (BioLegend; 801201), and GFAP (Dako; Z0334). The antibody for SAMD1 is custom-made. It is directed against the SAM domain of human SAMD1 (amino acids 452 to 538) and was produced as described previously ⁴.

Proliferation Assay

To determine proliferation rates, cells were seeded on 6-well plates at a density of 1x10⁵ cells per well. Cell viability was determined at 1, 3, and 6 days after seeding using the MTT assay by adding 90 µl of 5 mg/ml thiazolyl blue ≥ 98% (Carl Roth; 4022) to each well. The medium was aspirated after 1 h, and stained cells were dissolved in 400 µl of lysis buffer (80% isopropanol, 10% 1 M HCl, 10% Triton X-100). Absorption was measured at 595 nm using a plate reader. All values were normalized to day 1 to compensate for variations in seeding density. The mean value of three biological replicates was determined.

Immunofluorescence staining

WT or SAMD1 KO mESCs were seeded on 0.2% gelatin-coated coverslips. WT or SAMD1 KO differentiated neural precursor and neural cells were seeded on 0.1% gelatin-coated and 20 µg/mL poly-L-ornithine hydrobromide-coated coverslips Cells were fixed with 4% formaldehyde (w/v), methanol-free (Thermo Fisher Scientific; PI28906), and subsequently permeabilized with wash buffer (0.5% Triton X-100 in PBS). Blocking was performed with 10% FCS in wash buffer. Primary antibodies were diluted 1:500 in blocking solution and incubated in a wet chamber overnight at 4°C. Three washing steps of the cells were performed before incubation with secondary antibodies, using Alexa Fluor 488 goat anti-rabbit IgG (H + L) and Alexa Fluor 546 goat anti-mouse IgG (H + L) (Thermo Fisher Scientific; A-11008 and A-11003), at a 1:2000 dilution each. Following three washing steps, the coverslips were mounted onto microscopy slides using VECTASHIELD® Antifade Mounting Medium with DAPI (Vector Laboratories; H-1200) and sealed. Microscopy was performed using a Leica DM5500 microscope, and data were analyzed using ImageJ (Fiji).

Immunofluorescence quantification

To determine the cellular fluorescence in microscopy images using ImageJ, cells of interest were outlined and measured for their area, integrated intensity and mean gray value. Adjacent background regions were measured for the same parameters to calculate the corrected total cell fluorescence (CTCF) according to the following formula: CTCF = integrated density – (area of selected cell x mean gray value of background reading). Average CTCF values and standard deviations for 5 cells each in 3 different fluorescence microscopy images per sample are presented in a graph.

Bioinformatics analysis

Gene expression data from mouse embryo tissues were obtained from ENCODE (https://www.encodeproject.org/, under “Mouse Development”)¹⁷ and from GSE45719¹⁶. GSEA was performed with standard settings⁴⁷ using previously published data (GSE144396)⁴.

Statistical Analysis

Quantitative results are expressed as mean ± standard deviation of the mean (SD). Deviation from the mendelian ratio was evaluated using Chi Square tests. The significance of differences between two independently measured groups were evaluated using a two-tailed unpaired t-test. Two-way ANOVA followed by Tukey’s postdoc test was applied for multiple group comparisons. Significance of the Lipoprint analysis was evaluated using a two-tailed Mann–Whitney U test. The p-values were calculated using the GraphPad Prism Software. The p-values of the GSEA analysis were obtained from the GSEA software⁴⁷. A value of p < 0.05 was considered statistically significant.

Acknowledgements

We thank Dr. Andrew Lichtman (Brigham & Women’s Hospital, Boston, MA) for guiding discussions, critical inputs, unwavering intellectual support, and for managing members of his lab throughout this research. We thank Dr. Margaret Tarrio (Brigham & Women’s Hospital, Boston, MA) for breeding, raising, performing microscopy and IHC studies on KO mice, as well as performing mouse treatment studies. We thank Dr. Bastian Stielow for creating the SAMD1 KO mouse embryonic stem cells and Iris Rohner for technical assistance.

Author Contributions

B.C., S.J.E., and R.L. wrote the manuscript. L.M.W. and R.L. performed and supervised, respectively, neuronal differentiation experiments with mESCs. S.J.E. was responsible for the generation, breeding, and phenotyping of the SAMD1 mouse line. T.R.S.O. was responsible for the histological and embryonic lethality analyses of the WT and KO embryos. P.B. performed histology and histological analyses of WT and KO embryos. R.A. performed oversight and leadership for the Pfizer research activity including planning, execution, and mentorship, and provided materials, reagents, laboratory samples, animals, instrumentation, computing resources, and other analysis tools. All authors reviewed the manuscript.

Data availability

The datasets generated during and/or analyzed during the current study are available from the corresponding authors on reasonable request. Gene expression data analyzed during the study are available from ENCODE¹⁷ (https://www.encodeproject.org/, under “Mouse Development”) and from the GEO repository (GSE45719¹⁶ and GSE144396⁴).

Funding

This study received funding from Atherex and Pfizer. The funders were not involved in the study design, collection, analysis, interpretation of data, the writing of this article or the decision to submit it for publication. R.L. acknowledges support from the German Research Foundation (DFG, TRR 81/3 –109546710).

Ethics declarations

Competing interests: Authors P. Bourassa and R. Aiello are currently employed by Cybrexa Therapeutics. Author S. J. Engle is currently employed by Biogen. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. Thus, all authors declare no competing interests.

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Ablation of SAMD1 in Mice Causes Failure of Embryonic Blood Vessel Maturation and Embryonic Lethality

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Abstract

Figures

Introduction

Results

Discussion

Methods

Animal handling

SAMD1 gene inactivation

RNA and cDNA measurements for mouse experiments

Isolation of mouse embryonic fibroblasts

Phenotypic studies

Immunohistochemistry

Cell culture

Neuronal differentiation of mouse ES cells – Embryoid body system

RT-qPCR experiments

Antibodies for neuronal studies

Proliferation Assay

Immunofluorescence staining

Immunofluorescence quantification

Bioinformatics analysis

Statistical Analysis

Declarations

Author Contributions

Data availability

Funding

Ethics declarations

References

Additional Declarations

Supplementary Files

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