SUMMARY
The lung microvasculature is essential for gas exchange and commonly considered homogeneous. We show that Vascular endothelial growth factor A (Vegfa) from the epithelium specifies a distinct endothelial cell (EC) population in the postnatal mouse lung. Vegfa is predominantly expressed by alveolar type 1 (AT1) cells and locally required to specify a subset of ECs. Single cell RNA-seq identified 15-20% lung ECs as transcriptionally distinct and marked by Carbonic anhydrase 4 (Car4), which are specifically lost upon epithelial Vegfa deletion. Car4 ECs, unlike bulk ECs, have extensive cellular projections and are separated from AT1 cells by a limited basement membrane without intervening pericytes. Without Car4 ECs, the alveolar space is aberrantly enlarged despite the normal appearance of myofibroblasts. Lung Car4 ECs and retina tip ECs have common and distinct transcriptional profiles. These findings support a signaling role of AT1 cells and shed light on alveologenesis.
INTRODUCTION
Endothelial cells (ECs) lining the blood vessels fulfill their transport function via region- and organ-specific specialization, such as the widely-known artery-capillary-vein relay and the non-leaky blood-brain barrier (Aird, 2007a, b; Potente and Makinen, 2017). Additional EC heterogeneity and plasticity are illustrated during development by the opposing duo of leading tip cells and trailing stalk cells within sprouting vessels, as well as the transition of ECs to hematopoietic, mesenchymal, and lymphatic lineages (Dejana et al., 2017; Gariano and Gardner, 2005). These functional and morphological differences in ECs are underlain by distinct gene expression profiles that have been extensively studied in the more tractable vasculature of the postnatal retina and, recently, have begun to be systematically tackled across organs using single cell RNA-seq (scRNA-seq) (Han et al., 2018; Sabbagh et al., 2018). An emerging theme for cell types that exist in multiple organs, as exemplified by macrophages (Lavin et al., 2014), is that they are endowed with organ-specific molecular signatures.
The pulmonary circulation consists of arterial and venous trees that parallel the branched airways and alveolar ducts and connect distally via a dense network of capillaries covering the gas-exchange alveoli – a high level of spatial coordination that presumably requires precise epithelial-endothelial crosstalk (Morrisey and Hogan, 2010). Although differences between lung macro- and micro-vasculature as well as lung-specific EC gene expression have been noted (Sabbagh et al., 2018; Stevens et al., 2008), the molecular, cellular, and genetic basis of these differences are poorly understood, especially in vivo (Durr et al., 2004). Deciphering lung EC heterogeneity and its developmental origin is also critical to our understanding of bronchopulmonary dysplasia, a severe lung disease often associated with premature birth and characterized by simplified alveoli and dysmorphic vasculature (Thebaud and Abman, 2007).
Our published work shows that (1) the lung capillaries are embedded within grooves of folded alveolar type 1 (AT1) cells, which constitute >95% of the alveolar epithelium; (2) developing AT1 cells, instead of alveolar type 2 (AT2) cells, express a potent angiogenic factor Vascular endothelial growth factor A (Vegfa); and (3) genetically blocking AT1 cell development decreases alveolar angiogenesis (Yang et al., 2016). These results indicate an instructive role of the alveolar epithelium for lung ECs. In this study, using scRNA-seq and single cell imaging, conditional and mosaic genetic models, and cross-organ comparisons, we found that epithelial-derived Vegfa is required to specify a transcriptionally-distinct lung EC population, which features net-like cellular extensions, embraces the epithelial contours, and promotes alveologenesis independent of myofibroblasts – contractile mesenchymal cells generally considered to drive alveologenesis (Bostrom et al., 1996).
RESULTS
Vegfa is predominantly expressed by AT1 cells and locally promotes alveolar angiogenesis
To confirm and extend our previous finding of Vegfa expression in developing AT1 cells (Yang et al., 2016), we immunostained developing and mature lungs carrying a nuclear LacZ knock-in reporter, VegfaLacZ (Miquerol et al., 1999). While scattered and at a low level throughout the embryonic lung, the LacZ reporter in postnatal lungs co-localized with nuclei that were positive for NK2 homeobox 1 (NKX2.1; a lung epithelial lineage factor (Minoo et al., 1999)) but were not outlined by cuboidal E-Cadherin (a cell junction protein) staining – a characteristic feature of AT2 cells – indicating that AT1 cells, instead of AT2 cells, express Vegfa (Fig. 1A).
To test the functional relevance of this AT1 specific Vegfa expression, we conditionally deleted Vegfa in AT1 cells using Aqp5Cre (Flodby et al., 2010), as validated by in situ hybridization probing for the deleted exon (Fig. S1A). The resulting mutant lungs had sparser alveolar vasculature without apparent changes in alpha-smooth muscle actin (SMA)-expressing myofibroblasts (Fig. 1B). This vascular phenotype was quantified as a persistent decrease in vessel volume and EC number (Fig. 1B, C). However, EC proliferation was not affected (Fig. 1B, C), suggesting that VEGFA does not function appreciably as a mitogen in the postnatal lung. Interestingly, the remaining vessels in the mutant were not randomly distributed: they failed to occupy the surface of alveolar “islands” that, in a control lung, contained grooves that were associated with myofibroblasts and thus considered secondary septation (Fig. 1B, D), raising the possibility that VEGFA is required for a specific subset of alveolar vessels.
The role of AT1-derived Vegfa was confirmed using another AT1 cell driver HopxCreER (Takeda et al., 2011; Yang et al., 2016). A single dose of the chemical inducer, tamoxifen, resulted in mosaic recombination in AT1 cells, as indicated by the juxtaposition of recombined GFP+ and unrecombined GFP- AT1 cells using a Cre reporter RosamTmG (Muzumdar et al., 2007)(Fig. 2A). As expected for the control lung, the alveolar islands, as defined in Fig. S1B, were invariably covered with vessels with no difference between GFP and non-GFP islands (Fig. 2). In contrast, up to 50% of the alveolar islands in the Vegfa mutant had no vessels – a phenotype more frequently observed for the GFP islands, presumably due to their higher likelihood of recombining both Vegfa alleles (Fig. 2). Notably, islands with and without vessel coverage were frequently found to be juxtaposed (Fig. 2 and Fig. S1B), indicating that AT1 derived VEGFA functions locally to promote vessel formation.
To exclude other functional sources of VEGFA, we conditionally deleted Vegfa from AT2 cells and ECs using SftpcCreER and Cdh5-CreER, respectively (Barkauskas et al., 2013; Wang et al., 2010). As expected from the lack of appreciable expression (Fig. 1A), these mutants had normal vasculature (Fig. S1C, D).
Lastly, we noticed that the AT1-specific Vegfa mutant had normal vasculature at postnatal day (P) 2 (Fig. S2A), 5 days after the initiation of AT1 cell differentiation at embryonic day (E) 17 (Desai et al., 2014; Li et al., 2018a; Yang et al., 2016). This was likely because Cre recombination occurred after commitment to the AT1 identity – a requirement for a cell-specific Cre driver – such that enough VEGFA protein had accumulated and perdured after DNA deletion. To test this and to achieve more efficient deletion, we resorted to ShhCre, which targets the entire epithelium during lung specification (Harris et al., 2006). Despite such early targeting, and different from reported results in another epithelial Vegfa deletion model (Yamamoto et al., 2007), branching morphogenesis was unaffected and vascular defects were only observed after E17, concomitant with AT1 cell differentiation (Fig. S2B, C), suggesting that early embryonic lung angiogenesis is supported by non-epithelial sources of Vegfa or Vegfa-independent mechanisms. The pan-epithelium Vegfa mutant recapitulated the AT1 specific Vegfa mutant phenotypes, including a decrease in vessel volume and EC number without affecting proliferation, as well as the failure to cover alveolar islands where secondary septation normally occurred (Fig. 3A and 3B). Taking into account all the cell type-specific Vegfa mutant models, we concluded that AT1 derived Vegfa is required locally for alveolar angiogenesis and possibly regulates a specific EC population in the developing lung.
Single cell RNA-seq identifies a molecularly distinct lung EC population
To understand the molecular basis of the Vegfa mutant phenotype, we optimized a cell dissociation and sorting protocol using lungs with genetically-labeled fluorescent ECs and found that adding a CD45 (an immune cell marker) negative selection step allowed better separation of ICAM2 (or CD31; both EC markers) positive and negative cells and that ICAM2 selection was consistent with, but more robust than, the more commonly-used CD31 selection (Fig. S3). Bulk RNA-seq comparison of purified ECs from control and epithelial Vegfa mutant lungs revealed downregulation of genes that were, intriguingly, recently identified as markers of sprouting tip ECs, such as Signal-regulatory protein alpha (Sirpa) and Carbohydrate sulfotransferase 1 (Chst1) (Fig. 3C and Table S1) (Sabbagh et al., 2018). This, together with the established role of Vegfa in inducing retinal tip ECs (Gariano and Gardner, 2005) and the non-random distribution of remaining ECs in our Vegfa mutant lungs, led us to hypothesize that Vegfa specifies a subset of ECs in the lung that are analogous to tip ECs in the retina.
To examine such potential lung EC heterogeneity, we performed scRNA-seq on 4,857 sorted P14 lung ECs (out of 5,175 cells) using 10x Genomics. Intriguingly, 14% of the sequenced ECs comprised a transcriptionally distinct cluster, which we named Car4 EC because Carbonic anhydrase 4 (Car4) was its most specific marker (Fig. 4). All remaining ECs expressed Plasmalemma vesicle associated protein (Plvap) and included two transcriptionally distinct clusters – non-capillary ECs that expressed Von Willebrand factor (Vwf; thus named Vwf EC; 8.5% of all ECs) and lymphatic ECs that expressed Prospero homeobox 1 (Prox1; 1.5% of all ECs) (Fig. 4A, B), both of which were confirmed to be spatially distinct from capillary ECs (Fig. S4). The remainder of the Plvap-expressing ECs (76% of all ECs) did not display a strong gene signature (Fig. 4C) – possibly because they were not as specialized as Car4 ECs, Vwf ECs, and lymphatic ECs – but were named Plvap EC for both simplicity and the availability of a PLVAP antibody for tissue localization. We note that Car4, Plvap, Vwf, and Prox1 were used as population-specific markers, but not investigated for their functions in this study.
We further investigated the localization of Car4 and Plvap ECs by immunostaining in conjunction with a pan-EC nuclear marker ETS transcription factor (ERG) (Fish et al., 2017; Shah et al., 2016). Although predominantly on the cell membrane, both CAR4 and PLVAP also accumulated around the nucleus, allowing assignment of each ERG nucleus to the Car4 or Plvap EC population (Fig. 5A). Interestingly, CAR4 ECs covered the aforementioned alveolar islands that were undergoing secondary septation, whereas PLVAP ECs surrounded those islands, a distribution reminiscent of the remaining vessels in the Vegfa mutants (comparing Fig. 5A with Fig. 1B and 3A). This spatial difference in CAR4 and PLVAP staining was also evident for the corresponding nuclei (Fig. 5A). Furthermore, although CAR4 staining was abundant, the number of CAR4+ cells was under-represented (Fig. 4A, 5A), a discrepancy between cell number and vessel contribution suggesting that CAR4 ECs are disproportionally larger.
Single cell imaging revealed extended Car4 EC morphology
Unlike the retina vasculature, which can be visualized in 2D after flat-mounting the tissue, lung ECs reside in thin tubes winding through a three-dimensional alveolar structure, making it challenging to examine their cell morphology. To overcome this, we adopted the sparse cell labeling method that we have used to study the similarly-complex AT1 cells (Yang et al., 2016). Specifically, we used a pan-endothelial inducible driver Cdh5-CreER (Wang et al., 2010), combined with an optimized, limited dose of tamoxifen, to sparsely label ECs so that individual ECs could be readily demarcated. As lungs were harvested within 24 hr after tamoxifen administration, labeled cells were not intended to be clonally related. We also used a strong reporter RosatdT (Madisen et al., 2010) to fill the entire cell body to (1) visualize the nucleus to confirm the intended single cell labeling and (2) colocalize with the membrane marker CAR4 to identify EC populations. Labeled cells were identified as Car4 ECs if they satisfied two criteria: (1) nuclear tdT completely overlapped with perinuclear CAR4 staining; (2) cytoplasmic tdT completely aligned with membrane CAR4 staining. Remarkably, in comparison to the bulk Plvap ECs, Car4 ECs were highly branched, often contributing to 5-10 vessel segments, and also much larger, as quantified by measuring the cell perimeter (Fig. 5B, C). Both Car4 and Plvap ECs had distinct morphology from the non-capillary Vwf ECs, which were much elongated along the direction of blood flow (Fig. S5A).
Closer examination of RosatdT labeled ECs in relation to neighboring Car4 ECs, an apical lumen marker Intercellular adhesion molecule 2 (ICAM2), and a cell adherens junction protein Cadherin 5 (CDH5) showed that (1) the sprawling labeled CAR4+ ECs did not exclude other ERG nuclei within those vessel segments; (2) the labeled CAR4- ECs could be juxtaposed and share a common lumen with CAR4+ ECs, the latter being closer to the air space; (3) although the boundary of labeled cells coincided with CDH5, additional cell junctions were present within the labeled vessel segments (Fig. 5B and S5A). These observations suggested that individual lung capillaries are multi-cellular and can be comprised of both CAR4+ and CAR4- ECs.
Comparison of retina and lung ECs revealed common and organ-specific EC heterogeneity
The aforementioned similarity of Car4 ECs to tip ECs prompted us to examine tip EC-enriched genes in our distinct subsets of lung ECs. Intriguingly, Car4 ECs specifically expressed Apln as well as additional tip EC genes recently identified by scRNA-seq analysis of developing brain ECs (Sabbagh et al., 2018), such as Plaur, Serpine1, Sirpa, Piezo2, and Chst1 (Fig. S6A, C). Plvap ECs specifically expressed several stalk cell genes including Apelin receptor (Aplnr) and TEK receptor tyrosine kinase (Tek; also known as Tie2) (Fig. S6A). However, the analogy of Car4/tip ECs and Plvap/stalk ECs did not extend to other known tip and stalk EC markers, as exemplified by Esm1 and Dll4 for tip ECs and Hes1 and Flt1 for stalk ECs (Blanco and Gerhardt, 2013) (Fig. S6A, B, C).
We further compared retina and lung ECs by immunostaining for ESM1, CAR4, and DLL4 (Fig. S6D). As reported (Rocha et al., 2014), ESM1 was restricted to tip ECs in the peripheral retina and excluded from mature vessels in the more central region. Interestingly, ESM1 was expressed in sporadic ECs near the lobe edge in embryonic lungs, raising the possibility that the lobe edge represents a growing front similar to the peripheral retina. However, this edge-specific expression was lost postnatally and ESM1 was enriched in ECs in a transition zone between capillaries and non-capillaries (Fig. S6D). In contrast, CAR4 was not detected in either the tip or stalk ECs in the retina, whereas its expression in the lung initiated at E19, concomitant with AT1 cell differentiation (Fig. S6D). Lastly, although Dll4 regulates tip ECs (Blanco and Gerhardt, 2013), its protein was only slightly enriched in tip ECs and, in the central region, were more evident in arteries than veins. Similarly, DLL4 was widely expressed in embryonic lungs, but became more concentrated in a cord-like pattern postnatally (Fig. S6D). Such frequency and distribution of ESM1 and DLL4-expressing cells were also reflected in scRNA-seq (Fig. S6A, B). These data suggested that there are additional layers of EC heterogeneity in the lung that do not correspond to our four EC subpopulations.
Next, we examined the morphology of retina ECs in comparison to that of lung ECs using our sparse cell labeling method (Fig. S5B). In the central retina, ECs either were elongated along large vessels, similar to those in non-capillaries of the lung, or had a limited number of projections, similar to the lung Plvap ECs. In the peripheral retina, tip ECs displayed their characteristic blind-end filopodia, which was never found in the lung. Conversely, the expansive net-like morphology of Car4 ECs of the lung was also never found in the retina (Fig. S5B).
Among the tip EC genes shared by the lung Car4 ECs, we focused on Apln because of its reported role in sprouting angiogenesis (del Toro et al., 2010) and because its receptor Aplnr was specific to Plvap ECs (Fig. 4B). Using an AplnCreER knock-in, knock-out allele (Liu et al., 2015), we found that Apln underwent X-inactivation and, when tested with a RosaL10GFP reporter, recombination was variable and enriched in but not specific to Car4 ECs (Fig. S7A, B). In addition, scRNA-seq comparison of sorted lung ECs from mutant (AplnCreER/Y) and littermate control males showed, as expected, a complete loss of Apln specifically in Car4 ECs, but no difference in either the number or gene expression of Car4 and Plvap ECs (Fig. S7C). Therefore, despite their intriguing expression patterns, the functional relevance of Apln and Aplnr in the lung is uncertain.
Taken together, the cross-organ comparison identified similarities and differences in EC gene expression and cell morphology, suggesting a common core vascular pathway superimposed with organ-specific adaptation.
CAR4 ECs are specifically lost upon epithelial Vegfa deletion
Given their similarity to the retina tip ECs, which require Vegfa (Gerhardt et al., 2003), we asked if Car4 ECs of the lung also depended on Vegfa. To test this, we performed scRNA-seq on sorted ECs from control and epithelial Vegfa mutant lungs. Remarkably, the mutant lung specifically and completely lost the Car4 EC population, which represented 18% of all ECs in the control lung (Fig. 6A). In comparison, Plvap, Vwf, and lymphatic ECs were unaffected and proliferating ECs, marked by Mki67 and frequent in early postnatal lungs, were also unaffected, consistent with our immunostaining-based quantification (Fig. 6A and 3B). This specific loss of Car4 ECs was confirmed by immunostaining showing that, in the mutant lung, CAR4 was rarely detected in the remaining vessels but unaffected in alveolar macrophages where CAR4 was normally present (Fig. 6A).
The specific loss of Car4 ECs in the Vegfa mutant lung allowed us to examine Car4 ECs at a higher resolution on sections and in the context of nearby non-ECs (Fig. 6B). Specifically, we used the pan-EC apical membrane marker ICAM2 to label the vessel lumen, which was mostly collapsed for capillaries, and used Aquaporin 5 (AQP5), Collagen type IV (COL4), and Chondroitin sulfate proteoglycan 4 (CSPG4; also known as NG2) to label the AT1 cell membrane, basement membrane, and pericyte membrane, respectively. We found that CAR4+ ECs – those that expressed CAR4 in the control but were absent in the Vegfa mutant – abutted the AT1 cell membrane and shared a thin basement membrane with the epithelium without intervening pericytes (Fig. 6B). In contrast, CAR4- ECs – those that did not express CAR4 in the control or comprised all remaining vessels in the mutant – were away from AT1 cell membrane, underlain with a thicker basement membrane, and surrounded by pericyte processes (Fig. 6B). For vessels abutting the AT1 cell membrane, the side further away from the air space could consist of CAR4- ECs, as described in Fig. 4B, and were found to be covered by a thicker basement membrane and pericyte processes (Fig. 6B and the diagram herein). The CAR4- ECs in such hybrid vessels, as predicted by the lack of Car4 EC-containing vessels in the mutant, did not occupy the aforementioned secondary septae of the alveolar islands by themselves and depended on CAR4+ ECs for such locations (Fig. 3A, 6A), reminiscent of stalk ECs depending on tip ECs to spread to avascular regions. Taken together, these data suggested that Car4 ECs are induced by epithelial VEGFA, locate closest to the AT1 cell surface, and orchestrate the distribution of capillaries to secondary septae, as diagramed in Fig. 6B.
CAR4 ECs contribute to alveolar morphogenesis independent of myofibroblasts
The CAR4+ vessels covering alveolar islands undergoing secondary septation and their specific absence in the Vegfa mutant (Fig. 1B, 3A and 6A) led us to examine the role of Car4 ECs in alveolar morphogenesis. Indeed, without the CAR4+ vessels coursing through the previously described grooves of the folding AT1 cells, the AT1 surface was smoother in the mutant (Fig. 7A). Consistent with a failure in secondary septation, the air space in the mutant lung was aberrantly enlarged with fewer recognizable alveoli (Fig. 7A), as quantified using a mean linear intercept method (Liu et al., 2017) as well as a D2 method that measured area-weighted air space diameters and bypassed the difficulty in distinguishing alveolar ducts and alveoli in the mouse lung (Jacob et al., 2009; Massaro and Massaro, 1996; Parameswaran et al., 2006). Notably, such defective alveologenesis occurred despite the normal specification and localization of myofibroblasts, a widely-accepted driver of secondary septation (Bostrom et al., 1996) (Fig. 7B).
We further examined the mesenchymal lineage using a 4-marker panel: Platelet derived growth factor receptor, alpha polypeptide (PDGFRA), SMA, Transgelin (TAGLN; also known as SM22), and Platelet derived growth factor receptor, beta polypeptide (PDGFRB) (Fig. 7B). The peri-nuclear accumulation of PDGFRA, PDGFRB, and TAGLN allowed colocalization and assignment of cell types, while the commonly used myofibroblast marker SMA was not useful for such application but largely matched the cytoplasmic TAGLN staining. In the control lung, PDGFRB-expressing pericytes were distinct from PDGFRA cells, a subset of which expressed TAGLN and SMA and were thereby considered myofibroblasts. In the Vegfa mutant lung, there were fewer PDGFRB pericytes presumably as a result of loss of CAR4+ capillaries, but the PDGFRA/TALGN/SMA-expressing myofibroblasts were unaffected (Fig. 7B). These data are consistent with the notion that Car4 ECs, possibly together with the associated pericytes but independent of myofibroblasts, are required for alveolar morphogenesis.
DISCUSSION
In this study, we show that the pulmonary microvasculature is heterogeneous and harbors a transcriptionally distinct EC population that is defined by CAR4 expression and specifically requires epithelial derived Vegfa, which is predominantly supplied by AT1 cells. These Car4 ECs feature an extended net-like morphology, situate seamlessly over the alveolar epithelium and specifically in regions undergoing secondary septation, and are required for alveolar morphogenesis independent of myofibroblasts (Fig. 7C). This work opens a new avenue of vascular research and has implications in alveolar development, physiology, and pathogenesis.
Our study reveals remarkable parallels between retinal tip ECs and lung Car4 ECs: (1) retinal astrocytes express Vegfa and provide the scaffold for tip ECs (Gariano and Gardner, 2005), whereas lung AT1 cells express Vegfa and provide the surface for Car4 ECs; (2) VEGFA in the retina induces tip ECs with characteristic filopodia and tip EC genes, whereas VEGFA in the lung induces Car4 ECs with characteristic net-like morphology and a subset of tip EC genes, such as Apln. However, there are also substantial differences: (1) the retina vasculature undergoes sprouting angiogenesis to spread from the central vascularized region to the peripheral avascular region in response to a hypoxia-induced VEGFA gradient, whereas the postnatal lung vasculature is surrounded by inhaled oxygen, always covers the alveolar epithelium as a dense net, and is believed to undergo intussusceptive angiogenesis (Burri et al., 2004) to match the expansion of the epithelial surface; (2) tip ECs disappear after development, but Car4 ECs persist even in the mature lung (Fig. S7C). Future mechanistic dissection of the common and distinct signaling events downstream of Vegfa in the two organs should shed light on the poorly understood process of intussusceptive angiogenesis and may possibly establish lung Car4 ECs as a novel model for vascular biology. More broadly, understanding organ-specific function of Vegfa may pave the way for targeted anti-VEGF therapy (Meadows and Hurwitz, 2012).
Alveologenesis, formation of alveoli, divides primary alveolar sacs – resulting from expansion of embryonic branch tips – into mature alveoli via the process of secondary septation (Yang and Chen, 2014). These secondary septae are marked by myofibroblasts, SMA-expressing contractile mesenchymal cells. Notably, we previously showed that such SMA-marked septae are the grooves of folded AT1 cells and coincide with capillaries that, as found in this study, are composed of Car4 ECs. Furthermore, although myofibroblasts are required for secondary septation in response to PDGFA signaling (Bostrom et al., 1996; Li et al., 2018b), our Vegfa mutant is missing Car4 ECs without affecting myofibroblasts or PDGFRA expression and yet displays aberrantly enlarged alveoli – consistent with a failure in secondary septation. This result raises the possibility that the force-generating myofibroblasts initiate secondary septation and AT1 cell folding, which is stabilized and maintained by CAR4+ vessels or their associated pericytes (Kato et al., 2018). This possibility is also consistent with the observation that myofibroblasts disappear or adopt other cell fates after the lung matures, whereas the vasculature persists (Li et al., 2018b; Yang et al., 2016). Future work will examine Car4 ECs in mutants that directly affect myofibroblasts and examine both myofibroblasts and Car4 ECs in experimental BPD models and patients with BPD, which is characterized by defective alveologenesis (Abman, 2001).
Car4 ECs are likely to have additional functions. First, compared to other ECs, Car4 ECs co-develop with the gas-exchanging AT1 cells, have a larger surface area, and are located closest to the alveolar epithelium, separated by a thinner basement membrane without intervening pericytes – all features suggesting a high efficiency in gas exchange. Testing this would require monitoring gas exchange in Car4+ versus Car4- capillaries because conventional blood gas measurement of the Vegfa mutant will not distinguish a general loss of capillaries from a specific loss of high-efficiency Car4+ capillaries. Second, Car4 ECs specifically occupy the secondary septae and are required for alveolar morphogenesis, suggesting a structural role during lung development. Third, Car4 ECs specifically express secreted ligands including Apln and Kitl while the corresponding receptors, Aplnr and Kit, are expressed by Plvap ECs, suggesting possible signaling roles of Car4 ECs toward other ECs. However, our examination of the Apln mutant does not revealed any vascular phenotype. Future studies should focus on Car4 EC-specific genes, including CAR4 itself, which intriguingly is an enzyme involved in carbon dioxide formation (Crandall and O’Brasky, 1978; Fleming et al., 1993). Finally, the persistence of Car4 ECs in the mature lung calls for a better understanding of their role in homeostasis and injury-repair.
AUTHOR CONTRIBUTIONS
LVE, JDW, and JC designed research; LVE, MPC, VH, EJO, and JC performed research; PF, EDC, and ZB provided the Aqp5Cre mice; BZ provided the AplnCreER mice; LVE, JDW, and JC wrote the paper; all authors read and approved the paper.
DECLARATION OF INTERESTS
The authors declare no competing interests.
METHODS
Mice (Mus musculus)
The following mouse strains were used: VegfaLacZ (Miquerol et al., 1999), VegfaCKO (also called VEGF-LoxP) (Gerber et al., 1999), Aqp5Cre (Flodby et al., 2010), HopxCreER (Takeda et al., 2011), SftpcCreER (Barkauskas et al., 2013), Cdh5-CreER (Wang et al., 2010), AplnCreER (Liu et al., 2015), RosamTmG (Muzumdar et al., 2007), RosatdT (Madisen et al., 2010), RosaL10GFP (Liu et al., 2014). The day of observing a vaginal plug was designated as E1. To induce Cre recombination, tamoxifen (T5648, Sigma) dissolved in corn oil (C8267, Sigma) was injected intraperitoneally. The tamoxifen dosage used is specified in the figure legends. Outliers were excluded only if there were technical errors, such as failed immunostaining. The number of control-mutant pairs and sections analyzed is stated in the figure legends. Unless specified, mice of both genders were used. Investigators were not blind to the genotypes. Control and mutant samples were processed in the same tube or block to minimize experimental variation. No power analysis was used to determine the sample size. All animal experiments were approved by the Institutional Animal Care and Use Committee at Texas A&M Health Science Center Institute of Biosciences and Technology and MD Anderson Cancer Center.
Antibodies
The following antibodies were used: rabbit anti-Aquaporin 5 (AQP5, 1:2500, ab78486, Abcam), goat anti-Carbonic anhydrase IV (CAR4, 1:500, AF2414, R&D), BV786 rat anti-CD31 (1:250, 740870, BD Biosciences), PE/Cy7 rat anti-CD45 (1:250, 103114, BioLegend), mouse anti-Claudin 5 (Cldn5, 1:500, Invitrogen, 352588), rabbit anti-collagen IV (COL4, 1:2500, LSL-LB-1403, CosmoBioUSA), goat anti-Delta like canonical Notch ligand 4 (DLL4, 1:250, AF1389, R&D), Alexa Fluor 488 rat anti-CD324 (ECAD, 1:500, 53-3249-80, eBioscience), goat anti-Endothelial cell specific molecule 1 (ESM1, 1:500, AF1999, R&D), rabbit anti-Avian erythroblastosis virus E-26 (v-ets) oncogene related (ERG, 1:5000, ab92513, Abcam), goat anti-Vegfr3/Flt4 (1:1000, R&D, AF743), chicken anti-beta Galactosidase (LacZ, 1:500, Ab9361, Abcam), chicken anti-Green fluorescent protein (GFP, 1:5000, AB13970, Abcam), Alexa Fluor 647 rat anti-Intercellular adhesion molecule 2 (ICAM2, 1:500, A15452, ThermoFisher), rat anti-Intercellular adhesion molecule 2 (ICAM2, 1:2500, 16-1021-82, eBioscience), goat anti-Intercellular adhesion molecule 2 (ICAM2, 1:500, AF774, R&D systems), eFluor 570 rat anti-Ki67 (1:500, 41-5698-82, eBioscience), rabbit anti-Ki67 (1:1000, RM9106S0, ThermoFisher), rabbit anti-Chondroitin sulfate proteoglycan 4 (CSPG4, 1:1000, AB5320, Millipore), rabbit anti-NK2 Homeobox 1 (NKX2.1, 1:1000, sc-13040, Santa Cruz), rat anti-Platelet derived growth factor receptor alpha (PDGFRA, 1:1000, 14-1401-82, eBioscience), goat anti-Platelet derived growth factor receptor beta (PDGFRB, 1:1000, AF1042, R&D systems), rat anti-Plasmalemma vesicle associated protein (PLVAP, 1:125, 553849, BD Biosciences), rabbit anti-Prospero Homeobox 1 (PROX1, 1:250, 11-002, AngioBio), rat anti-Advanced glycosylation end-product specific receptor (RAGE, 1:1000, MAB1179, R&D systems), rabbit anti-Red fluorescent protein (RFP, 1:1000, 600-401-379, Rockland), Cy3-conjugated mouse anti-alpha-Smooth muscle actin (SMA, 1:1000, C6198, Sigma), rabbit anti-SM22 (TAGLN, 1:2500, Abcam, ab14106), Alexa Fluor 647 rat anti-Vascular endothelial cadherin (VECAD/CDH5, 1:250, 562242, BD Biosciences), rabbit anti-Von Willebrand Factor (VWF, 1:2500, Abcam, ab6994).
Section immunostaining
Postnatal lungs were inflation-harvested as described with minor modifications (Yang et al., 2016). Briefly, mice were anaesthetized with Avertin (T48402, Sigma) and perfused through the right ventricle with phosphate-buffered saline (PBS, pH 7.4). The trachea was cannulated and the lung was inflated with 0.5% paraformaldehyde (PFA; P6148, Sigma) in PBS at 25 cm H2O pressure, submersion fixed in 0.5% PFA at room temperature for 4-6 hr, and washed in PBS at 4 °C overnight. Section immunostaining was performed following published protocols with minor modifications (Alanis et al., 2014; Chang et al., 2013). Fixed lung lobes were cryoprotected in 20% sucrose in PBS containing 10% optimal cutting temperature compound (OCT; 4583, Tissue-Tek) at 4°C overnight and then embedded in OCT. OCT sections at 10 um thickness were blocked in PBS with 0.3% Triton X-100 and 5% normal donkey serum (017-000-121, Jackson ImmunoResearch) and then incubated with primary antibodies diluted in PBS with 0.3% Triton X-100 in a humidified chamber at 4 °C overnight. Sections were washed with PBS in a coplin jar for 1 hr and incubated with donkey secondary antibodies (Jackson ImmunoResearch) and 4’,6-diamidino-2-phenylindole (DAPI) diluted in PBS with 0.3% Triton X-100 at room temperature for 1 hr. After another 1 hr wash with PBS, sections were mounted with Aquamount (18606, Polysciences) and imaged on a confocal microscope (A1plus, Nikon).
Wholemount immunostaining
This was performed following published protocols with minor modifications (Yang et al., 2016). In brief, ∼3 mm wide strips from the edge of the cranial or left lobes of postnatal lungs or whole lobes of embryonic lungs were blocked with PBS with 0.3% Triton X-100 and 5% normal donkey serum (017-000-121, Jackson ImmunoResearch) and then incubated with primary antibodies diluted in PBS with 0.3% Triton X-100 overnight at 4 °C. The next day, the strips were washed with PBS+1% Triton X-100+1% Tween-20 (PBSTT) on a rocker at room temperature for one hour, and the process was repeated three times. Secondary antibodies and DAPI were added and incubated overnight at 4 °C. On the third day, the strips were washed as described before with PBSTT and fixed with PBS with 2% PFA for at least 2 hr on a rocker. For tissues expressing green or red fluorescent protein, native fluorescence was quenched after immunostaining by overnight incubation with methanol containing 6% hydrogen peroxide (H1009, Sigma) at 4 °C. Finally, the strips were mounted on slides using Aquamount (18606, Polysciences) with the flat side facing the coverslip. Embryonic lungs were also immunostained as a whole and imaged with an optical projection tomography microscope (Bioptonics, UK) as published (Alanis et al., 2014; Chang et al., 2013). Enucleated eyes were fixed in 0.5% PFA for 3-6 hr at room temperature and then retinas were dissected free of retinal pigmented epithelium, lens, and hyaloid vessels, immunostained in the same tube with matching lung strips, and mounted with the vitreous side facing the coverslip. Z-stack images of 20-40 um thick at 1 um step size were taken from the top of the tissue to obtain an en face view.
Section in situ hybridization
Postnatal lungs were harvested and processed as described for immunostaining except 0.5% PFA was included for the sucrose/OCT overnight incubation to minimize RNA degradation. Colorimetric section in situ hybridization was carried out following published protocols (Alanis et al., 2014; Chang et al., 2013). The entire exon 3 of Vegfa was amplified with the following primers for probe: 5’-TGATCAAGTTCATGGATGTC-3’ and 5’-agcttataatacgactcactatagggCTGCATGGTGATGTTGCTCT-3’ (lower case indicates the T7 promoter sequence). Images were acquired on an upright Olympus BX60 microscope.
Vasculature analysis
Wholemount samples were imaged on a confocal microscope (A1plus, Nikon) using the 40x oil objective with a field size of 318 μm x 318 μm x 20 μm and a pixel dimension of 512 x 512 x 20. At least three confocal Z-stacks per lung were analyzed with Imaris software (Bitplane) to obtain the EC number (ERG) and the percentage of proliferating (KI67) ECs. Surface rendering ICAM2 staining was used to measure vessel volume using the automatic threshold and a cut-off of 50 voxels for both control and mutant lungs. For automatic analysis of CAR4 and PLVAP staining, ERG surface rendering was filtered by the mean intensity of CAR4 or PLVAP, which was then used to mask ERG staining. To measure the EC perimeter, the viewpoint was set for the largest projection area and then all visible cell projections were measured. Cells in contact were split midway between them.
D2 air space analysis and mean linear intercept analysis
The D2 index is an area-weighted alveolar diameter that takes into account the heterogeneous distribution of airspace sizes in the mouse lung (Jacob et al., 2009; Parameswaran et al., 2006). In this study, the D2 was measured and calculated on 5 um-thick frozen lung sections stained with hematoxylin and eosin (H&E). For each mouse, three images were acquired on an upright Olympus BX60 microscope with a 10x objective. Airway and main vessels that could not be avoided during imaging were filled in manually using ImageJ prior to analysis, which was performed using a 225-pixel intensity threshold and a 400-pixel area size cut-off. The same images were used to quantify the mean linear intercept with Photoshop based on a published protocol (Liu et al., 2017). Two horizontal and two vertical gridlines were drawn evenly-spaced on each picture. The distance from one alveolar wall to the next along the gridlines was measured with the Photoshop ruler tool. Airways and main vessels were excluded, as well as alveoli where one wall was not visible in the image. At least 32 intercepts per image and 3 images per mouse were measured.
Cell dissociation and FACS
Postnatal mouse lungs were dissected in PBS, minced into pieces with forceps and digested in RPMI (ThermoFisher, 11875093) with 2 mg/mL Collagenase Type I (Worthington, CLS-1, LS004197), 2 mg/mL mL Elastase (Worthington, ESL, LS002294), and 0.5 mg/mL DNase I (Worthington, D, LS002007) for 30 min at 37 °C. The tissue was mechanically triturated after 15 min of digestion. Fetal bovine serum (FBS, Invitrogen, 10082-139) was added to a final concentration of 20% and the tissue was triturated until homogenous. The sample was transferred to the cold room and kept on ice, filtered with a 70 μm cell strainer (Falcon, 352350), and spun down at 5000 rpm for 1 min. The cells were resuspended in red blood cell lysis buffer (15 mM NH4Cl, 12 mM NaHCO3, 0.1 mM EDTA, pH 8.0) for 3 min, washed with RPMI with 10% FBS and filtered into a 5 ml glass tube with a cell strainer cap (Falcon, 352235). The cells were then incubated with CD45-PE/Cy7 (BioLegend, 103114), ICAM2-A647 (Invitrogen, A15452), CD31-BV786 (BD Biosciences, 740870) at a concentration of 1:250 for 30 minutes, spun down at 5000 rpm for 1 min, washed for 5 minutes and resuspended with RPMI with 10% FBS. The sample was refiltered and incubated with SYTOX Blue (Invitrogen, S34857), then sorted on a BD FACSAria Fusion Cell Sorter. After exclusion of dead cells, CD45 negative cells were selected and from those ICAM2 positive cells were collected.
RT-PCR and bulk RNA-seq
RNA was extracted from FACS-purified ECs using Trizol reagents (Invitrogen, 15596018) and the RNeasy Micro kit (Qiagen, 74004). 100 ng of RNA was used for RT-PCR with the SuperScript™ IV First-Strand Synthesis System (Invitrogen, 18091050). For bulk RNA-seq, 100-200 ng total RNA was used to prepare an RNAseq library using an mRNA isolation kit (New England BioLabs, E7490) and a NEBNext Ultra RNA library prep kit (New England BioLabs, E7530S) with a final double (0.65 x −1 x bead volume) size selection step using a SPRIselect reagent kit (Beckman Coulter, B23318). The libraries were indexed (New England BioLabs, E7335S) and sequenced on an Illumina NextSeq500. Seventy-six nucleotide pair-end reads were generated for each sample and aligned to the UCSC mm10 reference genome using tophat2 and bowtie2 in R (Kim et al., 2013; Langmead and Salzberg, 2012). Transcript abundance, differential expression, and isoform quantitation were calculated using the cufflinks suite in R (Roberts et al., 2011a; Roberts et al., 2011b; Trapnell et al., 2013; Trapnell et al., 2012). Raw data have been deposit in GEO under the accession number GSE124325.
Single-cell RNA-seq
FACS-purified lung ECs were processed through the Chromium Single Cell Gene Expression Solution Platform (10X Genomics) using the Chromium Single Cell 3’ Library and Gel Bead Kit in accordance with the manufacturer’s user guide (v2, rev D). The libraries were sequenced on an Illumina NextSeq500 using a 26X124 sequencing run format with 8 bp index (Read1). Chromium single-cell RNA-seq output was processed with Cell Ranger using “cellranger count” and “cellranger aggr”. Further analysis was carried out using Loupe Cell Browser (10x Genomics) or an R package Seurat https://satijalab.org/seurat/pbmc3k_tutorial.html. In brief, we first selected cells with at least 200 detected genes and filtered out cells with unique gene counts over 5000 or less than 200; no mitochondrial genes were found. Following filtering, we performed log normalization, identification of highly variable genes, scaling, PCA dimensionality reduction, and K-means clustering. Next, we identified differentially expressed genes per cluster and generated tSNE, dot, and violin plots, and heat maps. Raw data have been deposited in GEO under the accession number GSE124325.
ACKNOWLEDGEMENTS
We thank Drs. Napoleone Ferrara (Genentech; currently University of California San Diego, USA), Ralf Adams (University of Münster, Germany), Brigid Hogan (Duke University, USA) for providing the VegfaCKO, Cdh5-CreER, and SftpcCreER mice, respectively. We thank Kamryn Gerner-Mauro for assisting with scRNA-seq data analysis. The University of Texas MD Anderson Cancer Center DNA Analysis Facility and Flow Cytometry and Cellular Imaging Core Facility are supported by the Cancer Center Support Grant (CA #16672). This work was supported by the University of Texas MD Anderson Cancer Center Start-up Fund and National Institutes of Health R01-HL130129 (JC).