Abstract
The mesothelium forms epithelial membranes that line the bodies cavities and surround the internal organs. Mesothelia widely contribute to organ homeostasis and regeneration, and their dysregulation can result in congenital anomalies of the viscera, ventral wall defects, and mesothelioma tumors. Nonetheless, the embryonic ontogeny and developmental regulation of mesothelium formation has remained uncharted. Here, we combine genetic lineage tracing, in toto live imaging, and single-cell transcriptomics in zebrafish to track mesothelial progenitor origins from the lateral plate mesoderm (LPM). Our single-cell analysis uncovers a post-gastrulation gene expression signature centered on hand2 that delineates distinct progenitor populations within the forming LPM. Combining gene expression analysis and imaging of transgenic reporter zebrafish embryos, we chart the origin of mesothelial progenitors to the lateral-most, hand2-expressing LPM and confirm evolutionary conservation in mouse. Our time-lapse imaging of transgenic hand2 reporter embryos captures zebrafish mesothelium formation, documenting the coordinated cell movements that form pericardium and visceral and parietal peritoneum. We establish that the primordial germ cells migrate associated with the forming mesothelium as ventral migration boundary. Functionally, hand2 mutants fail to close the ventral mesothelium due to perturbed migration of mesothelium progenitors. Analyzing mouse and human mesothelioma tumors hypothesized to emerge from transformed mesothelium, we find de novo expression of LPM-associated transcription factors, and in particular of Hand2, indicating the re-initiation of a developmental transcriptional program in mesothelioma. Taken together, our work outlines a genetic and developmental signature of mesothelial origins centered around Hand2, contributing to our understanding of mesothelial pathologies and mesothelioma.
Introduction
As a key feature of the vertebrate body plan, the mesothelium is composed of several continuous, epithelial monolayers surrounding the internal organs (visceral mesothelium) and lining the body cavities (parietal mesothelium). The mesothelium provides a protective layer against invasive microorganisms, produces serous fluid that both decreases friction of moving organs, and enables the transport of cells and nutrients across serosal cavities (Mutsaers, 2002; Mutsaers and Wilkosz, 2007). Moreover, cell tracking studies have established that the mesothelium contributes to a multitude of downstream cell fates including smooth muscles and fibroblasts during organogenesis, tissue homeostasis, and regeneration (Carmona et al., 2018; Koopmans and Rinkevich, 2018; Rinkevich et al., 2012). While mesothelium-lined body cavities are a fundamental trait across bilaterian animals (Hartenstein and Mandal, 2006; Monahan-Earley et al., 2013; Technau and Scholz, 2003), little is known about how the vertebrate mesothelium lineage initially emerges during development.
The embryonic mesothelium, also called coelomic epithelium after establishing its baso-apical polarization, is a highly dynamic cell layer that undergoes epithelial-to-mesenchymal transition (EMT) during development and seeds mesenchymal cells to underlying tissues (Carmona et al., 2018; Rinkevich et al., 2012). Several prior observations tie mesothelial lineage origins to the lateral plate mesoderm (LPM) (Ahn et al., 2002; Carmona et al., 2016; Winters et al., 2012), a mesodermal progenitor territory that forms at the periphery of the early vertebrate embryo (Prummel et al., 2020). In vertebrates, the coelomic cavity forms stereotypically by splitting the LPM into dorsal and ventral layers (Onimaru et al., 2011). A subset of cells within both layers differentiates into polarized epithelial cells that form visceral (splanchnic) mesothelial layers and parietal (somatic) mesothelial layers (Funayama et al., 1999; Sadler and Feldkamp, 2008; Sheng, 2015). Ultimately, the coelom in amniotes spans from the neck to the abdomen and outlines four main body compartments: two pleural cavities (around the lungs), a pericardiac cavity (around the heart), and a peritoneal (abdominal) cavity, each with their associated mesothelial layers (Fig. 1A). Representative for teleosts, zebrafish feature mesothelium-lined cardiac and abdominal cavities. Which territories within the emerging LPM initially harbor the mesothelial progenitors, how and when the mesothelium diverges from other LPM lineages, and how the visceral and parietal layers form remain uncharted.
Embryonic studies of the mesothelium have predominantly focused on developmental stages after the coelomic epithelium has formed. Several genes, including Mesothelin (Msln), Gata4, Tbx18, Tcf21, and Wilms Tumor 1 (Wt1), have enabled labeling and genetic lineage tracing of mesothelial lineages in mouse and chick (Alghamdi et al., 2020; Ariza et al., 2016; Chau et al., 2014; Delgado et al., 2014; Rinkevich et al., 2012). The Wt1-expressing coelomic epithelium contributes to the mature mesothelium, fibroblasts, stellate cells, smooth muscles, and white adipose tissue associated with the gastrointestinal tract, liver, lungs, urogenital system, and heart (Asahina et al., 2011; Cano et al., 2013; Carmona et al., 2016; Delgado et al., 2014; Lee et al., 2019; Lua et al., 2014; Martínez-Estrada et al., 2010; Rinkevich et al., 2012; Sebo et al., 2018; Zhou et al., 2008). Further, regional specification within the mesothelial components along the developing gut is recognizable early in mouse development (Han et al., 2020; Kishimoto et al., 2020). Studies using zebrafish have documented expression of wt1a/b, tcf21, and tbx18 in the epicardium, the visceral mesothelial layer covering the heart (Peralta et al., 2013; Peralta et al., 2014). Despite these advances across models, expression of these conserved genes is initiated after the onset of coelomic epithelium formation, leaving the earliest differentiation steps obscure.
Compromised integrity of the adult mesothelium can result in pathologies including intra-abdominal organ adhesion (Tsai et al., 2018), serosal fibrosis (Mutsaers et al., 2015), pericarditis (Chiabrando et al., 2020), and mesothelioma tumors (Carbone et al., 2019; Yap et al., 2017). Malignant mesothelioma is a rapidly fatal solid tumor that can arise within the visceral or parietal mesothelia, predominantly as the result of environmental exposure to asbestos (Carbone et al., 2019; Felley-Bosco and Macfarlane, 2018; Odgerel et al., 2017; Wagner et al., 1962; Yap et al., 2017). While mesothelioma cases are increasing globally despite regulatory means to curb the use of causative agents, treatment remains limited (Hinz and Heasley, 2019; Yap et al., 2017). Presenting predominantly as epithelioid, sarcomatoid, and biphasic phenotypes, malignant mesothelioma frequently harbor genetic alterations affecting the tumor suppressors BAP1, NF2, CDKN2AB, and TP53 (Bueno et al., 2016; Cheng et al., 1994; Hmeljak et al., 2018; Quetel et al., 2020; Tate et al., 2019). Nonetheless, the cell of origin and the underlying aberrant molecular mechanisms leading to mesothelioma remain uncertain.
In addition to the mesothelium, the LPM forms a vast array of downstream cell fates that include the cardiovascular system, blood, kidneys, and limb connective tissue (Prummel et al., 2020). How the LPM partitions into its diverse fates and what regulatory programs specify the individual progenitor fields remains unclear. Emerging as a dedicated mesendoderm domain, the post-gastrulation LPM segments into recognizable bilateral territories discernible by the expression of several transcription factor genes including Scl/Tal1, Lmo2, Pax2a, Nkx2.5, and Hand1/2 (Chal and Pourquié, 2017; Davidson and Zon, 2004; Prummel et al., 2020; Takasato and Little, 2015). dHand/Hand2, encoding a conserved basic helix-loop-helix transcription factor, is expressed during segmentation stages in the most laterally positioned LPM progenitors in amphioxus and zebrafish (Onimaru et al., 2011; Perens et al., 2016; Prummel et al., 2019). In the developing mouse embryo, Hand2 expression has been described in the flank at comparable embryonic stages (Charité et al., 2000; Firulli et al., 1998; Srivastava et al., 1995). Studies across vertebrate models have revealed key insights into the contribution of Hand2 and its paralog eHand/Hand1 in anterior LPM (ALPM) progenitors that contribute to the heart, (fore)limbs, and branchial arches (Funato et al., 2009; Yelon et al., 2000; Yin et al., 2010). Additionally, in the posterior LPM of zebrafish and chick, Hand2 has been linked to refining the fate divergence between smooth muscle versus hemangioblast and kidney fates during somitogenesis (Gays et al., 2017; Perens et al., 2016; Shin et al., 2009). The definitive fate of especially the posterior Hand2-expressing progenitors has remained unclear.
Here, we establish several lines of evidence that hand2 in zebrafish, both by expression and by function, is the earliest specific transcription factor gene demarcating the emerging mesothelial progenitors within the LPM. We provide further evidence for conservation of this property in mouse. We link the developmental function of Hand2 in mesothelium formation to a reactivation of an early coelomic epithelium-focused LPM program in mouse and human mesothelioma tumors. Our findings propose that Hand2 expression contributes to the unique properties of mesothelial progenitor cells in development and in mesothelioma.
Results
Zebrafish mesothelium is LPM-derived
To formally assess whether in zebrafish mesothelial membranes are bona fide LPM lineages, we performed genetic lineage tracing using drl:creERT2 that is active in LPM-primed progenitor cells from the onset of gastrulation until early-to-mid somitogenesis (Mosimann et al., 2015; Prummel et al., 2019). We induced drl:creERT2;hsp70l:Switch embryos with 4-OH-Tamoxifen (4-OHT) at shield stage and analyzed the resulting EGFP-based LPM lineage labeling in transverse sections at 3 days post-fertilization (dpf) when LPM-derived organs are clearly detectable (Fig. 1A,B). EGFP expression recapitulated broad labeling of LPM-derived organs including endothelium, blood, and cardiac lineages, and sparse labeling of endoderm-derived organs, in line with previous observations using drl:creERT2 (Mosimann et al., 2015; Prummel et al., 2019) (Fig. 1C-G, Fig. S1A-C). Extending the previously reported LPM lineage labeling in the Transgelin (Tagln)-positive smooth muscle layers around the zebrafish gut (Fig. 1C)(Gays et al., 2017), we also observed smooth muscles around the swim bladder and ducts within the liver by double-positive staining for EGFP and Tagln (Fig. 1C). In addition, we consistently observed EGFP-positive, yet Tagln-negative cells as thin epithelial layers surrounding the gut and swim bladder (Fig. 1C) as well as around other endodermal organs including the liver and the pancreas (Fig. 1D,E). We also found EGFP lineage labeling in the pericardial and epicardial layers surrounding the heart, confirming their LPM origin (Fig. 1F, Fig. S1A,D-I). Of note, at this stage, the pericardium directly adheres to the body wall, forming a sac around the cardiac cavity, while in adult zebrafish the pericardium establishes a dedicated body cavity. Lastly, we observed prominent drl-based LPM lineage labeling within the body wall as an epithelial layer right underneath the skin surrounding the yolk and the yolk extension along the anterior-posterior axis (Fig. 1G,H). From these observations, we conclude that the prospective coelomic epithelium and most, if not all, developing mesothelial layers in zebrafish are LPM-derived.
The early LPM consists of distinct progenitor populations
We next sought to chart the mesothelial progenitors within the emerging LPM. To probe whether early mesothelial progenitors can be recognized within the emerging LPM, we analyzed the transcriptome of individual zebrafish LPM cells by single-cell RNA-sequencing (scRNA-seq) at tailbud stage (Fig. 2A). At this stage, the end of gastrulation, the drl:mCherry reporter-labeled LPM comprises approximately 7% of all cells in the zebrafish embryo (Fig. 2B, Fig. S2). Upon quantifying the transcriptomes from individually sorted drl:mCherry-positive cells using CEL-Seq2 (Muraro et al., 2016), we obtained 1039 cells that passed filtering and quality control. Using graph-based clustering (Louvain algorithm (Blondel et al., 2008)), we called 15 distinct cell populations within the drl-positive LPM at tailbud stage (Fig. 2C). We annotated these as contributing to 8 major subpopulations based on canonical markers and published gene expression patterns in conjunction with marker genes identified through differential expression analysis (Fig. 2E, see Methods for details).
While mRNA in situ hybridization (ISH) and transgene expression for the earliest markers of individual LPM fate potentials renders them chiefly detectable from early somitogenesis on (Davidson and Zon, 2004), our analysis at tailbud stage resolved seemingly determined LPM progenitor fields already at the end of gastrulation, in line with and extending previous findings in zebrafish and mouse embryos (Farrell et al., 2018; Mohammed et al., 2017; Mosimann et al., 2015; Pijuan-Sala et al., 2019; Scialdone et al., 2016; Wagner et al., 2018). Our uncovered clusters broadly represent the cardiopharyngeal, emerging head mesoderm, hatching gland progenitors, endoderm, and endothelial and hematopoietic progenitors (Fig. 2C,E). Three clusters were composed of presumptive cardiopharyngeal progenitors based on the expression of tbx20, nkx2.5, nkx2.7, hey2, gata4/5/6, ttn.1, and ttn.2 (Fig. 2C,E) (Bloomekatz et al., 2017; Gibb et al., 2018; Lee et al., 1996; Lu et al., 2017; Peterkin et al., 2009; Shih et al., 2015; Wang et al., 2019a). One cluster was positive for designated markers of putative head mesoderm progenitors, including fsta, foxc1a, gsc, meox1, and prrx1a (Wang et al., 2019b). Based on the expression pattern of alx1, pitx2, and pitx3, we assigned a cluster to represent the hatching gland progenitors (John et al., 2013; Zilinski et al., 2005), in line with drl reporter activity in these cells (Mosimann et al., 2015). We additionally uncovered cells expressing the endodermal genes sox32, sox17, gata5, gata6, foxa2, and fox3a (Farrell et al., 2018; Schier and Talbot, 2001; Wagner et al., 2018; Warga and Nüsslein-Volhard, 1999) as distinct group of clusters within the analyzed cells (Fig. 2C,E). This observation is in line with previous findings that the drl reporter-positive cells at tailbud stage represent either mixed endoderm- and LPM-primed populations or a bi-potential LPM-fated mesendoderm population (Prummel et al., 2019).
Towards reconstructing rudimentary positional information back to the dissociated single cells, we mapped the homeobox transcription factor gene cdx4 and several hox genes to assign whether a cluster likely represented anterior or posterior cells within the embryo (Fig. 2D, Fig. S3). cdx4 at 5 somite stages broadly demarcates in the posterior half of the developing zebrafish embryo (Fig. 2D). cdx4-positive and thus posterior clusters composed of cells expressing marker genes for endothelial and hematopoietic precursors (fli1a, lmo2, znfl2a) (Hogan et al., 2006; Lawson and Weinstein, 2002; Zhu et al., 2005), and for the pronephros (pax2a, pax8, lbx2) (Naylor et al., 2016; Perens et al., 2016; Pfeffer et al., 1998) (Fig. 2E). The overlapping expression of hemangioblast and kidney markers possibly indicates that these two clusters represent a mixed multi-lineage progenitor pool at the end of gastrulation.
Notably, among all the prominent LPM genes we detected hand2 expression across several clusters. In addition to clusters encompassing the expected cardiopharyngeal precursors, hand2 transcripts were abundant in four clusters we accordingly named hand2-high_1-4 (Fig. 2C,E). Further, expression of hand2 fell into both anterior and posterior expression domains, correlating with its native expression pattern in zebrafish (Yelon et al., 2000) (Fig. 2C,E). As anticipated, hand2 transcripts coincided with the expression of cardiac genes (Fig. 2E). However, the anterior hand2-high_3 and hand2-high_4 clusters showed no detectable expression of cardiac markers including hey2, tbx20, nkx2.5, and nkx2.7, suggesting that they encompass cells with another fate potential. We also found posterior hand2-positive cells that appeared distinct from endothelial, hematopoietic, and kidney progenitors (hand2-high_1 and hand2-high_2) (Fig. 2E). Taken together, our scRNA-seq captured hand2 expression as a central feature of several LPM progenitor clusters.
We next aimed to determine what genes are co-expressed with hand2. Our analysis revealed that several genes were enriched and individually even among the cluster-defining genes in one or more of the hand2_high clusters, including sfrp5, foxh1, gata5, gata6, and meis3 (Fig. 2F-K, Fig. S4). In fluorescent mRNA ISH (RNAscope) and colorimetric mRNA ISH on whole-mount tailbud and early somitogenesis staged embryos, we observed that the endogenous expression pattern of meis3 expression overlaps with the hand2 domain in the anterior and posterior LPM, among with meis3 expression in other non-LPM domains (Fig. 2G,I). Moreover, foxh1 expression has previously been described in the LPM during somitogenesis stages (Pogoda et al., 2000; Slagle et al., 2011), yet the exact domain within the LPM has remained unclear. Akin to meis3, our fluorescent ISH confirmed that foxh1 is expressed in the lateral-most posterior LPM territory overlapping with hand2 expression (Fig. 2H). Notably, the in part redundant transcription factor genes gata4, gata5, and gata6 that play key roles in cardiac and endoderm development (Tremblay et al., 2018), are also expressed in a domain lateral to the forming heart field (Jiang et al., 1999; Reiter et al., 1999). The cells in this domain take on a spread-out, mesh-like pattern over the yolk during somitogenesis (Fig. 2J). Similarly, we found that sfrp5 demarcates lateral-most LPM cells that form a spread-out expression domain along the body axis over the yolk and yolk extension (Fig. 2K). This mesh-like pattern can also be recognized within the endogenous expression of hand2 within the ALPM (Yelon et al., 2000). Taken together, our data documents a collection of genes co-expressed with hand2 in the lateral-most LPM domain of a yet-unknown fate.
hand2 expression identifies mesothelial progenitors
Our analysis indicated that expression of hand2 and several associated genes demarcate a lateral-most LPM domain that is distinct from the cardiopharyngeal, blood, vasculature, and kidney progenitors at the end of gastrulation. Based on the previous association of Hand1/2 expression with differentiated mesenchymal structures in mouse and chick (Ahn et al., 2002; Charité et al., 2000; Fernandez-Teran et al., 2000; Firulli et al., 1998; Perens et al., 2016; Prummel et al., 2019), we hypothesized that the hand2-expressing LPM in zebrafish, in particular the posterior lateral-most stripe, forms the mesothelium. We turned to the transgenic line hand2:EGFP based on a BAC encompassing the zebrafish hand2 locus that faithfully recapitulates endogenous hand2 expression (Yin et al., 2010). Using time-lapse SPIM imaging and panoramic projections, we captured the dynamics of hand2:EGFP reporter activity during segmentation stages, demarcating the lateral-most drl-expressing LPM (Fig. 3A,B) (Perens et al., 2016; Prummel et al., 2019). Notably, and in contrast to the more medial LPM stripes that progressively migrated to the midline, a subset of the hand2:EGFP-expressing cell population sprawled out laterally over the yolk as a single-cell layer (Fig. 3C).
In transverse sections of hand2:EGFP-transgenic embryos at 3 dpf, we observed the previously described hand2 reporter expression in Tagln-positive, LPM-derived intestinal smooth muscle cells that layer around the endodermal gut tube (Gays et al., 2017) (Fig. 3D). In higher magnification analysis of transverse sections, we also observed Tagln-negative hand2:EGFP-expressing cells surrounding the gut at 3 dpf, reminiscent of coelomic epithelial cells (Fig. 3D). At 3 dpf, the hand2 reporter-expressing layers had also wrapped around other endoderm-derived organs, including the liver and the pancreas (Fig. 3D,E). In addition, in transverse sections capturing the yolk at 3 dpf, we detected hand2:EGFP expression underneath the skin in the body wall, the prospective parietal peritoneum (Fig. 3F). Extending previous observations (Gays et al., 2017; Mao et al., 2015; Perens et al., 2016; Yin et al., 2010), these data suggest that hand2:EGFP reporter expression at 3 dpf in zebrafish delineates the visceral and parietal peritoneal membranes.
To extend our interpretation of zebrafish hand2 as marker for the emerging mesothelium, we next turned to the mouse to uncover any mesothelial lineage contribution of cells expressing either of the two partially redundant murine Hand genes Hand1 and Hand2. Crossing either the Hand1EGFPCreΔNeo/+ (Barnes et al., 2010) or Hand2Cre (Ruest et al., 2003) transgenic strains into the R26R loxP reporter strain (Soriano, 1999) resulted in lineage labeling along the mesothelium lining of the liver lobes at E14.5 and E15.5, respectively (Fig. 3G,H). We found lineage labeling to be more apparent and widespread using the Hand1EGFPCreΔNeo/+ strain, with strong staining in most of the mesothelium and in a subset of cells within the liver of currently unknown identity (Fig. 3G). Labeling was also present in some cells lining sinusoids, though more sporadic (data not shown). Lineage analysis of Hand2Cre-descendant cells also revealed lineage labeling in the mesothelium (Fig. 3H), though staining was more restricted and less robust than that observed in Hand1 daughter cells. These differences could reflect differences between the two Cre drivers (knockin vs transgenic) or a larger proportion of Hand1-expressing cells contributing to the liver mesothelium.
To further investigate a possible conserved link of Hand gene expression to mesothelium formation in mouse, we mined recent scRNA-seq and scATAC-seq data of early mouse embryos (Pijuan-Sala et al., 2020) (Fig. S5). This data harbored Hand2- expressing cell clusters that were previously assigned as “mesenchymal” and that share co-expression of transcription factor genes we found in our zebrafish LPM dataset as associated with mesothelial progenitors, including Meis3, Gata5, and Gata6 as well as the previously LPM-associated genes Gata4, FoxF1, and Hand1 (Prummel et al., 2020) (Fig. 2, Fig. S5). Together, these observations indicate that also in mammals Hand gene expression is a conserved feature of the mesothelial progenitors.
To further confirm and define the hand2-expressing mesothelia, we turned to Wt1 expression that characterizes the developing visceral mesothelium in mammals (Chen et al., 2014; Parenti et al., 2013; Walker et al., 1994). Expression of the zebrafish Wt1 paralogs wt1a and wt1b becomes detectable at 6-8 ss as previously analysed in kidney and epicardium development (Bollig et al., 2009; Endlich et al., 2014; Peralta et al., 2014; Perner et al., 2016). During somitogenesis, we found wt1a:EGFP and wt1b:EGFP activity lateral of the differentiating kidney structures within the drl:mCherry-expressing LPM corresponding to the hand2-positive territory (Fig. S6A,B). In transverse sections of wt1a- and wt1b:EGFP-expressing embryos, we detected EGFP signal in the coelomic epithelium surrounding the gut, liver, and pancreas in addition to the previously reported labeling of the glomerulus and pronephric tubules (Fig. 3J-K) (Bollig et al., 2009; Endlich et al., 2014; Peralta et al., 2014; Perner et al., 2016). Further, we observed wt1a/b reporter-expressing Tagln-positive smooth muscle cells around the gut and hepatic and pancreatic ducts (Fig. 3J-K). Genetic lineage tracing using wt1a- and wt1b:creERT2 from the onset of transgene expression at 6-8 ss robustly marked the visceral peritoneum around the gut, liver, and pancreas (Fig. S6C-E). Notably, at 3 dpf, we neither detected any EGFP-labeling of the visceral peritoneum around the more posterior gut, nor did we detect any EGFP-expressing parietal peritoneum cells around the yolk, indicating these cells do not express wt1 genes in zebrafish at our analyzed time points (Fig. S6F-G). Extending previous wt1a-based lineage tracing (Sánchez-Iranzo et al., 2018a), we observed that also wt1b-expressing cells contribute to the cardiac mesothelial layers in the dorsal and ventral pericardium, and in the pro-epicardial clusters (Fig. S6H,I). These observations are consistent with LPM lineage tracing using drl:creERT2 to ventral and dorsal pericardium and to proepicardium (Fig. 1F, Fig. S1D-I). We conclude that, akin to the mammalian mesothelium, expression of both Wt1 orthologs is also a feature of developing mesothelia in zebrafish. Nonetheless, in addition to the absence of wt1a/b expression in the parietal peritoneum lineage, hand2:EGFP presents an earlier and more complete marker of all developing mesothelial membranes in zebrafish.
Primordial germ cells associate with mesothelial progenitors
We noted the chemokine-encoding gene cxcl12 significantly co-expressed with hand2 in our scRNA-seq data set (Fig. 2E, Fig. 4A). Cxcl12a provides directional cues for the migration of several cell types (Lewellis and Knaut, 2012), including the guidance of primordial germ cells (PGCs) (Grimaldi and Raz, 2020). Through complex migration paths, the PGCs reach their final destination in the region where the gonad develops, dorsally and medially bordered by the developing pronephros, while the identity of the ventrally located tissue has not been precisely defined (Doitsidou et al., 2002; Richardson and Lehmann, 2010; Stebler et al., 2004). We asked whether the drl- and hand2-expressing mesothelium interacts with the migrating PGCs and if PGCs are possibly already associated with the developing LPM during gastrulation. In time-lapse imaging of transgenic zebrafish embryos expressing EGFP in the LPM (drl:EGFP transgene) and the farnesylated mCherry in their PGCs (kop:mCherry-f’-nos3’UTR transgene) (Blaser et al., 2005), we observed that the migrating PGCs associate with the drl reporter-expressing mesendoderm already during gastrulation (Fig. 4B,C, Movie 1). The drl:EGFP-expressing progenitor field is gradually specifying into the LPM at the beginning of somitogenesis (Prummel et al., 2019) and the PGC clusters continued to associate with, and migrate within, the forming LPM (Fig. 4B,C). By 24 hpf, the PGCs reach the region where the gonads will form (Raz, 2003) and we speculated the hand2-positive mesothelium-primed LPM serves as the final destination for the PGCs. Imaging mCherry-f’-nos3’UTR;hand2:EGFP embryos documented how the PGCs migrated within the hand2:EGFP-expressing LPM, and the PGCs eventually arrive at a domain within the hand2-expressing forming mesothelium (Fig. 4D). Antibody staining for the kidney transcription factor Pax2a confirmed that the PGCs end up ventral of the pronephros within the hand2-positive LPM (Fig. 4,EF), with the developing gut located medially between the PGC clusters (Paksa et al., 2016). These observations further support our conclusion that the PGCs migrate in close association with the forming coelomic epithelium, as marked by hand2 expression, already during early development.
hand2-expressing LPM forms the visceral and parietal mesothelium
To clarify their mesothelial identity and to resolve the dynamics of hand2:EGFP-expressing mesothelial progenitor cells in the zebrafish embryo, we performed long-term and in toto light sheet imaging of developing hand2:EGFP;drl:mCherry double-transgenic embryos. Applying a multi-sample imaging and processing workflow (Daetwyler et al., 2019), we captured the dynamics of the hand2:EGFP expression over the course of embryonic development from 18 to 82 hpf in toto (n = 6 embryos). At imaging onset, the hand2:EGFP-expressing cells had initiated their lateral migration over the yolk, while expression of drl:mCherry refined to cardiovascular and blood lineages (Fig. 5A, Movie 2) (Mosimann et al., 2015).
In the resulting time lapse imaging, we captured the hand2:EGFP-positive LPM as a dynamic cell population with distinct migration behaviors in different regions of the developing embryo. First, we observed the hand2:EGFP-expressing cells contributing to the previously described processes of pharyngeal arch formation, pectoral fin bud outgrowth, and primary heart tube extension (Fig. 5B, Movie 2,3). Additionally, our light sheet imaging registered that two seemingly separated populations of hand2:EGFP-expressing cells migrate ventro-laterally, crawling over the yolk and yolk extension, and continue to form the parietal layer of the cardiac cavity (pericardium) and abdominal cavity (parietal peritoneum), respectively (Fig. 5C-G). Already at 18 hpf, we observed a dorso-ventral split in the hand2-expressing progenitor field positioned lateral of the cardiopharyngeal progenitors, forming the cardiac cavity (Fig. 5E). The dorsal and ventral pericardium surround the forming heart tube and maintain the pericardial fluid that supports the beating, moving heart (Peralta et al., 2013). The cells that will form the dorsal pericardium migrated over the yolk as a connected cell field with a clearly visible leading edge and were already interacting with the forming primary heart tube (Fig. 3B, 5D, Movie 2,3). We observed that the cells forming the more posterior-located parietal peritoneum are split from the forming pericardium by the circulation valley forming over the anterior yolk, where later the common cardinal vein (Duct of Cuvier) migrates into to ultimately enclose the circulating blood (Hamm et al., 2016) (as visualized by drl:mCherry, Movie 1, Fig. S7). In addition, we documented how the hand2:EGFP-cells forming the presumptive parietal mesothelium reach the ventral midline earlier around the caudal yolk extension than around the main yolk, possibly due to the shorter migration distance over the yolk extension’s smaller diameter (Fig. 5F-H). Simultaneously, the more medially located hand2:EGFP-expressing cells migrated towards the midline (Fig. 5F-H), corresponding to the formation of visceral peritoneum including intestinal smooth muscles (Gays et al., 2017; Yin et al., 2010). Taken together, these imaging data further support the notion that hand2:EGFP-expressing LPM progenitors in zebrafish form the visceral, parietal, and pericardiac mesothelium.
hand2 mutants fail to form their mesothelia and to close the ventral body wall
The prominent expression of hand2 in mesothelial progenitors of the LPM prompted us to investigate whether the loss of hand2 has an impact on the formation of the mesothelial layers. Phenotype analyses of mutant mice and zebrafish have documented that Hand2 (partially redundantly with Hand1 in mice) contributes to the development of the heart, forelimbs/pectoral fins, and smooth muscle lineages (Galli et al., 2010; Laurent et al., 2017; Santoro et al., 2009; Srivastava et al., 1997; Yelon and Stainier, 2005; Yelon et al., 2000). Additionally, zebrafish hand2 mutants feature disorganized migration of the intestinal smooth muscle progenitors towards the gut and overall gut mislooping due to disorganized extracellular matrix remodeling (Gays et al., 2017; Yin et al., 2010).
We revisited the previously established zebrafish mutant hanS6 that harbors a presumptive hand2 null allele (Yelon et al., 2000). In addition to the well-described cardiac and pectoral fin defects, we observed that hanS6-homozygous embryos displayed ventral defects of their yolk extensions and subsequent deterioration of the embryo due to yolk herniation at 3 dpf (Fig. 6A-D). Moreover, we noticed selective blistering and shriveling of the ventral fin fold. At the cellular level, in contrast to the connected epithelial layer forming mesothelia in wildtype embryos, we observed that LPM-derived cells (marked by genetic lineage tracing using drl:creERT2) are sparse in the abdominal cavity in hanS6-homozygous embryos (Fig. 6E). The sparse distribution of LPM-derived cells could potentially result from mosaic labeling from incomplete loxP-reporter recombination following drl-driven CreERT2 activation, as common in Cre/loxP-based experiments (Carney and Mosimann, 2018). However, hanS6-homozygous embryos displayed similar migration issues when we directly visualized for the mesothelial cells with the wt1a:EGFP and hand2:EGFP transgenic reporters, respectively (Fig. 6F,G). We therefore conclude that loss of hand2 perturbs mesothelial progenitor migration, in line with previous observations indicating that coelomic epithelial progenitors migrated to the midline in hand2 mutants but failed to properly wrap around the gut (Yin et al., 2010). Of note, we only observed few LPM lineage-labeled cells and hand2:EGFP-expressing cells lining the body wall, respectively, indicating that also the ventrally migration of the parietal peritoneum progenitors is perturbed in hand2 mutants (Fig. 6E,G). Consequently, hanS6-homozygous embryos also fail to form a proper body wall, likely resulting in herniation of the yolk along the anterior-posterior axis over time. These observations in zebrafish connect hand2 expression and function in dedicated LPM progenitor cells to the proper execution of mesothelial membrane formation.
Mesothelioma activates early LPM transcription factors including Hand2
Upon exposure to asbestos fibers, mesothelial cells within the visceral and parietal pleura can transform into mesothelioma tumors (Carbone et al., 2019). As the cell(s) of origin have been hypothesized to reside within the mesothelium, we sought to determine whether mesothelioma tumors feature a transcriptional signature akin to the mesothelial progenitors we found in zebrafish (Fig. 2).
First, we revisited our transcriptomics data set obtained from mouse mesothelioma triggered by repeated exposure to crocidolite fibers (blue asbestos) (Rehrauer et al., 2018) (Fig. 7A). As previously established, genes found upregulated in human mesothelioma development (including Msln and Wt1) or downregulated (including tumor suppressors Nf2 and Bap1) behaved analogously in this mouse model, both in pre-neoplastic lesions and in fully formed tumors when compared to healthy adult mesothelium (Fig. 7B) (Bianchi et al., 1995; Bott et al., 2011; Bueno et al., 2016; Hmeljak et al., 2018; Rehrauer et al., 2018; Sekido et al., 1995). Early LPM lineage markers, including Scl/Tal1, Etv2, and Pax2, were not detected with RNA-seq in the adult mouse mesothelium samples. Notably, the sham-treated mesothelial tissue also expressed no or low levels of transcripts for LPM-associated genes including Hand2, Gata4/5/6, Meis3, FoxF1, Mixl1, and Lmo2 (Fig. 7B). In contrast, the transition to pre-neoplastic lesions and to mesothelioma after crocidolite exposure was accompanied by the upregulation of mouse orthologs of several genes associated with mesothelium progenitors in zebrafish: expression of Hand2, Wt1, Gata4/5/6, and Meis3 increased in crocidolite-exposed mesothelium, with a particularly striking upregulation in fully formed tumors (Fig. 7B). Also FoxF1 was upregulated in crocidolite-induced tumors, a gene which has been associated with the developing splanchnic mesoderm in lamprey, chicken, and mouse, but not in zebrafish (Funayama et al., 1999; Mahlapuu et al., 2001; Onimaru et al., 2011; Ormestad et al., 2004) (Fig. 7B). Following immunohistochemistry for Hand2, we detected specific and prominent nuclear Hand2 immunoreactivity in crocidolite-induced mesothelioma tumor sections (Fig. 7C). Altogether, these observations suggest that the transformative events leading to mesothelioma in a crocidolite-based mouse model are accompanied by the upregulation (or re-initiation) of early LPM genes we found associated with mesothelial progenitors in development (Fig. 2).
We next sought to corroborate whether human mesothelioma also feature HAND2 expression. Analyzing expression levels of HAND2 using qPCR in human pleural mesothelioma (n = 36) compared to healthy pleura (n = 4), we observed that individual tumors showed heterogeneous yet consistent upregulation of HAND2, with individual tumors expressing high levels of HAND2 mRNA (p = 0.0596) (Fig. 7D). We further detected nuclear HAND2 immunoreactivity in an epithelioid mesothelioma sample with high HAND2 mRNA levels (Fig. 7E). To further establish if expression of HAND2 and other mesothelial progenitor-associated LPM genes are re-activated in mesothelioma, we mined the mesothelioma-associated transcriptome data deposited in the The Cancer Genome Atlas (TCGA) as generated by the TCGA Research Network (https://www.cancer.gov/tcga). We compared gene expression for HAND2 and other mesothelial progenitor genes with a collection of mesothelioma-associated genes, and several tissue-specific and ubiquitous house-keeping genes. We found that increased or re-activated HAND2 expression was not an obvious signature across the analyzed mesothelioma samples, but rather a feature of a subset of tumors across the epithelioid, sarcomatoid, and biphasic classifications (Fig. 7F). HAND2 expression coincided with differential expression of the previously mesothelium- and mesothelioma-associated genes MSLN and WT1, as well as with GATA5 and MEIS2 (Fig. 7F). Notably, while we found in zebrafish mesothelial progenitors meis3 as the most-prominent MEIS family gene co-expressed with hand2 (Fig. 2), its redundant paralogs meis2a and meis2b were also enriched in hand2-expressing zebrafish progenitors (Fig. S4) (Schulte and Geerts, 2019). This HAND2-expressing group of mesothelioma tumors did not cluster notably with tumors featuring differential expression of the common mesothelioma-associated tumor suppressor genes BAP1 and NF2 (Fig. 7F). Taken together, these observations provide first indications that mesothelioma tumors can upregulate or reactivate several developmental factors associated with an mesothelium progenitor state, in particular Hand2 that in zebrafish provides the earliest mesothelium progenitor marker. We postulate that, following malignant transformation, mesothelioma tumors acquire features of a developmental progenitor program deployed by the early LPM to specify mesothelium progenitors, endowing transformed cells with unique migratory and lineage properties.
Discussion
Despite their various contributions to development and homeostasis, the early embryonic origins of mesothelia and their genetic control have remained uncertain. Uniquely positioned within the developing embryo to cover inner organs and the forming body cavities, our results uncover that mesothelial progenitor cells emerge among the earliest distinguishable LPM cell fates following hand2 expression.
Our finding that expression of the transcription factor Hand2 demarcates the merging mesothelial progenitors within the post-gastrulation LPM provides new means to study the mesothelium in development and disease. Hand gene function centrally contributes to cardiac, limb, and pharyngeal arch development across several models (Barnes et al., 2011; Firulli et al., 1998; Han and Olson, 2005; Shin et al., 2009; Yelon et al., 2000). Our results indicate that Hand genes also act as conserved regulators of mesothelial fates. Gene expression studies have defined hand2 as a marker for the forming body wall in tilapia and medaka, and have found Hand1/2 in the flank of the developing mouse embryo (Srivastava et al., 1995; Srivastava et al., 1997; Tanaka, 2011; Tanaka et al., 2015). Complementing studies in mice have demonstrated that Hand1-expressing progenitors contribute to cell layers of the gut including smooth muscles (Barnes et al., 2010; Srivastava et al., 1995; Srivastava et al., 1997). A possible ancient association of Hand genes with the LPM and thus mesothelium progenitors followed the expression of amphioxus AmpiHand and lamprey LjHandA within their presumptive LPM (Onimaru et al., 2011). While zebrafish have seemingly lost their Hand1 ortholog (Howe et al., 2013), our data suggest an early role for Hand2 and Hand factors in general in the differentiation of mesothelial progenitors from the LPM (Fig. 3).
Several fates emerging from the LPM stem from multi-lineage progenitors, such as the Scl/Tal1-positive hemangioblasts that contribute to endothelial and hematopoietic lineages (Choi et al., 1998; Murray, 1932; Sabin, 1917; Vogeli et al., 2006). Our single-cell transcriptomics analysis of the early drl reporter-expressing LPM now captured a surprising heterogeneity within the forming LPM already at the end of zebrafish gastrulation, including putative cardiopharyngeal, endothelial/hematopoietic, and here defined mesothelial progenitor clusters (Fig. 2). These data complement and expand prior scRNA-seq in mouse, zebrafish, and tunicate embryos that captured numerous mesendodermal and mesodermal cell fate territories at various developmental stages (Farrell et al., 2018; Mohammed et al., 2017; Pijuan-Sala et al., 2019; Scialdone et al., 2016; Wagner et al., 2018; Wang et al., 2019a). With wt1a/b expression appearing later than hand2 and seemingly restricted to the visceral mesothelial layers (Fig. 3), hand2 provides the earliest and most complete marker for mesothelial progenitors in zebrafish and possibly beyond. Our findings are applicable to in vitro differentiation protocols for multilineage organoids with desirable stromal components, such as to guide pluripotent stem cell differentiation into representative cell types of the gastro-intestinal tract. Further, our work provides a framework to interpret cell types deemed mesenchymal in numerous systems.
Our results provide several new insights into mesothelial progenitor biology. Combined expression of hand2 with sfrp5, meis3, and foxh1 potentially defines a rudimentary mesothelial progenitor signature in the early LPM (Fig. 2E-K; Fig. S4). Individually, these genes have all previously been linked to mesothelial biology. sfrp5 expression has been reported in mesenchymal cells around internal organs in zebrafish and in the foregut-surrounding epithelium in Xenopus (Li et al., 2008; Stuckenholz et al., 2013). meis3 expression has been linked to mesenchymal cells lining the developing intestine and to enteric neuron migration in zebrafish (Reichenbach et al., 2008; Uribe and Bronner, 2015); we note that the potentially redundant meis2a/b are also expressed in the early LPM and mesothelial progenitors (Fig. S4). During segmentation stages, foxh1 is involved in controlling left-right asymmetry in response to Nodal signaling in a lateral LPM domain of so-far unassigned fate (Slagle et al., 2011; Tavares et al., 2007); our data now proposes that this activity of foxh1, and possibly the response to left-right cues, is confined to the hand2-positive mesothelial progenitors (Fig. 2H). We also observed that the PGCs migrate already from gastrulation stages associated with the mesothelium-primed LPM that establishes their ventral migration border (Fig. 4). Recent scRNA-seq studies of early mouse embryos have uncovered Hand2-expressing cell clusters that were deemed mesenchymal and that we now likely recognize as mesothelial progenitors by their complement of LPM-associated transcripts (Fig. S5) (Han et al., 2020; Pijuan-Sala et al., 2020). Altogether, these observations connect the coelomic epithelium progenitors to key aspects of body plan organization already in early development. The exact composition of a mesothelial progenitor-controlling program and its evolutionary conservation beyond the genes uncovered here warrants further attention.
Expanding previous loss-of-function analyses that documented Hand gene contribution to heart and forelimb development (Firulli et al., 1998; Firulli et al., 2010; Osterwalder et al., 2014; Yelon et al., 2000), we here add that mesothelial progenitors fail to properly migrate in hand2-mutant zebrafish, resulting in ventral herniation of the yolk at 3 dpf due to incomplete closure of the ventral body wall (Fig. 6). How loss of hand2 leads to dysregulated migration of the mesothelial progenitors warrants future efforts. In congenital malformations affecting the body wall including omphalocele and gastroschisis, the future ventral abdominal body wall fails to form properly, resulting in internal organ protrusion and herniation of abdominal muscles, among other phenotypes (Boylan et al., 2020; Brewer and Williams, 2004; Sadler and Feldkamp, 2008). Among the candidate factors involved in this process is TGF-β signaling that has been repeatedly associated with Hand2 function and midline migration (Gays et al., 2017; Laurent et al., 2017; Yin et al., 2010), as well as Six4 and Six5 (Takahashi et al., 2018). Further, Hand2 regulates extracellular matrix components including repression of Fibronectin production, disruption of which lead to migration defects in the heart and prospective visceral mesothelium (Garavito-Aguilar et al., 2010; Yin et al., 2010). Altogether, our data indicate that hand2 is not only active early on during mesothelium emergence but also functionally contributes to correct mesothelium formation in zebrafish.
Asbestos-induced mesothelioma remains a global health challenge (Carbone et al., 2019; Felley-Bosco and Macfarlane, 2018). While asbestos fibers have been causally linked to pleural mesothelioma (Wagner et al., 1962), the transformative mechanism and the exact cell of origin await clarification. Different patient stratifications have been assigned to mesothelioma with biphasic, epithelioid, or sarcomatoid tumors based on histopathology (Carbone et al., 2019). Nonetheless, only a few genetic markers are known, including YAP activity and altered BAP1 expression, and their role in mesothelioma formation and maintenance remains unclear (Carbone et al., 2019; Hmeljak et al., 2018). Work in mouse models has uncovered activation of stemness-related signaling pathways and an increase in Msln-expressing cells upon asbestos exposure (Rehrauer et al., 2018). Human epithelioid mesothelioma show upregulated expression of coelomic epithelium-associated genes, including of WT1 (Bozzi et al., 2016; de Reyniès et al., 2014). Notably, the non-coding RNA Fendrr (Fetal-lethal noncoding) is strongly upregulated in mouse and human epithelioid mesothelioma and shares a bidirectional promoter with the LPM transcription factor gene FOXF1 (Bueno et al., 2016; Felley-Bosco and Rehrauer, 2018; Grote et al., 2013). These observations indicate that at least some populations of mesothelioma cells feature a gene expression profile reminiscent of late developmental markers for the coelomic epithelium.
Aberrant reactivation of early developmental programs is increasingly recognized as contributor to tumorigenesis (Aiello and Stanger, 2016; Kaufman et al., 2016; Pomerantz et al., 2020). Extrapolating from our developmental findings in zebrafish, our data propose that mesothelioma might reactivate LPM-specific factors involved in initial mesothelial progenitor formation: in both mouse and human mesothelioma, we found the expression of Hand2 and additional transcription factor genes associated with early mesothelium-primed LPM (Fig. 2,7). Notably, crocidolite-exposed tissue showed intermediate expression levels for mesothelial progenitor genes, further supporting a model wherein mesothelioma formation involves a temporal re-activation of LPM-expressed genes deployed in initial mesothelium development (Fig. 7). While more work is warranted to establish whether and which LPM transcription factors contribute to mesothelioma formation and maintenance, our findings provide several mRNA transcript and protein expression assays to potentially further stratify mesothelioma in the clinic. We speculate that the malignant transformation of adult mesothelium, and in particular Hand2, endows altered cells with features that recapitulate the migration, proliferation, and stemness that distinguished mesothelial progenitor cells already during their initial development in the LPM.
Materials and Methods
Zebrafish husbandry
Zebrafish (Danio rerio) husbandry and experiments were performed according to the European Communities Council Directive (86/609/EEC), the recommendations of the Swiss authorities (Animal Protection Ordinance), and according to IACUC regulations at the University of Colorado School of Medicine, Anschutz Medical Campus. Protocols and experiments were performed as approved by the cantonal veterinary office of the Canton Zurich (Kantonales Veterinäramt, permit no. 150, TV4209), in accordance with the European Union (EU) directive 2011/63/EU as well as the German Animal Welfare Act, and as by the IACUC at the University of Colorado School of Medicine, Anschutz Medical Campus (protocol no. 00370). All zebrafish were raised, kept, and handled according to established protocols (Westerfield, 2007) if not noted otherwise.
Transgenic zebrafish lines and transgene activity
Established transgenic and mutant lines used in this study includes Tg(drl:EGFP) (Mosimann et al., 2015), Tg(drl:mCherryzh705) (Sánchez-Iranzo et al., 2018b), Tg(hand2:EGFP) (Kikuchi et al., 2011), Tg(pax2.1:EGFP) (Picker et al., 2002), Tg(lmo2:dsRed2) (Zhu et al., 2005), Tg(-6.8kbwt1a:EGFP) (Bollig et al., 2009), Tg(wt1b:EGFP), Tg(kop:mCherry-f’-nos3’UTR) (Blaser et al., 2005), Tg(drl:creERT2;alpha-crystallin:YFP) (Mosimann et al., 2015), Tg(−6.8kbwt1a:creERT2) (Sánchez-Iranzo et al., 2018a), Tg(–3.5ubb:loxP-EGFP-loxP-mCherry) (Mosimann et al., 2011), Tg(−1.5hsp70:loxP-STOP-loxP-EGFP;alpha-crystallin:Venus) (Felker et al., 2018),n and hand2 mutants (hanS6) (Yelon et al., 2000).
The construct to generate the transgenic line TgBAC(wt1b:rtTA-p2A-creERT2)cn19 (referred to in the text as wt1b:creERT) was generated by BAC recombineering using EL250 bacteria (Lee et al., 2001). Fragments were amplified by PCR, adding 50 nucleotide homology arms. First, the iTol2Amp-γ-crystallin:RFP cassette (Sánchez-Iranzo et al., 2018b) was amplified using primers 1. pTarBAC_HA1_iTol2_F 5’-gcgtaagcggggcacatttcattacctctttctccgcacccgacatagat CCCTGCTCGAGCCGGGCCCAAGTG-3’ and pTarBAC_HA2_iTol2CrystRFP_R 5’-gcggggcatgactattggcgcgccggatcgatccttaattaagtctact aTCGAGGTCGACGGTATCGATTAAA-3’ and recombined into the backbone of the BAC clone CH73-186G17 to replace the BAC-derived loxP site. Then, the rtTA-p2A-iCreERT2 cassette was amplified and recombined replacing the ATG of the wt1b coding sequence, with primers wt1b_HA1_rtTA_F 5’-gacattttgaactcagatattctagtgttttgcaacccagaaaatccgtc ACCATGGTCGACGCCACAACCAT-3’ and wt1b_HA2_FRT_R 5’-gcgctcaggtctctgacatccgatcccatcgggccgcacggctctgtc agGGAGGCTACCATGGAGAAG-3’. Finally, the Kanamycin resistance cassette was removed by inducing expression of Flipase recombinase in the EL250 bacteria. The final BAC was purified with the HiPure Midiprep kit (Invitrogen) and injected along with synthetic Tol2 mRNA into wildtype strain zebrafish embryos. Sequence information and primer details are freely available upon request.
6-18 somite staged (ss) dual-fluorescent embryos were imaged using a Leica SP8 upright confocal microscope with HCX-Apochromat W U-V-I 20x/0.5 water correction objective.
Mouse transgenic strains
The Hand1EGFPCreΔNeo/+ (Barnes et al., 2010), Hand2Cre (dHand-Cre) (Ruest et al., 2003) and ROSA26 Reporter (R26R) (Soriano, 1999) mouse strains have all been used as previously described. The Hand2Cre transgenic strain was produced using a 7.4 kb genomic fragment from the mouse Hand2 locus immediately upstream of the transcriptional start site. The Hand1EGFPCreΔNeo/+ strain was produced by knocking a Cre cassette into the mouse Hand1 locus.
Human tumor specimens
Mesothelioma tumor specimens from the previously described collection were used (Oehl et al., 2018). Non-tumor pleural tissue was received from four patients undergoing mesothelioma-unrelated thoracic surgery. The study was approved by the Institutional Ethical Review Board of the University Hospital Zurich under reference number StV 29-2009.
Zebrafish CreERT2-based lineage tracing
Genetic lineage tracing experiments were performed using the Cre/loxP recombination system with 4-OHT-inducible creERT2 transgenic lines. hsp70l:Switch reporter zebrafish (Felker et al., 2018) were crossed with creERT2 drivers drl:creERT2 (Mosimann et al., 2015), wt1a:creERT2, and wt1b:creERT2. Lineage-labeling was induced at indicated time points using fresh or pre-heated (65°C for 10 min) 4-Hydroxytamoxifen (4-OHT) (Sigma H7904) stock solutions in DMSO at a final concentration of 10 μM in E3 embryo medium (Felker et al., 2016). To activate the EGFP transcription in hsp70l:Switch strains, embryos were incubated at 37°C for 1 hour, 2-3 hours before fixation. Subsequently, the embryos were fixed in 4% paraformaldehyde (PFA) at 4°C overnight, washed and stored in PBS, and processed for imaging.
Zebrafish histology and sectioning
Transverse sections were performed as previously described (Gays et al., 2017). 4% PFA fixed zebrafish embryos were embedded in 6% low-melting agarose (LMA, Sigma-Aldrich) dissolved in 0.1% Tween-20 (Sigma-Aldrich) in PBS and dissected into 130 μm slices with a vibratome (Leica VT 1000S). For immunostaining, the sections were incubated for 30 minutes in permeabilization buffer (1% BSA (AMRESCO), 1% DMSO (Sigma-Aldrich), and 0.3% Triton X-100 (Sigma-Aldrich) in PBS) and washed with PBT (0.1% Triton X-100 and 1% BSA in PBS). Antibodies were dissolved in PBT and incubated overnight at 4°C (primary antibody) or for 4 hours at room temperature (secondary antibody). Primary antibody used was α-SM22 alpha antibody (ab14106). Secondary antibody used: goat-anti-rabbit 594 (Alexa Fluor, Life Technologies, 1:300). Sections were mounted with DAPI-containing Vectashield (Cat#H-1200, Vector Laboratories). Images were obtained with the Zeiss LSM710 confocal microscope equipped with Plan-Apochromat 40x/1.3 oil DIC M27 or 40x/1.2 immersion correction DIC M27 objectives. Images were adjusted for brightness and cropped using ImageJ/Fiji (Schindelin et al., 2012).
Embryos for whole-mount immunostainings were processed as described above for the sections. Primary antibodies used were anti-Pax2 (GeneTex, GTX128127, 1:250) and anti-Myosin heavy chain (DSHB, MF20, 1:20). Secondary antibodies used were Alexa Fluor conjugates (Invitrogen). Embryos were either imaged on a Leica SP8 upright confocal microscope or Zeiss Z.1 light sheet microscope (see details below).
Gene expression analysis and in situ hybridization (ISH)
First-strand complementary DNA (cDNA) was generated from pooled zebrafish RNA isolated from different developmental stages using SuperScript III first-strand synthesis kit (Invitrogen). DNA templates were generated using first-strand cDNA as a PCR template and the primers as specified for each gene of interest; for in vitro transcription initiation, the T7 promoter 5′-TAATACGACTCACTATAGGG-3′ was added to the 5′ ends of reverse primers. Specific primers per gene were meis3: forward 5’-TACCACAGCCCACTACCCTCAGC-3’, reverse 5’-TAATACGACTCACTATAGGGTCAGCAGGATTTG GTGCAGTTG-3’; gata5: forward 5’-CACCATGTATTCGAGCCTGGCTTTATCTTCC-3’, reverse 5’-TAATACGACTCACTATAGGGTCACGCTTGAGAC AGAGCACACC-3’; sfrp5: forward 5’-GAATCACAGCAGAGGATG-3’, reverse 5’-TAATACGACTCACTATAGGGCATCTGTACTAATG GTCG-3’; cdx4: 5’-CACCATGTATCACCAAGGAGCG-3’, reverse 5’-TAATACGACTCACTATAGGGTAATCCACAACCCA CGCC-3’. PCR reactions were performed under standard conditions as per manufacturer’s protocol using Phusion high-fidelity DNA polymerase (Thermo Fisher Scientific). RNA probes were generated via overnight incubation at 37°C using 20 U/µL T7 RNA polymerase (Roche) and digoxigenin (DIG)-labeled dNTPs (Roche) as per manufacturer’s protocol. The resulting RNA was precipitated in lithium chloride and EtOH. Wildtype strain zebrafish embryos were fixed in 4% PFA in PBS overnight at 4°C, dechorionated, transferred into 100% MeOH, and stored at −20°C. ISH of whole-mount zebrafish embryos was performed essentially as per standard protocols (Thisse and Thisse, 2008).
RNAscope assay
The RNAscope assays were carried out based on a previously published protocol (Gross-Thebing et al., 2014) and the manufacturer’s instruction (Advanced Cell Diagnostics). Wildtype strain or transgenic zebrafish embryos of tailbud stage until early somitogenesis were fixed in 4% PFA in PBS for 1 h at RT or overnight at 4°C, hand dechorionation, and transferred into 100% MeOH and stored at −20°C. Protease digestion of embryos using Protease III was performed for 20 min at RT followed by rinsing the embryos three times in 0.01% PBS-T (0.01% Tween-20). Target probe hybridization was performed overnight at 40°C. Target probes were designed by Advanced Cell Diagnostics for hand2, foxh1, and meis3. For RNA detection, incubation with the different amplifier solutions was performed in an incubator at 40°C. After each hybridization step, the embryos were washed three times with 0.2x SSCT for 15 min. The embryos were then incubated with ACD’s DAPI solution overnight at 4°C. Prior to imaging, embryos were rinsed in 0.01% PBT.
Tumor RNA extraction, cDNA synthesis, and qPCR
RNA extraction, cDNA synthesis, and qPCR were performed as previously described (Thurneysen et al., 2009). The expression of HAND2 was determined using the following primers: forward 5’-CAGGACTCAGAGCATCAACAG-3’ and reverse 5’-TCCATGAGGTAGGCGATGTA-3’. Normalization was performed based on the expression of Histone H3 detected with primers 5’-GGTAAAGCACCCAGGAAGCA-3’ and 5’-CCTCCAGTAGAGGGCGCAC-3’ (Andre and Felley-Bosco, 2003). Data were analyzed using Mann-Whitney test.
Tumor immunohistochemistry
Immunohistochemistry was performed as previously described (Frei et al., 2011) using anti-HAND2 antibody (A-12, Sc-398167, 1:100) and secondary universal antibody (Vectastain, Vector laboratories Inc., Burlingame, CA, USA), according to manufacturer instructions. Control staining was performed by omission of the primary antibody.
Light sheet sample preparation, microscopy, and image analysis
Embryos used for long-term imaging or imaging after 1 dpf were treated with 0.003% 1-phenyl-2-thiourea (PTU, Sigma-Aldrich) to prevent melanin pigment formation.
Panoramic SPIM for hand2:EGFP;drl:mCherry imaging during somitogenesis
At 50% epiboly, embryos in the chorion were mounted in an FEP tube into 1% LMA. Up to 6 embryos were positioned on top of each other with a minimum gap between them. The FEP tube was mounted in the microscope imaging chamber filled with E3 medium. Time-lapse acquisition was performed with a four-lens SPIM set up by a standardized image acquisition pipeline (Schmid et al., 2013). The microscope was equipped with four identical Olympus UMPlanFL N 10x/0.3 NA detection objectives. The subsequent real-time image processing, registration of time points, and 2D map (Mercator) projections were performed with previously described and published Fiji scripts (Schmid et al., 2013). A Z-stack was obtained from every embryo with an interval of 2 min for a period of 14–17 h. Images were processed using ImageJ/Fiji.
Multidirectional SPIM (mSPIM) for long-term hand2:EGFP;drl:mCherry imaging
To prevent movement of zebrafish embryos during imaging, we injected 30 pg of α-bungarotoxin mRNA (Swinburne et al., 2015) at the 1-cell stage. At the 20 ss, embryos that showed no sign of muscular contractions were selected and mounted in FEP tubes filled with 0.1% LMA. A detailed mounting protocol can be found in Daetwyler et al., 2019. Long-term time-lapse imaging was performed using a custom multidirectional SPIM (mSPIM) setup with multi-sample capacity (Daetwyler et al., 2019). The microscope was equipped with two Zeiss 10x/0.2 NA illumination objectives and an Olympus UMPlanFL N 10x/0.3 NA detection objective. The embryos were imaged every 20 min for up to 4 days starting around 20 ss from 4 different angles. Custom-made data processing tools were used as previously described (Daetwyler et al., 2019) to stitch the individual volumes per angle. For successful fusion and deconvolution of the 4 individual angles, we manually registered them, and then applied the Fiji “Multiview Reconstruction” and “BigStitcher” plugins (Hörl et al., 2019; Preibisch et al., 2010; Preibisch et al., 2014) for precise registration and deconvolution. ImageJ/Fiji and Imaris (Bitplane AG, Zurich, Switzerland) were used for visualization.
Zeiss Z.1
The Zeiss Z.1 microscope equipped with a Zeiss W Plan-Apochromat 20x/0.5 NA objective was used for all other light sheet microscopy and accordingly mentioned in the figure legends. Embryos were either within or out of the chorion embedded in 1% LMA in respectively a 50- or 20-µL glass capillary. Alive embryos older than 16 ss were additionally mounted with 0.016% ethyl 3-aminobenzoate methanesulfonate salt (Tricaine, Sigma) in the LMA and added to the E3 medium to prevent movement during imaging. For live imaging of the heart, the heartbeat was stopped with 30 mM 2,3-butanedione monoxime (BDM, Sigma). Images were processed using ImageJ/Fiji and Imaris.
Map projections drl:EGFP;kop:mCherry-F-actin-nanos-3’UTR
All pre-processing steps and map projection were performed using custom-developed algorithms in MATLAB, as also described (Naganathan et al., 2020). The overall size and shape of the fused embryo were first determined in an image that was binned four times to increase the processing speed. The binned image was binarized using an adaptive threshold (imbinarize function with a sensitivity of 0.4) and the isosurface of the resulting image was determined. A sphere fit was then performed using linear least squares with the ellipsoid_fit function from Mathworks (https://ch.mathworks.com/matlabcentral/fileexchange/24693-ellipsoid-fit). The center and radius of the sphere were then used for performing Mercator map projection of the fused images.
For each fused image, a multi-layered projection was performed by generating concentric circles with different radii (step size: 2 µm). This allowed for unwrapping different layers of the 3D fused embryo onto 2D surfaces. For each layer of the map projection, a dummy image in polar coordinates was first generated that extended from -90° to +90° in latitude and 0° to 360° in longitude. Using Mercator projection formulas, the latitudes and longitudes that correspond to each position in the projected map were then determined. The cartesian (x,y,z) positions that correspond to each of these (latitude, longitude) points in the projected map was obtained using standard spherical to cartesian coordinate system conversion formulas. Once this was obtained, a direct mapping of the pixel values corresponding to an x,y,z position in the fused image onto the projected map was performed yielding the map-projected image.
Single-cell RNA sequencing of LPM
Sample preparation and sorting
Wildtype stain and drl:mCherry-positive zebrafish embryos were grown until tailbud stage and chorions were removed by incubating in 1 mg/mL Pronase (Roche), followed by washing in E3 medium. Around 300 embryos were used to generate the sample. The E3 medium was replaced by 1X PBS and the embryos were stored on ice until further processing. The embryos were dissociated using 2 mg/mL collagenase IV (Worthington) in DMEM (high glucose (4.5g/l) and NaHCO3, without L-glutamine and sodium pyruvate, Sigma-Aldrich) and incubated for 5 min in a water bath at 37°C. The embryonic tissues were triturated into a single-cell suspension by pipetting carefully up and down. When the embryos were not yet dissociated sufficiently, they were incubated for another 5 min. Cells were filtered through a 35 μm cell-strainer (Falcon, round-bottom tubes with cell-strainer cap) and centrifuged at 6000 rpm for 30 sec. Cell pellets were resuspended in 1X HBSS (Gibco) containing 2% FBS and subjected to an additional round of centrifugation and resuspension. After washing, the cells were resuspended in 1X PBS.
drl:mCherry-positive cells were sorted using a FACS Aria III cell sorter (BD Bioscience). Cells were gated based on size and forward scattering, to exclude debris and doublets. The gating for the negative population was determined based on wildtype tailbud stage embryos. See also Supplementary Figure 2 for gate setting. The SORTseq single-cell RNA-sequencing protocol was carried out as described previously (Muraro et al., 2016). Live mCherry-positive single cells were sorted in four 384-well plates (BioRad) containing 5 μl of CEL-Seq2 primer solution in mineral oil (24 bp polyT stretch, a 4 bp random molecular barcode (UMI), a cell-specific barcode, the 5′Illumina TruSeq small RNA kit adaptor and a T7 promoter), provided by Single Cell Discoveries. After sorting, the plates were immediately placed on ice and stored at −80°C.
Single Cell Discoveries further processed the four plates. In brief, ERCC Spike-in RNA (0.02 μL of 1:50000 dilution) was added to each well before cell lysis with heat shocking. Reverse transcription and second-strand synthesis reagents were dispensed using the Nanodrop II (GC biotech). After generation of cDNA from the original mRNA, all cells from one plate were pooled and the pooled sample was amplified linearly with in vitro transcription. The amplified RNA was then reverse transcribed to cDNA using a random hexamer primers (Hashimshony et al., 2016). To generate sequencing libraries, RPI-series index primers were used for library PCR. Libraries were sequenced on an Illumina Nextseq500 using 75 bp paired-end sequencing.
Single-cell analysis
The CEL-Seq2 barcoded sequences were obtained from Single Cell Discoveries. STAR v2.5.3a (Dobin et al., 2013) was used to create an index based on the Ensembl GRCz10.91 genome and annotation, after manually adding drl:mCherry. Gene abundances were estimated separately for each plate using zUMIs v0.0.4 (Parekh et al., 2018), retaining cells with at least 100 reads and setting the Hamming distance threshold to 1 for both UMI and cell barcode identification. Finally, the exonic UMI counts for all four plates were merged into a single matrix.
Quality control and filtering were performed using the scater R package (McCarthy et al., 2017). Upon removal of genes that were undetected across all cells, we removed cells whose percentage of mitochondrial genes fell beyond 2 Median Absolute Deviations (MADs) of the median. Secondly, features with a count greater than 1 in at least 10 cells were retained for downstream analysis. Finally, we discarded cells measured on plate 4, as it was of overall low quality (low number of counts, high percentage of mitochondrial genes).
Next, we used Seurat (Stuart et al., 2019) for clustering and dimension reduction. Clustering was performed using the 2000 most highly variable genes (HVGs) identified via Seurat’s FindVariableFeatures function with default parameters; clustering and dimension reductions were computed using the first 20 principal components. For clustering, we considered a range of resolution parameters (0.2-2.4); downstream analyses were performed on cluster assignments obtained from resolution 1.8 (15 subpopulations).
Cluster annotations were performed manually on the basis of canonical markers in conjunction with marker genes identified programmatically with Seurat’s FindAllMarkers function, and complimentary exploration with iSEE (Rue-Albrecht et al., 2018).
For Supplementary Figure 4, the three endoderm clusters were excluded and the remaining clusters manually merged into 6 major subpopulations. For each subpopulation, genes that were differentially expressed (DE) against at 3 others were identified using scater’s findMarkers function with pval.type = “some” (McCarthy et al., 2017). The top 50 DE genes (in terms of effect size) at FDR < 5% and with an average positive log-fold change (summary.logFC > 0) were selected for visualization.
Software specifications and code availability
All analyses were run in R v4.0.2, with Bioconductor v3.11. Data preprocessing and analysis code are deposited at DOI:10.5281/zenodo.4267898, and available as a browsable workflowr website (Blischak et al., 2019). All package versions used throughout this study are captured in the session info provided therein.
Mesothelioma RNA-sequencing data
Mouse model and sample preparation
As previously described, mice were repeatedly intra-peritoneally over a time course of 21 weeks injected with crocidolite or sham-exposed and sacrificed at 33 weeks, 12 weeks after the last crocidolite exposure (Rehrauer et al., 2018) (Fig. 7). Tissues, among which mesothelium and tumor masses, were collected from euthanized mice and consecutively processed for histopathological diagnosis and analysis and for RNA-seq analysis. The mice sacrificed corresponded to three groups: sham, crocidolite-exposed with pre-neoplastic lesions, and mice exposed to crocidolite-bearing tumors.
Mouse RNA-seq analysis
The RNA isolation, library generation, and RNA-seq analysis pipelines are previously described (Rehrauer et al., 2018), In this study, we extracted the expression values of genes associated with early LPM and mesothelium development and diagnostic markers (Msln, Wt1, Bap1, Nf2). Counts were normalized with DESeq2’s estimateSizeFactors function (Love et al., 2014); heatmap values correspond to log10-transformed, scaled and centered (scale with default parameters) normalized counts. Right-hand side row annotations display (unscaled) log10-transformed count ranges.
Human RNA-seq analysis
Analysis of human mesothelioma was performed using TCGA data downloaded from cBioportal (www.cBioportal.org) (Gao et al., 2013). To perform unsupervised clustering analysis on the mRNA of TCGA mesothelioma samples, we used the complexHeatmap R package (Gu et al., 2016). Heatmap values correspond to log10-transformed, batch-normalized mRNA counts.
Funding
This work has been supported by Swiss National Science Foundation (SNSF) professorship PP00P3_139093 and SNSF R’Equip grant 150838 (Light sheet Fluorescence Microscopy), Marie Curie Career Integration Grant from the European Commission [CIG PCIG14-GA-2013-631984], the Canton of Zürich, the UZH Foundation for Research in Science and the Humanities, the Swiss Heart Foundation, the ZUNIV FAN/UZH Alumni, the University of Colorado School of Medicine, Department of Pediatrics and Section of Developmental Biology, and the Helen and Arthur E. Johnson Foundation and Children’s Hospital Colorado Foundation to C.M.; a UZH CanDoc to S.N.; EuFishBioMed and Company of Biologists travel fellowships to K.D.P; SNSF postdoc mobility fellowship P400PB_191057 to J.K.R.; SNSF grant 320030_182690 and Stiftung für Angewandte Krebsforschung to E.F.B.; SNSF 310030L_182575 and ERC Starting Grant 2013 337703 to N.M.; SNSF grants 310030_175841 and CRSII5_177208 and University of Zurich’s Research Priority Program Evolution in Action to M.D.R..
Author contributions
K.D.P. and C.M. conceived the project and designed the study. K.D.P. performed the zebrafish experiments together with E.C.B. and Z.L. K.D.P., S.N., and A.K. performed and interpreted gene expression analysis in zebrafish embryos. K.D.P. and S.N. prepared and interpreted the scRNA-seq. H.L.C., C.S., and M.D.R. processed and analyzed the scRNA-seq data. S.D. and K.D.P. acquired data with multidirectional SPIM (mSPIM) and processed the data. A.E. and H.S. generated wt1b:creERT2. D.E.C and A.B.F. performed mouse lineage tracing experiments. M.R. performed the IHC of mouse and human tumor samples, M.R. and J.K. performed qPCR on mesothelioma human patient samples, E.F.B., M.R., and J.K. analyzed the mouse and human samples, J.K. and R.O. analyzed TCGA data. E.R., N.M., J.H., E.F.B., A.B., M.D.R., and C.M. supervised. K.D.P. compiled the data and K.D.P. and C.M. wrote the manuscript with input from all co-authors, E.F.B wrote sections of the manuscript related to mesothelioma.
Competing interests
The authors declare no competing interests.
Data and material availability
All sequencing datasets generated for this publication will be deposited on the ArrayExpress database upon publication; all reagents used are available upon request.
Raw scRNA-seq data, intermediate, and metadata files to reproduce all analyses and figures are available at DOI:10.6084/m9.figshare.13221053.v1.
Movies
Movie 1: Primordial germ cells are migrating within the drl-positive LPM during gastrulation and early somitogenesis
Multiview time-lapse SPIM (Zeiss Z.1) of a representative embryo expressing drl:EGFP and primordial germ cell (PGC) marker nos-3’UTR:mCherry from 6 hpf until 14 hpf. Already during gastrulation, the PGCs migrate within the drl-expressing mesendoderm and concentrate within the LPM during early somitogenesis. Note the cells that end up more posterior in the most lateral territory of the drl-positive LPM.
Movie 2: hand2 is expressed in the forming mesothelial layers
Lateral view of dual-color time-lapse imaging of a hand2:EGFP;drl:mCherry embryo from 18 hpf until 82 hpf. drl:mCherry drives expression in mainly the circulating blood cells from 24 hpf onwards. EGFP expression is restricted to the mesothelial layers, pectoral fin, pharyngeal arches, and heart (endo-, myocardium).
Movie 3: hand2-expressing cells form the pericardium and visceral and parietal peritoneum
Zebrafish embryo expressing hand2:EGFP imaged from four different angles over a time course from 18 hpf until 82 hpf. Full 3D-rendering shows how the EGFP-expressing cells migrate over the yolk and yolk extension to form the parietal peritoneum and the pericardium. A front of cells with a medial-directed migration pattern forms the visceral peritoneum along the anterior-to-posterior axis.
Acknowledgments
We thank Dr. Sundar Naganathan for help with the Matlab scripts for the panoramic projections. We thank Single Cell Discoveries for executing the single-cell RNA-sequencing, as well as the Center for Microscopy and Image Analysis (ZMB) and the Cytometry Facility (FCF) of the University of Zurich for technical support and equipment. We also thank Dr. Deborah Yelon, Dr. Richard M. White, Dr. Charles K. Kaufman, and the members of the Mosimann lab for critical input on the manuscript. Species schematics in Figure 1 and Figure 6 were adjusted from templates provided by PhyloPic (phylopic.org).