Mechanical stretch scales centriole number to apical area via Piezo1 in multiciliated cells

Organelles compartmentalize cells into discrete functioning units. Cells must regulate the number of organelles to achieve proper function (Marshall, 2007; Marshall, 2016; Nigg and Holland, 2018; Rafelski and Marshall, 2008). For example, multiciliated cells (MCCs) line the epithelia of the brain ventricles, the airway, and the oviduct where motile cilia propel extracellular fluid to circulate cerebrospinal fluid, remove pathogens, and move the ova (Spassky and Meunier, 2017). Depending on the location, MCCs synthesize between 30 and 300 motile cilia (Spassky and Meunier, 2017). Assembly of too few or too many cilia impairs MCC function and is associated with several diseases including Primary Ciliary Dyskinesia, suggesting the existence of an active mechanism that controls cilia number (Boon et al., 2014; Spassky and Meunier, 2017; Wallmeier et al., 2014). Yet, the cellular and molecular mechanisms that control the number of cilia in MCCs remain unknown.

To shed light on mechanisms, we used the Xenopus embryonic epidermis, an established, versatile, in vivo model to study MCCs (Walentek and Quigley, 2017; Werner and Mitchell, 2013). There, MCCs are first specified in the basal epithelia, where they begin to synthesize centrioles using specialized structures called deuterosomes (Figure 1a, Step 0) (Klos Dehring et al., 2013; Zhao et al., 2013). As MCCs intercalate into the outer epithelial cell layer and expand their apical surface, centrioles migrate apically, dock at the apical surface, and provide the platform for assembly of motile cilia (Figure 1a, Steps 1–4) (Deblandre et al., 1999; Kulkarni et al., 2018a; Stubbs et al., 2006; Zhang and Mitchell, 2015). As such, in this study, we focused our efforts on counting centrioles as a simple, efficient proxy for cilia number using chibby-GFP, a marker for mature centrioles (Burke et al., 2014).

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MCC development is closely linked to embryonic development (Figure 1a). Therefore, we collected embryos at different developmental stages (from stage 20 to stage 28) to examine MCCs at various stages of apical expansion, ranging from MCCs that have just intercalated (Step 2) to fully mature MCCs (step 4). We measured the number of centrioles at the apical surface and the apical area. Surprisingly, we observed a strong correlation between the apical area and the number of centrioles at the apical surface (Figure 1b,c).

Next, we wanted to test if this relationship would persist if the MCC apical area became larger. We employed four different approaches to increase the apical area of MCCs. First, two species of Xenopus are common models for cell biology (X. laevis and X. tropicalis), and due to the evolutionary variation in embryonic sizes of the two species, they are useful for scaling experiments (Figure 1d – compare relative sizes of the eggs and embryo) (Levy and Heald, 2012). Compared to X. tropicalis, the X. laevis embryo is larger with significantly larger MCCs (Figure 1d–g, median ± SD, 391 ± 86 μm² vs. 270 ± 53 μm² in X. tropicalis). X. laevis MCCs also have significantly more centrioles (Figure 1h, median ± SD, 195 ± 27 compared to 150 ± 20 in X. tropicalis). Interestingly, by combining data from both species, we observed a clear trend where centriole number scales with MCC apical area (Figure 1i).

Second, in X. laevis, we converted epithelial goblet cells (which normally secrete mucus) to MCCs by overexpressing the master regulator of multiciliogenesis, mcidas (multiciliate differentiation and DNA synthesis-associated cell cycle protein) (Figure 1—figure supplement 1a; Stubbs et al., 2012). These induced MCCs in X. laevis are larger than X. tropicalis MCCs and have proportionately more centrioles (Figure 1—figure supplement 1b,c). Interestingly, the apical area of mature MCCs and the induced MCCs of X. laevis were similar and so were the number of centrioles (Figure 1—figure supplement 1c).

Third, we induced cytokinesis defects by knocking down ccdc11 using a morpholino oligo (MO) in X. tropicalis (Kulkarni et al., 2018b). MCCs are mitotically mature; however, if their progenitor fails to undergo cytokinesis, then the resultant MCC can be much larger (Figure 1—figure supplement 2a). With this strategy, we identified MCCs with significantly larger apical areas (median ± SD, 437 ± 154 μm² vs. 202 ± 38 μm² in controls) (Figure 1—figure supplement 2a,b), and these cells had proportionately more centrioles (median ± SD, 227 ± 74 vs. 138 ± 26 in controls) (Figure 1—figure supplement 2c,d).

Finally, in X. laevis, we fused MCCs with neighboring (most likely) non-MCCs (confirmed by the presence of two nuclei – dashed lines in Figure 1—figure supplement 2e), resulting in much larger cells with increased apical area (median ± SD, apical area of 534 ± 149 μm² vs. 337 ± 49 μm² in controls) and more centrioles (median ± SD, 235 ± 40 vs. 168 ± 21 in controls) (Figure 1—figure supplement 2e–h). Interestingly, in each experiment, centriole number increased in proportion to the apical area suggesting that centriole amplification is a plastic process and cells can calibrate centriole number in response to cell size perturbations. By combining the data from controls and manipulated embryos, we found that this scaling relationship could be observed over a 40-fold change in the apical area, with the smallest apical area being ~25 μm² and the largest about 1000 μm² (Figure 1j). However, these experiments are limited in two ways. First, in our experiments, we increased the entire volume of the cell not just the apical area. Additionally, in cells with a cytokinesis defect or cell-cell fusion, we have combined two or more cells leading to an increase in the centriole number. Nevertheless, despite these limitations, the correlation between apical area and centriole number appears robust, and in subsequent experiments, we strived to overcome these limitations.

While these experiments suggest that the apical area of an MCC may regulate centriole number, we sought to test the alternative hypothesis that centriole number may determine apical area or that there may be a feedback mechanism between centriole number and apical area. We can manipulate centriole number in two ways: increase the number of centrioles by overexpressing cep152 (Collins et al., 2020; Klos Dehring et al., 2013) or decrease the number of centrioles with Centrinone treatment, a PLK4 inhibitor (Wong et al., 2015). We first increased the centriole number by overexpressing cep152 in X. laevis (Figure 2a), which increased the number of centrioles (median ± SD, 467 ± 113 vs. 160 ± 27 in controls) and was accompanied by a correlated increase in the apical area (median ± SD, 692 ± 296 μm² vs. 236 ± 58 μm² in controls) (Figure 2a–d). Next, we reduced centriole numbers with Centrinone. Centriole synthesis begins during intercalation, when the cells are in the basal layer (Figure 1a, Step 0). To allow the chemical inhibitor to access the cells in the basal layer, we generated Xenopus embryonic ‘stem cell’ explants (commonly referred to as animal caps), which auto-differentiate into an embryonic epidermis replete with MCCs (Figure 2e). We harvested animal caps from X. tropicalis embryos and grew them on fibronectin-coated slides with exposure to Centrinone or vehicle alone until control embryos reached stage 25–26 (Figure 2e,f). We successfully reduced the median number of centrioles from 104 in controls to 25 in Centrinone-treated MCCs (Figure 2f,g). Despite a dramatic reduction in the number of centrioles, we found a slight increase in the apical area of Centrinone-treated MCCs as compared to controls (median ± SD, 175 ± 67 vs. 141 ± 38 μm² in controls) (Figure 2h). From this result, we conclude that a minimum apical size can be achieved independent of the centriole amplification (Figure 2f–i). Once this minimum size is reached, then centrioles may contribute to apical expansion (Figure 2a–d). Moving forward, we focused on the hypothesis that the apical area may fine tune centriole number.

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While we initially focused on the number of centrioles at the apical surface, this may not reflect the total number of centrioles in the cell. Previous studies have noted the presence of centrioles in the intercalating MCCs (Figure 1a, Steps 0–1), but the number of these centrioles is unknown (Collins et al., 2020; Werner et al., 2014). One possibility is that MCCs assemble all of the centrioles (~150 in X. tropicalis) during intercalation with subsequent waves of either synthesis or degradation depending on the final apical area. Alternatively, MCCs may continuously produce centrioles and halt this process based on the final apical area. In either model, MCCs would require a cellular and molecular mechanism to measure the apical area.

To differentiate between these hypotheses, we set out to image centrioles in the intercalating cells. To image deep within the cytoplasm, we used animal caps which are relatively transparent compared to whole embryos and imaged centrioles (Chibby-GFP) along the apico-basal axis of intercalating MCCs. Using segmentation and 3D reconstruction, we could observe the process of centriole migration and count all the centrioles in the cytoplasm of intercalating MCCs (presumptive MCC border marked by white dotted line, Figure 3a–d, Videos 1 and 2). In these intercalating cells, the number of centrioles was 75 (median), approximately half the number in mature X. tropicalis MCCs (Figure 3e, blue, Step 4; magenta, Steps 0–1). Interestingly, X. laevis MCCs also make 90 (median) centrioles during intercalation, again half the number of centrioles in mature MCCs (Figure 3f blue, Step 4; magenta, Steps 0–1). From these results, we conclude that (1) Xenopus MCCs synthesize half the total number of centrioles prior to intercalation, (2) these centrioles dock to the apical surface in a manner that scales with apical area, and (3) the remaining half of the number of centrioles must be synthesized in a second round that is regulated based on the apical area.

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The apical area of the cell is dependent on multiple parameters including but not limited to the overall size of the cell, cell autonomous pushing forces, as well as pulling forces by neighboring cells (Guillot and Lecuit, 2013; Heisenberg and Bellaïche, 2013; Mao and Baum, 2015; Sedzinski et al., 2016). Specifically, during intercalation, the MCC is thought to cell autonomously push against its neighbors to expand its apical surface. Subsequently, neighboring cells pull on the MCC at cell junctions, expanding the apical area further (Sedzinski et al., 2016). To elucidate the contributions of pushing vs. pulling forces, we decided to examine the shape of the cells during apical expansion. Cell autonomous pushing forces would be radially symmetric so the apical surface should expand circularly (Sedzinski et al., 2016). On the other hand, cell non-autonomous pulling forces would depend on the relative positions of the neighboring cells and cell junctions leading to a polygonal apical shape (Sedzinski et al., 2016). The thinness ratio (TR) which relates the area of a shape to the square of its perimeter can detect these changes in cell shapes (Figure 4a). The TR is 1 for a circle and < 1 for polygons (Figure 4a). When we plotted the TR as a function of apical area, we found that MCCs with small apical areas have a TR of nearly 1, while the TR decreases to 0.8 as the apical area increases to ~300 μm² (Figure 4b,c, Video 3). Therefore, TR measurements support the notion that the initial apical expansion is driven by cell autonomous pushing forces, while subsequent apical expansion is driven largely by cell non-autonomous pulling forces.

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We speculated that cell non-autonomous pulling forces that drive the final phase of apical expansion might define the apical area and centriole number in MCCs. To test the hypothesis, we began with an embryological approach to manipulate the apical area. In a developing embryo, morphogenetic movements create forces that lead to dramatic shape changes that transform a spherical embryo (stage 9–10) to an elongated one (stage 28), presumably, exerting stretching forces on the epidermal MCCs to increase their apical area (Figure 1a). For example, Spemann’s Organizer, which is dependent on Wnt signaling, dorsalizes the mesoderm and ectoderm, which subsequently creates considerable elongation forces (De Robertis et al., 2000; Harland and Gerhart, 1997; Hikasa and Sokol, 2013; Keller and Sutherland, 2020; Kiecker, 2000). By depleting β-catenin, a key effector of the Wnt signaling pathway, we can eliminate the formation of Spemann’s Organizer and generate cylindrically symmetric embryos that lack dorsal structures and have much less elongation compared to control embryos (Figure 4—figure supplement 1a,c, Heasman et al., 1994; Khokha et al., 2005). While β-catenin-depleted embryos can form functioning MCCs that generate fluid flow, both the MCCs (median ± SD, 106 ± 26 μm² vs. 267 ± 64 μm² in controls) and non-MCCs (median ± SD, 319 ± 104 μm² vs. 428 ± 125 μm² in controls,) have smaller apical areas (Figure 4—figure supplement 1b,d–f). Interestingly, the centriole number in these embryos is also significantly decreased (median ± SD, 100 ± 16 vs. 148 ± 26 in controls, Figure 4—figure supplement 1g,h), approaching the 75 centrioles formed prior to intercalation. Further, the TR in β-catenin-depleted MCCs is significantly higher and closer to 1 (median ± SD, 0.93 ± 0.05 vs. 0.77 ± 0.04 in controls, Figure 4—figure supplement 1i,j), supportive of a significant reduction of pulling forces exerted on MCCs. This result suggests that the lack of embryonic elongation forces in β-catenin-depleted embryos causes the reduction in the MCC apical area and centriole number. However, a challenge in this experiment is the confounding effects generated by genetic manipulations, such as diminished Wnt signaling or potential changes in cell adhesion in β-catenin depleted embryos.

To avoid these confounding effects, we sought to manipulate MCCs using non-genetic tools. We returned to animal caps, Xenopus stem cell explants that auto-differentiate into an embryonic multiciliated epidermis and raised them in two different conditions. In the first condition, we harvested animal caps and cultured them on agarose. In this case, because the cells do not adhere to agarose, the animal caps roll up to form irregular spherical structures which we called ‘untethered’ explants (Figure 4—figure supplement 2a,b, Video 4). The MCCs in these explants had an apical area just slightly larger than in β-catenin-depleted embryos (median ± SD, 116 ± 37 μm² compared to 106 ± 26 μm² in β-catenin-depleted embryos and 263 ± 46 μm² in controls, Figure 4e,f,i) and a correlated decrease in centriole number (median ± SD, 105 ± 27 compared to 100 ± 16 in β-catenin-depleted embryos and 149 ± 14 in controls, Figure 4j). In the second condition, we harvested animal caps and cultured them on fibronectin-coated slides. In this case, the cells adhere to the slide and spread outward (Stepien et al., 2019). As a result, these ‘tethered’ explants are stretched along the slide to form flat epithelia (Figure 4—figure supplement 2c,d, Video 5; Stepien et al., 2019). In these tethered explants, the apical area of both non-MCCs (Figure 4—figure supplement 3, median ± SD, 372 ± 159 μm² compared to 170 ± 86 μm² in untethered caps and 465 ± 177 μm² in controls) and the MCCs (Figure 4e–g,i, median ± SD, 210 ± 43 μm² compared to 116 ± 37 μm² in untethered caps compared to 263 ± 46 μm² in controls) are increased compared to the untethered caps but are slightly smaller than epithelial cells in the embryo suggesting that additional forces or factors in the embryo may contribute to the apical area. Nevertheless, the tethered caps had a significant increase in the number of centrioles (Figure 4j, median ± SD, 130 ± 25 vs 105 ± 27 in untethered caps compared to 149 ± 14 in controls) in an area-dependent manner.

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To understand the contribution of pulling forces in defining the apical area, we analyzed cell shapes and measured the TR. Specifically, by binning the data based on our results, from 0 to 100 μm² (apical areas of MCCs in the initial stages of development, step 2, median ± SD, TR: 0.90 ± 0.05), 100–150 μm² (untethered caps, median ± SD, TR: 0.89 ± 0.05), 150–250 μm² (tethered caps, median ± SD, TR: 0.79 ± 0.06), 250–350 μm² (wildtype X. tropicalis MCCs, median ± SD, TR: 0.79 ± 0.06), the TR reduces significantly as the cells become larger, supporting the increasing contribution of pulling forces on defining the apical area (Figure 4d). Taken together, these results suggested that tension generated by stretching within the epithelial sheet is critical to achieve proper apical area and triggers centriole amplification over the initial set of 75 centrioles.

To directly test the role of stretching force on centriole number, we applied an artificial radial stretch to the explants. Specifically, we raised X. tropicalis explants on a silicone membrane coated with fibronectin until sibling control embryos reached stage 26. At this stage, MCCs are nearly mature, and we stretched the explants radially for 3 hr in a stepwise fashion (Figure 4—figure supplement 2e,f). This stepwise stretch created a force of 11.5 N and about 50–75% strain. We observed a significant increase in the apical area of MCCs (median ± SD, 409 ± 57 μm² compared to 210 ± 43 μm² in unstretched tethered caps) (Figure 4g–i). Stretching also led to a dramatic change in cell shape and a further significant reduction in the TR (apical area: 351–600 μm², median ± SD, TR: 0.71 ± 0.1, Figure 4c,d) compared to both tethered unstretched caps (median ± SD, TR: 0.79 ± 0.6) and WT X. tropicalis MCCs (median ± SD, TR: 0.79 ± 0.6), consistent with the expectation that external stretching will lead to more polygonality of apical shape. In these stretched MCCs, the number of centrioles also increased (median ± SD, 199 ± 33 vs 130 ± 25 in unstretched tethered caps) demonstrating that stretching forces trigger centriole amplification in an area dependent manner in MCCs (Figure 4j). Interestingly, just by stretching, we transformed MCCs in X. tropicalis explants to sizes more similar to X. laevis (median ± SD, 409 ± 57 μm² vs. 390 ± 87 μm² in X. laevis) and the number of centrioles generated were also similar (median ± SD, 199 ± 33 vs. 195 ± 27 in X. laevis), highlighting the conserved role of mechanical forces in establishing the scaling mechanisms across species (Figure 1g–j, Figure 4k).

Given the central role stretching plays in regulating centriole number, we decided to investigate the molecular mechanisms that sense the force. While there are several molecules that can act as mechanosensors (Luo et al., 2013; Martino et al., 2018; Wang, 2017), we were particularly struck by the punctate distribution pattern of Piezo1 (Figure 5—figure supplement 1a). Piezo1 is a mechanosensitive cation channel that responds directly to membrane stretch and is primarily expressed in epithelial cells exposed to fluid pressure and flow (Bagriantsev et al., 2014; Wang and Xiao, 2018; Wu et al., 2017). In addition to its expression at cell junctions (Figure 5—figure supplement 1b – dashed box), we unexpectedly discovered that Piezo1 is localized adjacent to the centrioles at the apical membrane (Figure 5a, Figure 5—figure supplement 1a,b). Piezo1 localization is diminished with MO-based Piezo1 depletion indicating that this anti-Piezo1 antibody signal is specific (Figure 5—figure supplement 1c–e).

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To test the role of Piezo1 in regulating centriole number in MCCs, we depleted Piezo1 using MO and CRISPR and also inhibited its activity using GSMTx4, a spider venom peptide that inhibits cationic mechanosensitive channels including Piezo1 (Gnanasambandam et al., 2017). Centrioles were measured in mature MCCs and all three treatments resulted in a significant decrease in centriole number compared to control embryos (Figure 5b,d–g, median ± SD, 105 ± 22 piezo1 MO, 116 ± 24 piezo1 CRISPR, and 110 ± 26 GSMTx4, compared to 140 ± 21 in uninjected controls and 133 ± 22 in standard MO controls). This reduction in centriole number did not appear to be due to a defect in docking as we did not detect centrioles inside the cell. Further, Piezo1 depletion also uncoupled the relationship between apical area and centriole number as evident by the increase instead of a decrease in the apical area (Figure 5c–g) and flattening of the regression line in the treated embryos (Figure 5h–k). Our results demonstrate that Piezo1 is essential for calibrating centriole number in relation to the apical area in MCCs.

In Piezo1-depleted embryos, the centriole number in MCCs was similar to β-catenin depleted MCCs (median ± SD 100 ± 16) and untethered animal caps (median 105 ± 27), both of which experienced diminished pulling forces. Thus, our data suggest that Piezo1 is not essential to generate the first 100 centrioles but calibrates the final 50 centrioles in response to pulling forces. To test this hypothesis, we depleted Piezo1 and raised animal caps in untethered and tethered conditions. In the context of untethered animal caps, centriole number did not significantly differ in controls and Piezo1 depleted MCCs (Figure 6a,c, median ± SD, 101 ± 15 vs. 94 ± 15). In contrast, in tethered animal caps, Piezo one depletion led to a significant reduction in centriole number compared to controls (Figure 6b,d, median ± SD, 119 ± 14 in controls vs. 109 ± 16). Interestingly, with Piezo1 depletion, the number of centrioles in MCCs of tethered caps was similar to wild-type MCCs of untethered caps (median 109 in Piezo1 depleted compared to 101 in controls), demonstrating that Piezo1 is required for stretch-induced centriole amplification in the MCCs of Xenopus. Taken together, our data demonstrated that the stretching of MCCs due to morphogenetic movements calibrates centriole number in proportion to the apical area via Piezo1.

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MCCs must regulate the number of cilia to optimize extracellular fluid flow. While a previous study using mouse tracheal epithelial cell culture suggested a correlation between cilia number and the apical area (Nanjundappa et al., 2019), the underlying mechanism was unknown. Our study, using the Xenopus embryonic epidermis, demonstrates that MCC apical area undergoes dramatic size changes as cell non-autonomous forces generated by morphogenetic movements pull on the epithelia. Importantly, centriole amplification occurs while the MCCs are being stretched and expanding their apical surface. Therefore, the cells must contend with a complex mathematical problem: how to count centrioles, how to measure the apical area, and how to coordinate the two. Our results show that Piezo1 translates the pulling forces that define the apical area into an appropriate number of centrioles (Figure 6f). Thus, Piezo1-mediated mechanosensation couples apical area and centriole number (Figure 6e,f).

There are a few possibilities that may explain how Piezo1 calibrates centriole number and controls the correlation between centriole number and the apical area. One possibility is that stretch may activate Piezo1, which leads to an influx of Ca²⁺ from the extracellular environment. This increase in intracellular Ca²⁺ may promote centriole amplification via either transcriptional or non-transcriptional means. Alternatively, Piezo1 has been shown to regulate the expression of focal adhesion kinases (FAK, Paxillin and Vinculin) in cancer cells leading to changes in tissue stiffness (Chen et al., 2018). These three focal adhesion kinases localize to the bases of cilia in MCCs, and their downregulation causes defects in actin and cilia assembly (Antoniades et al., 2014). However, their role in the regulation of centriole number and apical area remains unexplored. Finally, filamentous (F)-actin plays a critical role in mechanotransduction in all cells (Massou et al., 2020; Wang, 2017). At the apical membrane, MCCs are enriched in F-actin and the apparent loss of apical F-actin in Piezo1-depleted cells may lead to defective mechanotransduction resulting in the defects in centriole amplification and apical area.

In our work, we exploited the frog multiciliated embryonic epithelium because of two main considerations. First, studying the effects of tissue scale forces on MCC apical area and cilia number requires an in vivo system and therefore is challenging in mammals. Second, we could exploit the two-step process of MCC formation (Deblandre et al., 1999; Stubbs et al., 2006), radial intercalation followed by apical expansion to determine the number of centrioles at time zero (just prior to apical expansion) (Step 1, Figure 1a, Figure 2). Then we could compare that number to the final count to understand the contribution of stretching forces on centriole amplification. While mammalian MCCs do not radially intercalate like Xenopus MCCs, they still scale centriole number to the apical area suggesting a conserved mechanism (Nanjundappa et al., 2019). Indeed, the formation of MCCs in the mammalian respiratory epithelium is similar to the goblet cells converted to MCCs by mcidas overexpression in Xenopus, which also scale centriole number to the apical area. Our results define a molecular pathway in which tissue scale forces regulate apical area and cilia number in MCCs. These results provide a new perspective to study the role of cell and tissue level mechanical forces in shaping organelle number to optimize cell function.

Data is attached as a source files.

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Author details

1. Saurabh Kulkarni

   Pediatric Genomics Discovery Program, Department of Pediatrics and Genetics, Yale University School of Medicine, New Haven, United States

   Present address

   Department of Cell Biology, Department of Biology, Center for Membrane and Cell Physiology, University of Virginia, Charlottesville, United States

   Contribution

   Conceptualization, Resources, Data curation, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing

   For correspondence

   sk4xq@virginia.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-0882-6478

2. Jonathan Marquez

   Pediatric Genomics Discovery Program, Department of Pediatrics and Genetics, Yale University School of Medicine, New Haven, United States

   Contribution

   Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-3377-7599

3. Priya Date

   Pediatric Genomics Discovery Program, Department of Pediatrics and Genetics, Yale University School of Medicine, New Haven, United States

   Present address

   College of Arts and Sciences, University of Virginia, Charlottesville, United States

   Contribution

   Investigation, Methodology

   Competing interests

   No competing interests declared

4. Rosa Ventrella

   Department of Cell and Developmental Biology, Feinberg School of Medicine, Northwestern University, Chicago, United States

   Contribution

   Investigation

   Competing interests

   No competing interests declared

5. Brian J Mitchell

   Department of Cell and Developmental Biology, Feinberg School of Medicine, Northwestern University, Chicago, United States

   Contribution

   Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

6. Mustafa K Khokha

   Pediatric Genomics Discovery Program, Department of Pediatrics and Genetics, Yale University School of Medicine, New Haven, United States

   Contribution

   Resources, Supervision, Funding acquisition, Investigation, Writing - original draft, Writing - review and editing

   For correspondence

   Mustafa.khokha@yale.edu

   Competing interests

   is a founder of Victory Genomics

   "This ORCID iD identifies the author of this article:" 0000-0002-9846-7076

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