mTORC1 signaling facilitates differential stem cell differentiation to shape the developing murine lung and is associated with mitochondrial capacity

Zhang, Kuan; Yao, Erica; Chuang, Ethan; Chen, Biao; Chuang, Evelyn Y.; Chuang, Pao-Tien

doi:10.1038/s41467-022-34763-y

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Published: 25 November 2022

mTORC1 signaling facilitates differential stem cell differentiation to shape the developing murine lung and is associated with mitochondrial capacity

Nature Communications volume 13, Article number: 7252 (2022) Cite this article

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Abstract

Formation of branched organs requires sequential differentiation of stem cells. In this work, we find that the conducting airways derived from SOX2⁺ progenitors in the murine lungs fail to form without mTOR complex 1 (mTORC1) signaling and are replaced by lung cysts. Proximal-distal patterning through transitioning of distal SOX9⁺ progenitors to proximal SOX2⁺ cells is disrupted. Mitochondria number and ATP production are reduced. Compromised mitochondrial capacity results in a similar defect as that in mTORC1-deficient lungs. This suggests that mTORC1 promotes differentiation of SOX9⁺ progenitors to form the conducting airways by modulating mitochondrial capacity. Surprisingly, in all mutants, saccules are produced from lung cysts at the proper developmental time despite defective branching. SOX9⁺ progenitors also differentiate into alveolar epithelial type I and type II cells within saccules. These findings highlight selective utilization of energy and regulatory programs during stem cell differentiation to produce distinct structures of the mammalian lungs.

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Introduction

Developmental processes are executed through successful implementation of cellular functions in distinct cell types and in a spatially and temporally specific manner. These cellular functions underlie key processes such as cell proliferation, migration, differentiation and morphogenesis. Essential cellular function relies on energy supply. Thus, it is assumed that all developmental processes are energy dependent. However, it is unclear if different cell types and cellular processes display a similar level of energy dependence in vivo. Answers to this question will provide insight into how energy is utilized at tissue and organismal levels during development.

Generation of the elaborate respiratory tree in mammals requires the production of an appropriate number of cells and acquisition of cellular properties that enable cell migration, differentiation and morphological changes^1,2,3,4. Extensive studies have uncovered genes and pathways and the cellular events that control many aspects of these processes^5,6,7,8. The lungs undergo multiple rounds of stereotypic branching⁹ to increase the total surface of gas exchange. Once lung branches are produced, SOX2⁺ cells demarcate the proximal airway while SOX9⁺ cells are distributed in the airways distal to the SOX2⁺ domain¹⁰. Expansion of the distal SOX9⁺ cell population is proposed to give rise to proximal SOX2⁺ cells through changes in the transcription program. In particular, cessation of Sox9 expression is largely concomitant with Sox2 activation and subsequent formation of the conducting airways. As a result, the conducting airways extend distally due to the sequential addition of newly generated SOX2⁺ cells, which differentiate into various cell types including club cells, ciliated cells and goblet cells. The distal SOX9⁺ progenitors subsequently generate saccules and differentiate into alveolar epithelial type I (AT1) and type II (AT2) cells. The alveolus^11,12,13, the unit of gas exchange, forms within saccules. Little is known about how energy utilization impacts distinct processes during lung development.

mTOR complex 1 (mTORC1) controls many essential cellular functions^14,15. It is indispensable for cell growth, cell proliferation and cell cycle progression¹⁶. mTORC1 is also a sensor of energy and nutrients and controls biomass accumulation and multiple aspects of cell metabolism including anabolic and catabolic processes¹⁷. mTORC1 modulates ATP production via regulating the capacity of the mitochondria^18,19. Rptor (Raptor, rapamycin-sensitive regulatory associated protein of mTOR)²⁰ encodes an essential component of mTORC1, which includes the RPTOR scaffold protein, the mTOR kinase and several other proteins²¹. How mTORC1 signaling regulates lung organogenesis has not been reported. Manipulating mTORC1 or mitochondrial function thus provides a means to modulate energy utilization during organ formation in animals.

In this work, we have employed lung development in mice as a system to decipher energy dependence in different cell types and cellular processes in vivo. Through control of mTORC1 signaling and mitochondrial capacity during lung development, we have discovered energy dependence for distinct cellular processes to form lung branches and saccules. The transition of the distal SOX9⁺ progenitors to proximal SOX2⁺ progenitors to produce the conducting airways is vulnerable to energy reduction. By contrast, morphogenesis of the distal SOX9⁺ progenitors to generate saccules and differentiation into alveolar epithelial cells within saccules are more resistant to energy deficiency. Through RNA-Seq we have identified the downstream effectors (including Hedgehog signaling) of the mTORC1–mitochondrial axis in forming the conducting airways. These findings suggest that a given cellular process in a distinct environment in vivo displays different energy requirements. This approach provides a way to reveal how energy utilization impacts cellular processes and classify cellular processes in terms of energy utilization.

Results

Inactivation of mTORC1 signaling in the lung epithelium disrupts branching morphogenesis

To study the effect of energy utilization on lung branching, we eliminated Raptor (Rptor), a key component of mTORC1, in the murine lung epithelium. We produced Rptor^f/f; Shh^Cre/+ mice, in which broad expression of Shh^Cre²² in the lung epithelium converted a floxed (f) allele of Rptor (Rptor^f)²⁰ to a null allele (Supplementary Fig. 1c). As a result, mTORC1 function was disrupted. Consequently, the cellular processes controlled by mTORC1 were perturbed. For instance, mTORC1-dependent phosphorylation of ribosomal protein S6 (RPS6)²³ was reduced in the lung epithelium of Rptor^f/f; Shh^Cre/+ mice at 12.5 days post coitus (dpc) (Fig. 1a, b).

**Fig. 1: *Rptor* controls mammalian lung branching.**

While development of the trachea and main stem bronchi was normal in Rptor^f/f; Shh^Cre/+ lungs, branching morphogenesis was compromised (Supplementary Fig. 1a, b, d). Defective branching was first observed at 12.5 dpc and continued throughout lung development (Fig. 1c, d). In control lungs at 11.5 dpc, the five main branches (right cranial [RCr], right middle [RMd], right caudal [RCd], right accessory [RAc] and left [L]) were established to generate the five lobes in adult lungs⁹. Daughter branches have emerged from the five main branches in control lungs at 12.5 dpc (Fig. 1c, d). By contrast, Rptor^f/f; Shh^Cre/+ lungs maintained the rudimentary pattern of five main branches without new daughter branches at 12.5 dpc (Fig. 1c, d). Multiple solid sphere-shaped structures formed at the distal part of Rptor^f/f; Shh^Cre/+ lungs (Fig. 1c, d) and these animals died soon after birth due to respiratory failure. These results suggest that mTORC1 plays an essential role in controlling lung branching. Analysis of the branching defects in Rptor^f/f; Shh^Cre/+ mice provides a unique opportunity to explore how energy utilization impacts specific developmental processes.

To investigate whether loss of mTORC1 altered cellular behaviors and thus defective branching, we probed the properties of epithelial cells. Cell proliferation assessed by EdU incorporation²⁴ was unaffected in Rptor^f/f; Shh^Cre/+ lungs at 12.5–13.5 dpc (Supplementary Fig. 2). mTORC1-deficient epithelial cells in either the proximal SOX2⁺ or distal SOX9⁺ domain had a similar rate of EdU incorporation in comparison with control lungs. This observation suggests that the branching defect in Rptor-deficient lungs was not due to reduced cell proliferation. We speculate that production of new lung epithelial cells in the absence of stereotypic branching leads to the formation of sphere-shaped structures and subsequently cysts in Rptor^f/f; Shh^Cre/+ lungs.

We noticed reduced expression of several markers at the apical surface of epithelial cells in Rptor^f/f; Shh^Cre/+ lungs at 13.5 dpc. For instance, the actin cytoskeleton (labeled by phalloidin)²⁵, pMLC2 (indicative of mechanical force production)²⁶ and PKCζ (apical epithelial cell marker)²⁷ were preferentially concentrated at the apical surface of epithelial cells in control lungs. Instead, phalloidin, pMLC2 and PKCζ levels were reduced at the apical surface of Rptor^f/f; Shh^Cre/+ lungs (Fig. 1e, g and Supplementary Fig. 3). These results suggest that removal of mTORC1 was associated with loss of apical-basal polarity and mechanical force production in epithelial cells. These defects could contribute to abnormal branching morphogenesis in Rptor^f/f; Shh^Cre/+ lungs.

The SOX9⁺ population in the distal airway fails to generate SOX2⁺ cells in the absence of mTORC1

Since lung branching is disrupted in Rptor^f/f; Shh^Cre/+ lungs, we investigated if specification of the proximal-distal airway was perturbed in the absence of mTORC1 signaling. This could be the molecular basis of defective branching. During lung branching, patterning of the airway smooth muscle is closely coordinated with patterning of the airways. Alpha smooth muscle actin (SMA)²⁸ is highly expressed in mesenchymal cells along the epithelial tube except at the tip of lung branches in control lungs. SMA expression in Rptor^f/f; Shh^Cre/+ lungs was largely absent along the stratified proximal epithelium and was only detected in the most proximal region (Fig. 1f, h). This suggests that proximal-distal patterning of airway smooth muscles is disrupted in Rptor-deficient lungs. Moreover, proximal-distal patterning of blood vessels²⁹, which follow airway branching, was also perturbed in Rptor-deficient lungs (Fig. 2a). These findings suggest that removal of epithelial Rptor disturbs proximal-distal patterning of all structures, including the airways, smooth muscles and blood vessels.

**Fig. 2: SOX2–SOX9 distribution is perturbed in *Rptor*-deficient lungs.**

To affirm this notion, we examined the distribution of SOX2⁺ and SOX9⁺ epithelial cells during lung development. The segregation of SOX2⁺ and SOX9⁺ cells was already apparent at 11.5 dpc in Rptor^f/f; Shh^Cre/+ lungs when no apparent structural abnormality could be discerned between control and Rptor^f/f; Shh^Cre/+ lungs (Fig. 2b, c, d and Supplementary Fig. 4a). At this stage, SOX2⁺ cells were detected in the five main branches derived from the two primary buds in control lungs. By contrast, SOX2⁺ cells were absent in the corresponding lung branches in Rptor^f/f; Shh^Cre/+ lungs. As development proceeded, the SOX2⁺ domain remained confined to the trachea and main stem bronchi of the mutant lungs (Fig. 2b, e, f and Supplementary Fig. 4b). This result suggests that the program that produces SOX2⁺ cells from SOX9⁺ progenitor cells is not turned on in the absence of mTORC1. As a result, the conducting airways that are derived from SOX2⁺ cells fail to form in Rptor-deficient lungs. It also implies that loss of mTORC1 has a fundamental impact on the acquisition of cellular properties in SOX9⁺ progenitor cells.

SOX9⁺ cells lacking mTORC1 still form saccules and differentiate into AT1 and AT2 cells

Branching morphogenesis is followed by the formation of saccules at the end of lung branches and specification of cell types in both the proximal and distal airways. In control lungs at 16.5 dpc, the distal SOX9⁺ progenitors produced saccules, which were composed of AT1 (AQP5⁺/T1α⁺/HOPX⁺) and AT2 (SPC⁺) cells. By contrast, cavitation of solid sphere-shaped structures in the distal regions of Rptor^f/f; Shh^Cre/+ lungs produced cysts, which became apparent at 16.5 dpc (Fig. 3a and Supplementary Fig. 5). Rptor^f/f; Shh^Cre/+ lungs at 18.5 dpc consisted of a few large cysts (Fig. 3b) without forming the conducting airways.

**Fig. 3: Loss of *Rptor* does not disrupt saccule formation.**

Unexpectedly, despite lack of branching morphogenesis, the cysts in Rptor^f/f; Shh^Cre/+ lungs contained saccule-like structures on the surface (Fig. 3c, d). At 15.5 dpc, structures with focal SOX9 expression could be observed on the surface of Rptor-deficient cysts (Supplementary Fig. 6a, b). These SOX9⁺ foci coalesced into irregular patches and developed into saccule-like structures at 16.5 dpc (Fig. 3c and Supplementary 6c, d, e). These saccule-like structures on the cystic surface of Rptor^f/f; Shh^Cre/+ lungs further branched and became interconnected, reminiscent of saccule formation in control lungs (Supplementary Fig. 7). To confirm that the saccule-like structure in Rptor^f/f; Shh^Cre/+ lungs was derived from the distal lung epithelium, we examined marker expression in these regions. Indeed, SOX2 was absent in saccule-like structures (Fig. 4a, b, e, h, i and Supplementary Fig. 6).

**Fig. 4: Cell type specification occurs in *Rptor*-deficient saccules despite loss of cell types in the conducting airways.**

We then asked whether production of AT1 and AT2 cells from the SOX9⁺ domain was affected in Rptor^f/f; Shh^Cre/+ lungs. We found that SOX9⁺ saccule-like structures were populated by AT2 and AT1 cells (Figs. 3c, d and 4c, g and Supplementary Fig. 8c, d). SPC⁺ AT2 cells were interspersed within a layer of AT1 cells and preferentially distributed at the tip of saccules in a pattern similar to that in control saccules. By contrast, cell types in the conducting airways such as club cells (CC10⁺) and ciliated cells (Ac-tub⁺) were absent in Rptor^f/f; Shh^Cre/+ lungs (Fig. 4d, f and Supplementary Fig. 8a, b). These findings suggest that while production of SOX2⁺ cells from the SOX9⁺ domain was perturbed, SOX9⁺ cells were capable of forming saccules and differentiating into AT1 and AT2 cells.

RNA-Seq analysis of mTORC1-deficient lungs reveals perturbed pathways involved in branching morphogenesis

To gain additional insight into how mTORC1 controls lung branching, we collected lungs from control and Rptor^f/f; Shh^Cre/+ mice at 11.5 dpc and performed RNA-Seq analysis³⁰ to probe the changes in transcriptomes in the absence of mTORC1 (Fig. 5a, b, c). Pathway analysis revealed perturbation of the tubular branching process and cell junction formation (Fig. 5b). Other pathways that control extracellular matrix organization, mesenchymal cell differentiation, cytosolic ribosome organization, apicolateral plasma membrane and basement membrane organization were also affected (Fig. 5c). Moreover, genes involved in the glycolytic pathway were also altered (Fig. 5a). This suggests that disturbed energy utilization in Rptor-deficient lungs is linked to cellular processes that control lung branching. RNA-Seq analysis at 11.5 dpc also revealed a sharp reduction in Sox2 levels while the expression levels of Sox9 were unaltered. This is consistent with a blockade in producing SOX2⁺ cells from SOX9⁺ progenitors in the absence of mTORC1.

**Fig. 5: Mitochondrial capacity is reduced in *Rptor*-deficient lungs.**

Mitochondrial capacity is reduced in mTORC1-deficient lungs

Our previous work uncovered a role of mTORC1 in controlling mitochondrial capacity during alveolar development³¹. This prompted us to investigate if ablation of mTORC1 signaling during lung branching also impacted mitochondrial capacity and thus branching morphogenesis. We found that the relative ratio of mitochondrial DNA (mtDNA)³² to nuclear DNA (nDNA) was reduced in Rptor^f/f; Shh^Cre/+ lungs compared to controls (Fig. 5d, e), suggesting perturbed mitochondrial function. Indeed, relative ATP production was reduced in Rptor^f/f; Shh^Cre/+ lungs compared to controls (Fig. 5f). We also determined mitochondrial mass in Rptor-deficient lung epithelium using MPC1 (mitochondrial pyruvate carrier 1)³³ or MTCO1 (mitochondrially encoded cytochrome c oxidase I)³⁴ as markers that label mitochondria in vivo. Mitochondrial mass was greatly decreased in Rptor^f/f; Shh^Cre/+ lungs (Fig. 5g, h, i, j and Supplementary Fig. 9). These findings support a model in which mTORC1 signaling controls lung branching and saccule formation in part through its influence on mitochondrial capacity.

Reduced mitochondrial function leads to branching defects but preserves saccule formation

Our observations suggest that a main target of mTORC1 signaling during lung branching and saccule formation is the mitochondria. We tested if perturbation of mitochondrial function also generated branching defects. To this end, we created Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ mice. Tfam encodes a major transcription factor for mitochondria while Cox10 encodes an essential component of the respiratory chain in mitochondria. Likewise, expression of Shh^Cre in the lung epithelium converted a floxed allele of Tfam (Tfam^f)³⁵ or Cox10 (Cox10^f)³⁶ into a null allele (Supplementary Fig. 10a). Loss of either Tfam or Cox10 is expected to disrupt mitochondrial function. A global reduction of MTCO1 in lung epithelium of Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ mice indicated efficient removal of Tfam and Cox10 by Shh^Cre (Fig. 6a, b, c, d). A reduction in MTCO1 signal in Tfam-deficient cells was due to reduced transcription of MTCO1. By contrast, MTCO1 protein becomes unstable in the absence of COX10, resulting in decreased MTCO1 immunoreactivity. Moreover, the relative mtDNA/nDNA ratio was decreased in Tfam^f/f; Shh^Cre/+ lungs (Fig. 6e) but not in Cox10^f/f; Shh^Cre/+ lungs (Fig. 6f). Relative ATP production was compromised in Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs (Fig. 6g). These findings indicate that loss of epithelial Tfam or Cox10 reduces mitochondrial capacity.

**Fig. 6: Decreased mitochondrial function due to loss of *Tfam* or *Cox10* disrupts lung branching.**

We found that lungs from Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ mice displayed defective branching morphogenesis (Fig. 6h and Supplementary Fig. 10b). The defects appeared slightly later than those in Rptor^f/f; Shh^Cre/+ lungs. Abnormal branching could be discerned at 13.5 dpc in Tfam^f/f; Shh^Cre/+ lungs and 12.75 dpc in Cox10^f/f; Shh^Cre/+ lungs (Fig. 6i). Nevertheless, the main characteristics of the branching phenotype were similar among the three mutants. The SOX2⁺ domain was also restricted to the trachea and main stem bronchi and some of the early branches in Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs (Fig. 6i). Cell proliferation in the lung epithelium was unaffected in the absence of Tfam or Cox10 (Supplementary Fig. 10c). Thus, defective branching from SOX9⁺ cells failed to produce the conducting airways and led to cyst formation in Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs.

We then evaluated saccule formation and differentiation of AT1 and AT2 cells in the SOX9⁺ domain of Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs. Saccule-like structures formed at the edge of the cysts in Tfam- and Cox10-deficient lungs at 16.5 dpc (Fig. 7a, c) and progressed to more elaborate structures at 18.5 dpc (Fig. 7b, d). Thus, reduced mitochondrial capacity disrupts branching morphogenesis to generate the conducting airways but preserves saccule formation.

**Fig. 7: Saccular formation progresses in the absence of *Tfam* or *Cox10*.**

The saccule-like structures in Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs expressed AT1 markers (e.g., T1α) and contained SOX9/SPC double positive cells at the tips (Figs. 7c, d and 8b, d, e and Supplementary Fig. 11) despite loss of MTCO1 (Fig. 8a, c). These results suggest compromised ATP production in the mutant lungs blocked the generation of SOX2⁺ cells from distal SOX9⁺ progenitors but did not prevent them from producing saccules and AT1/2 cells.

**Fig. 8: Differentiation of AT1/2 cells within the saccules occurs in the absence of *Tfam* or *Cox10*.**

As an alternative approach to perturb mitochondrial function, we inactivated the mitochondria-associated protein, LRPPRC (leucine-rich PPR-motif containing), in the lung epithelium using Shh^Cre to convert a floxed allele of Lrpprc, Lrpprc^f³⁷, into a null allele. LRPPRC regulates multiple aspects of mitochondrial function. Lrpprc^f/f; Shh^Cre/+ mice displayed branching defects (Supplementary Figs. 12 and 13), which could be first detected at 15.5 dpc and were milder than those in Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs. Lung cysts were observed mainly in the more distal part of Lrpprc-deficient lungs (Supplementary Figs. 12 and 13). Likewise, saccule-like structures that contained AT1 and AT2 cells developed along the edge of cysts in Lrpprc^f/f; Shh^Cre/+ lungs (Supplementary Figs. 12 and 13).

Taken together, our findings further support the notion that saccule formation and AT1/2 differentiation can occur independently of lung branching and highlight a differential energy requirement for distinct cellular processes (Fig. 8f).

The Hedgehog pathway acts downstream of the mTORC1-mitochondria axis in regulating formation of the conducting airways

To explore the downstream targets of the mTORC1-mitochondria axis during formation of the conducting airways we performed RNA-Seq analysis of control and Rptor^f/f; Shh^Cre/+, Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs at 14.5 dpc (Fig. 9a, b, c). In all three mutant lungs, Sox2 expression levels were greatly reduced while Sox9 expression levels were unchanged, suggesting a persistent failure of generating SOX2⁺ cells from SOX9⁺ progenitors. Moreover, expression levels of Foxj1 (ciliated cells) and Ascl1 (neuroendocrine cells) were reduced while expression levels of Trp63 (p63) (basal cells) in the trachea and main stem bronchi were unaltered. The changes in molecular signature further confirmed a selective loss of the conducting airways through phenotypic analysis when the mTORC1-mitochondria axis is disrupted.

**Fig. 9: Defective Hh signaling perturbs the development of the conducting airways without affecting saccule formation.**

We found that the Hedgehog (Hh) signaling pathway was downregulated in the lungs of all three mutants (Fig. 9a, b, c). Loss of Sonic hedgehog (Shh) in the lung³⁸ generated phenotypes (Fig. 9d, e) similar to those in Rptor^f/f; Shh^Cre/+, Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs. The conducting airways located between the main stem bronchi (p63⁺) and saccules (SPC⁺HOPX⁺) failed to form in Shh^−/− lungs (Fig. 9f, g). Moreover, the cystic surface in Shh mutant lungs was populated by SOX9⁺ patches (Supplementary Fig. 14). These results suggest that the mTORC1-mitochondria axis controls a signaling network that includes the Hh pathway.

Discussion

Our work illustrates how distinct cellular processes exhibit differential energy dependence. Using lung development as a model system, we have provided insight into how different stages of lung development are controlled. In particular, lung branching and saccule formation can be independently regulated. These findings serve as the paradigm for examining other cellular processes during tissue and organ formation.

Proximal-distal patterning of lung branches by expansion of SOX9⁺ progenitors and production of SOX2⁺ cells from SOX9⁺ progenitors has been well described¹⁰. However, the molecular basis of this process remains elusive. Through manipulation of mTORC1 signaling and mitochondrial capacity, we were able to dissect distinct steps of SOX9⁺ cell expansion, SOX9–SOX2 transition to form the conducting airways, and differentiation of SOX9⁺ cells to generate alveolar epithelial cells and form saccules (Fig. 8f). In the absence of Rptor, Tfam or Cox10 in the lung epithelium, SOX9–SOX2 transition is blocked and the conducting airways fail to form. However, SOX9⁺ cell expansion and saccule formation are unaffected (Fig. 8f). Our results suggest that formation of the conducting airways and saccules can be regulated independently.

Our analysis of Rptor, Tfam or Cox10 mutant lungs reveals that the distal SOX9⁺ progenitor pool is preserved during branching morphogenesis despite a failure to produce SOX2⁺-conducting airways (Figs. 8f and 10). During sacculation of the mutant lungs, HOPX⁺ AT1 cells are adjacent to the SOX2⁺ trachea and main stem bronchi. By contrast, SOX9⁺ AT2 cells are located primarily at the tips of saccules. This suggests that the distal SOX9⁺ progenitors in control lungs proliferate to maintain a pool of progenitors at the tip of the lung buds (tip progenitors). Tip progenitors differentiate to precursors of AT1 cells (transition progenitors), which are located proximally to the progenitor pool at the tip. Transition progenitors would then form SOX2⁺ progenitors while downregulating SOX9 expression to construct the conducting airways. In Rptor, Tfam or Cox10 mutant lungs, the tip progenitors can still expand and produce transition progenitors. However, derivation of SOX2⁺ cells from transition progenitors is blocked in the mutant lungs, resulting in loss of the conducting airways.

**Fig. 10: A model of lung branching and saccule formation.**

Formation of saccules on the cystic surface of mTORC1-deficient lungs or from the cystic edge of Tfam- or Cox10-deficient lungs facilitates visualization of saccule formation to further test our model. We were able to examine the rudimentary saccule structure at 15.5 dpc, the progression of saccule development at 16.5 dpc and the maturation of saccules at 18.5 dpc in the mutant lungs. Rudimentary saccule progenitors consist of SOX9⁺ cells (tip progenitors). During saccule development, SOX9⁺ cells are confined to the tips of saccules and express SPC. By contrast, SOX9^– cells within the saccules are associated with HOPX expression. When saccules mature, the SOX9 signal sharply decreases; the tips of saccules are occupied by SPC⁺ AT2 cells while the bulk of saccules is composed of HOPX⁺ AT1 cells. This is consistent with the notion that the tip progenitors produce SPC⁺ AT2 cells while the transition progenitors fail to generate SOX2⁺ cells and become HOPX⁺ AT1 cells in the mutant lungs. This sequence of saccule formation is consistent with the published data using lineage tracing and scRNA-Seq³⁹. Our work provides a framework for further studies to trace saccule formation dynamically by combining live imaging, organ culture and molecular tools.

The main characteristics of the lung phenotypes are similar among the Rptor, Tfam or Cox10 mutants in this study. Nevertheless, their defects are not identical. We surmise that this is due to the fact that mTORC1 controls multiple cellular processes in addition to mitochondrial capacity. We noticed that the majority of the newly generated epithelial cells in Rptor^f/f; Shh^Cre/+ lungs would fill the inside of the distal end of lung branches, forming solid sphere-like structures. The distal SOX9⁺ progenitors sit on the surface of these structures. By contrast, newly produced epithelial cells in Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs would push outward, resulting in cyst formation. The distal SOX9⁺ progenitors settle to the edge of the cysts. Thus, in Rptor^f/f; Shh^Cre/+ lungs, saccules sprout from the surface of the defective lobules while saccules in Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ lungs emerge from the edges of the cystic lobules. These results form the basis of future investigation to study the molecular basis of lung branching and saccule formation.

Emergence of saccules in the absence of lung branching in Rptor, Tfam, Cox10 or Shh mutant lungs suggests that these two processes can proceed independently (Fig. 10). While saccule formation normally commences once lung branching is completed, saccule formation can occur at the proper developmental time when lung branching is blocked. This infers that a program for saccule formation can be activated without the completion of lung branching and without an existing scaffold. It also indicates the presence of a developmental clock. In Sox9 mutant lungs, certain marker genes for differentiated alveolar cells were upregulated or prematurely expressed^40,41. This suggests that SOX9 suppresses premature initiation of alveolar differentiation. It is possible that SOX9 is involved in controlling the timing of alveolar differentiation. Our work has set the stage for identifying the regulatory programs that execute distinct steps of lung development and the developmental clock that controls lung development.

Transcriptome analysis and genetic studies suggest that Hh signaling functions as a downstream effector of the mTORC1-mitochondria axis. Loss of the conducting airways while preserving sacculation are common features in Rptor, Tfam, Cox10 and Shh mutants. Nevertheless, the size of the lung and cysts are smaller in Shh mutant lungs. We speculate that Hh signaling has additional functions in addition to mediating the effects of the mTORC1-mitochondria axis. Moreover, KEGG pathway analysis revealed several pathways perturbed in Rptor, Tfam and Cox10 mutant lungs (Supplementary Fig. 15), including the Rap1 signaling pathway, the PI3K-Akt signaling pathway and the Ras signaling pathway. The mitochondrial pathway was not among the top 40 enriched pathways (Supplementary Fig. 15). This is likely because Rptor controls mitochondrial genes posttranscriptionally. We noted that Kras activation in the lung epithelium was reported to disrupt SOX9 to SOX2 transition⁴⁰, a common defect shared by Rptor-, Tfam- and Cox10-deficient lungs. Taken together, we propose that mTORC1 signaling functions in a signaling network, part of which involves mitochondrial capacity, and Hh and Kras signaling. These observations provide molecular insight into how mTORC1 signaling, mitochondrial capacity and the downstream signaling events control branching morphogenesis. Nevertheless, the detailed molecular mechanisms by which mitochondrial capacity and ATP production regulate branching require additional investigation.

mTORC1 regulates multiple cellular events including mitochondrial function, metabolism, protein synthesis and autophagy. Several targets (e.g., eukaryotic translation initiation factor 4E-binding protein (4E-BP), ribosomal protein S6 kinase (S6K) and unc-51 like kinase (ULK)) of mTORC1 have been identified. It is interesting to note that a connection established in cell lines is not necessarily recapitulated in the animals. For instance, disruption of 4e-bp1, 4e-bp2, S6K1 and Ulk1/2 in mice does not lead to severe branching defects in the lungs as reported in the literature^42,43,44. Thus, genetic studies are required to identify the functional targets of mTORC1 in vivo. Our results in this study suggest that mTORC1 controls mitochondrial capacity during lung branching. mTORC1 is a repressor of mTORC2 signaling⁴⁵. This raises the possibility that the phenotypes in mTORC1 mutant lungs were caused by mTORC2 activation. However, it was reported that active mTOR signaling in the lung does not lead to branching defects⁴⁶. Thus, it is unlikely that mTORC2 activation underlies the defects in mTORC1-deficient lungs. In summary, this work has advanced our molecular understanding of lung branching and sacculation and provides the infrastructure for future studies.

Methods

Animal husbandry

The mouse experiments in this study were performed following the protocols (AN187712) approved by the Institutional Animal Care and Use Committee (IACUC) of the University of California, San Francisco (UCSF). Mice were housed in a SPF (specific pathogen free) facility with controlled temperature and humidity and a 12-hour dark/12-hour light cycle. Details on the mouse ages and genotypes are included in the figures and main text. The mouse lines used are listed below. Rptor^f [B6.Cg-Rptor^tm1.1Dmsa/J], Tfam^f [B6.Cg-Tfam^tm1.1Ncdl/J], Cox10^f [B6.129×1-Cox10^tm1Ctm/J], Lrpprc^f [Lrpprc^tm1.1Lrsn/J], ROSA26^mTmG [Gt(ROSA)26Sor^{tm4(ACTB-tdTomato,-EGFP)Luo}/J], ROSA26^tdTomato [B6;129S6-Gt(ROSA)26Sor^{tm14(CAG-tdTomato)Hze}/J] and Shh^Cre [B6.Cg-Shh^{tm1(EGFP/cre)Cjt}/J] were obtained from Jackson Laboratory (Bar Harbor, ME, USA).

Histology and immunofluorescence

Histology and immunofluorescence were performed as previously described^31,47. In brief, mouse lungs at the indicated time points were dissected and fixed with 4% paraformaldehyde (PFA) (MilliporeSigma, Cat# P6148) on ice for 1 h. The lungs were embedded either in OCT or paraffin wax and sectioned at 7 μm.

For immunofluorescence, the primary antibodies used were: chicken anti-GFP (1:200, abcam, Cat# ab13970; RRID:AB_300798), rabbit anti-NKX2.1 (1:100, Epitomics, Cat# 2044–1; RRID:AB_1267367), goat-anti-CC10 (1:200, Santa Cruz Biotechnology, Cat# sc-9773; RRID:AB_2183391), mouse anti-acetylated tubulin (1:200, MilliporeSigma, Cat# T6793; RRID:AB_477585), rabbit anti-prosurfactant protein C (proSP-C) (1:200, MilliporeSigma, Cat# AB3786; RRID:AB_91588), hamster anti-T1α (1:200, Developmental Studies Hybridoma Bank, Cat# 8.1.1; RRID:AB_531893), mouse anti-HOPX (1:100, Santa Cruz Biotechnology, Cat# sc-398703; RRID:AB_2687966), mouse anti-p63 (1:100, Santa Cruz Biotechnology, Cat# sc-8431; RRID:AB_628091), rabbit anti-MPC1 (1:100, MilliporeSigma, Cat# HPA045119; RRID:AB_10960421), rat anti-E cadherin (1:200, Life Technologies, Cat# 13-1900; RRID:AB_2533005), mouse anti-β-catenin (1:100, BD Transduction Laboratories, Cat# 610154; RRID:AB_397555), mouse anti-MTCO1 (1:100, abcam, Cat# ab14705; RRID:AB_2084810), mouse anti-GM130 (1:100, BD Biosciences, Cat# 610822; RRID:AB_398141), mouse anti-ACTA2 (1:200, Thermo Scientific Lab Vision, Cat# MS-113-P0; RRID:AB_64001), rat anti-PECAM-1 (CD31) (1:150, Santa Cruz Biotechnology, Cat# sc-18916; RRID:AB_627028), rabbit anti-PDGFRA (1:150, Cell Signaling Technology, Cat# 3164; RRID:AB_2162351), mouse anti-S6 Ribosomal Protein (1:100, Cell Signaling Technology, Cat# 2317; RRID:AB_2238583), rabbit anti-Phospho-S6 Ribosomal Protein (Ser235/236) (1:100, Cell Signaling Technology, Cat# 4856; RRID:AB_2181037), rabbit anti-SOX2 (D9B8N) (1:200, Cell Signaling Technology, Cat# 23064; RRID:AB_2714146), goat anti-SOX9 (1:200, R&D Systems, Cat# AF3075; RRID:AB_2194160), rabbit anti-pMLC (S19) (1:100, Cell Signaling Technology, Cat# 3671; RRID:AB_330248), rabbit anti-PKC Zeta (C-20) (1:100, Santa Cruz Biotechnology, Cat# sc-216; RRID:AB_2300359). Secondary antibodies and conjugates used were: Alexa Fluor® 488 donkey anti-chicken antibody (1:1000, Jackson ImmunoResearch Laboratories, Cat# 703-546-155; RRID:AB_2340376), Alexa Fluor® 488 donkey anti-goat (1:1000, Life Technologies, Cat# A11055; RRID:AB_2534102), Alexa Fluor® 488 donkey anti-mouse (1:1000, Life Technologies, Cat# A21202; RRID:AB_141607), Alexa Fluor® 594 donkey anti-mouse (1:1000, Life Technologies, Cat# A21203; RRID:AB_141633), Alexa Fluor® 647 donkey anti-mouse (1:1000, Life Technologies, Cat# A31571; RRID:AB_162542), Alexa Fluor® 488 donkey anti-rabbit (1:1000, Life Technologies, Cat# A21206; RRID:AB_2535792), Alexa Fluor® 594 donkey anti-rabbit (1:1000, Life Technologies, Cat# A21207; RRID:AB_141637), Alexa Fluor® 594 donkey anti-rat (1:1000, Life Technologies, Cat# A21209; RRID:AB_2535795), Biotinylated goat-anti hamster (1:1000, Vector Laboratories, Cat# BA-9100; RRID:AB_2336137), Biotin-SP-conjugated AffiniPure donkey anti-rabbit (1:1000, Jackson ImmunoResearch Laboratories, Cat# 711-065-152; RRID:AB_2340593), Biotin-SP-conjugated AffiniPure donkey anti-rat (1:1000, Jackson ImmunoResearch Laboratories, Cat# 712-065-150; RRID:AB_2340646), Biotinylated horse anti-mouse (1:1000, Vector Laboratories, Cat# BA-2000; RRID:AB_2313581), Streptavidin, Alexa Fluor® 488 conjugate antibody (1:1000, Life Technologies, Cat# S11223), Streptavidin, Alexa Fluor® 594 conjugate antibody (1:1000, Life Technologies, Cat# S11227), Streptavidin, Alexa Fluor® 647 conjugate antibody (1:1000, Jackson ImmunoResearch Laboratories, Cat# 016-600-084; RRID:AB_2341101). The signal was detected using streptavidin-conjugated Alexa Fluor 488, 594, or 647 (1:1000, Life Technologies) or HRP-conjugated streptavidin (1:1000, TSA kit; Perkin-Elmer, Cat# NEL753001KT) coupled with fluorogenic substrate Alexa Fluor 594 or 488 tyramide for 30 s (1:200, TSA kit; Perkin-Elmer, Cat# NEL753001KT). F-actin was stained with rhodamine-conjugated phalloidin (1:200; Thermo Fisher, Cat# R415) in PBS for 2 h. Confocal images were captured on a Leica SPE laser-scanning confocal microscope. Adjustment of red/green/blue/gray histograms and channel merges were performed using LAS AF Lite (Leica Microsystems).

Whole-mount immunostaining

Whole-mount immunostaining was performed as previously described^48,49. Briefly, the embryonic lungs at the indicated time points were collected and fixed in 4% PFA on ice for 1 h. Samples were washed with 0.1% Tween-20/PBS for 30 min then dehydrated in graded methanol solutions (25%, 50%, 75%, 100%). After incubating in 5% H₂O₂/methanol overnight, the lungs were rehydrated through graded methanol solutions (100%, 75%, 50%, 25%, 0%) which diluted in 0.1% Tween-20/PBS and incubated in blocking buffer (1.5% BSA/0.5% Triton X-100/PBS) for 6 h. Primary antibodies were incubated at 4 °C overnight. On the next day, lungs were washed with blocking buffer for 5 h, then incubated with secondary antibodies at 4 °C overnight. Images were captured on a Nikon Eclipse E1000 microscope with a SPOT 2.3 CCD camera. The primary antibodies used were: rat anti-E Cadherin (1:200, Life Technologies, Cat# 13-1900; RRID:AB_2533005), rabbit anti-SOX2 (D9B8N) (1:200, Cell Signaling Technology, Cat# 23064; RRID:AB_2714146), goat anti-SOX9 (1:200, R&D Systems, Cat# AF3075; RRID:AB_2194160), rabbit anti-prosurfactant protein C (proSP-C) (1:200, MilliporeSigma, Cat# AB3786; RRID:AB_91588), rat anti-PECAM-1 (CD31) (1:150, Santa Cruz Biotechnology, Cat# sc-18916; RRID:AB_627028).

Cell proliferation assays

Proliferation of lung epithelial cells was evaluated by EdU incorporation assays as previously described⁴⁸. Briefly, pregnant females at the indicated time points were intraperitoneally injected with 300 μl of 5 mg/ml EdU (Research Products International, Cat# B71800-1.0)/PBS solution 1 h before dissection. The Click-iT EdU Alexa Fluor 488 Imaging Kit (Thermo Fisher, Cat# C10337) was performed to assess EdU incorporation. To measure the rate of epithelial cell proliferation, EdU staining was co-stained with E-Cad, a pan-epithelial cell marker. Nuclear DAPI helped identify individual cells. The EdU signal is nuclear and co-localizes with the DAPI signal. The number of proliferating epithelial cells (EdU⁺ E-Cad⁺ DAPI⁺) and the total number of epithelial cells (E-Cad⁺ DAPI⁺) were manually counted using ImageJ. The rate of epithelial cell proliferation was calculated as the ratio of the number of EdU⁺ E-Cad⁺ DAPI⁺ cells to the number of E-Cad⁺ DAPI⁺ cells, which was displayed as (EdU⁺ E-Cad⁺)/(E-Cad⁺) for simplicity in the figures or legends. For each animal, at least 5 sections were used for counting.

Purification of lung epithelial cells

Lungs from Rptor^f/f; Shh^Cre/+, Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ mice and their littermate controls were dissected at the indicated time points, placed in the digestion solution (1.2 U/ml dispase (BD Biosciences, Cat# 354235), 0.5 mg/ml collagenase B (Roche, Cat# 11088815001) and 50 U/ml DNase I (QIAGEN, Cat# 79254)) and rocked at 37 °C for 1 h to release individual lungs cells. An equal volume of culture medium (DMEM (Mediatech, Cat# 10-013-CV) with 10% FBS (Gibco, Cat# 10437-028), 2x penicillin/streptomycin (Gibco, Cat# 15070-063) and 1x L-glutamine (Gibco, Cat# 25030-081)) was added to the digested lung samples, which were filtered through 40 µm cell strainers. After centrifugation at 600 × g for 10 min, the cell pellets were resuspended in 200 µl purification buffer (Phenol red-free DMEM with 0.2% BSA and 2% FBS). Biotin-conjugated EpCAM antibody (anti-CD326 (EpCAM), 1:100, eBioscience, Cat# 13-5791-82; RRID:AB_1659713) and Dynabeads M-280 Streptavidin (Invitrogen, Cat# 11205D) were added to the samples and incubated at 4 °C for 1 h with shaking. Epithelial cells were captured using Dynabeads magnets following the manufacturer’s instructions. Purified lung epithelial cells were used for qPCR, ATP measurement and Western blotting analysis.

Analysis of mtDNA/nDNA ratio

mtDNA/nDNA ratio was evaluated as previously described³¹. Briefly, purified lung epithelial cells from Rptor^f/f; Shh^Cre/+, Tfam^f/f; Shh^Cre/+, Cox10^f/f; Shh^Cre/+ mice and their littermate controls at 13.5 and 17.5 dpc were incubated in lysis buffer (100 mM Tris pH 7.5, 5 mM EDTA, 0.4% SDS, 200 mM NaCl, 50 μg /ml Proteinase K) at 55 °C overnight. 100 μg/ml RNase A was added and incubated at 37 °C for 30 min to degrade the RNAs. After mixing with an equal volume of phenol/chloroform, the samples were centrifuged at 13,400 × g for 10 min, and the lung DNA was concentrated by ethanol precipitation. Mitochondrial 16S rRNA or ND1 gene and the nuclear Hk2 gene were used to assess the relative ratio of mitochondrial (mt) to nuclear (n) DNA copy number. Quantitative PCR was performed on an Applied Biosystems QuantStudio^TM 5 Real-Time PCR System. The primer pairs used for the indicated genes were: 16S rRNA forward, 5′-CCGCAAGGGAAAGATGAAAGAC-3′; reverse, 5′-TCGTTTGGTTTCGGGGTTTC-3′, mt-ND1 forward, 5′-CTAGCAGAAACAAACCGGGC-3′; reverse, 5′-CCGGCTGCGTATTCTACGTT-3′, Hk2 forward, 5′-GCCAGCCTCTCCTGATTTTAGTGT-3′; reverse, 5′-GGGAACACAAAAGACCTCTTCTGG-3′.

Western blotting analysis

Purified lung epithelial cells were lysed in 2x Western blotting buffer (120 mM Tris-Cl pH 6.8, 4% SDS, 20% Glycerol, 200 mM DTT, 0.02% Bromophenol Blue). The lysates were centrifuged at 16,100 x g for 15 min at 4 °C before loading onto SDS-PAGE. The primary antibodies used were rabbit anti-Raptor (1:2000, EMD Millipore Corporation, Cat# 09-217; RRID:AB_1659713), mouse anti-S6 ribosomal protein (1:2000, Cell Signaling Technology, Cat# 2317; RRID:AB_2238583), rabbit anti-phospho-S6 ribosomal protein (Ser235/236) (1:2000, Cell Signaling Technology, Cat# 4856), rabbit anti-MLC2 (1:500; Cell Signaling Technology, Cat#3672; RRID:AB_10692513), goat anti-TFAM (1: 250, Santa Cruz Biotechnology, Cat# sc-23588; RRID:AB_2303230), rabbit anti-COX10 (1:500, Proteintech, Cat# 10611-2-AP; RRID:AB_2084833), mouse anti-MTCO1 (1:500, Abcam, Cat# ab14705; RRID:AB_2084810), and mouse anti-alpha-tubulin (1:3000, Developmental Studies Hybridoma Bank, Cat# 12G10; RRID:AB_1157911). Western blotting images were captured on a LI-COR Odyssey Fc device. The uncropped images are available in the Source Data file.

RNA-Seq analysis

Bulk RNA-Seq of mouse lungs was performed as previously described³¹. In brief, embryonic lungs from Rptor^f/f; Shh^Cre/+, Tfam^f/f; Shh^Cre/+ and Cox10^f/f; Shh^Cre/+ mice and their littermate controls (Rptor^f/+; Shh^Cre/+, Tfam^f/+; Shh^Cre/+ and Cox10^f/+; Shh^Cre/+ mice) were collected at 14.5 dpc. Lungs from Rptor^f/f; Shh^Cre/+ mice and controls were also collected at 11.5 dpc. For lungs collected at 11.5 dpc, three or four lungs of the same genotype were combined to reach a sufficient amount of RNA for RNA-Seq. The samples were lysed in 500 μl TRIzol (Ambion, Cat# 15596018). After adding 100 μl chloroform, the samples were centrifuged at 4 °C for 15 min. The upper aqueous phase was mixed with an equal volume of 70% ethanol. RNA was extracted with the RNeasy Mini Kit (Qiagen, Cat# 74104) following the manufacturer’s instructions. After quality evaluation with an Agilent 2100 Bioanalyzer, the RNAs were then sequenced on BGISEQ-500 sequencer. Differential gene expression, gene ontology (GO) enrichment analyses and the barplot of gene ontology enrichment were performed with RStudio (R version 3.4.0). Heatmap images were generated using the online Heatmapper software [http://www.heatmapper.ca/]. Datasets have been deposited into NCBI’s Gene Expression Omnibus database and are accessible through GEO Series accession numbers GSE189327 and GSE213202.

Statistics and reproducibility

All the quantification including the cell number, diameter, thickness, distance and area of the lungs and the signal density of immunostaining were performed using ImageJ (Version 1.49). Measurement of the relative diameter of the branches and the relative thickness of the mesenchyme relied on the GFP and tdTomato signals, respectively, produced in Shh^Cre/+; ROSA26^mTmG/⁺ and Rptor^f/f; Shh^Cre/+; ROSA26^mTmG/⁺ lungs. For each lung, at least 5 distal buds were used for assessing the diameter and thickness. For detecting the signal density of immunostaining, at least 15 views of images (5 views per biological repeat) were measured. The images were taken under 40X magnification using a Nikon Eclipse E1000 Microscope with the same exposure.

The mutant phenotypes described in this study were completely penetrant. All the mice of the same genotype showed similar, if not identical, lung phenotypes at each timepoint and there was little variation between the mutant lungs. The mutant phenotypes were apparent and could be discerned by the naked eye or under the dissecting scope. Accordingly, changes in morphology and molecular markers in the mutants can also be readily distinguished from the controls under the microscope. In this regard, blinding and randomization are not applicable. Nevertheless, sample tissues or sections were randomly selected for analysis such as mtDNA quantification, EdU counting, ATP production and RNA-Seq.

For all the micrograph-related assays, at least three biological repeats were performed. The completely penetrant and invariant phenotypes have allowed us to provide representative images shown in the figures. It has also allowed us to obtain statistical significance with the number of mice used in this study. Since an average of two mutants were obtained in each litter, we combined mutants from different litters for statistical analysis. Again, the completely penetrant and invariant phenotypes facilitated our statistical analysis.

For all the biological repeats, greater than or equal to three repeats were performed. The n number for biological replicates are shown in the figure legends. All the graphs were generated using GraphPad Prism 7 and the statistics are shown as mean value ± SEM. Two-tailed Student’s t-tests were performed to assess the P values of the comparisons between the control and mutant groups; the statistical significance was evaluated as *P < 0.05; **P < 0.01; ***P < 0.001.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

All the related data for this study are available in the published article and the Supplementary Information file. Additional data that support the findings of this study are available from the corresponding authors upon reasonable request. Raw and analyzed data of RNA-Seq have been deposited to the Gene Expression Omnibus (GEO) database under accession numbers GSE189327 and GSE213202. Source data are provided with this paper.

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Acknowledgements

Some data for this study were acquired at the Nikon Imaging Center at CVRI. This work was supported by grants (R01 HL142876) from the National Institutes of Health to P.-T.C.

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Cardiovascular Research Institute, University of California, San Francisco, CA, USA
Kuan Zhang, Erica Yao, Ethan Chuang, Biao Chen, Evelyn Y. Chuang & Pao-Tien Chuang

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Kuan Zhang
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Erica Yao
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Ethan Chuang
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Biao Chen
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Contributions

K.Z., E.Y. and P.-T.C. conceived and designed the study. K.Z., E.Y., E.C., B.C. and P.-T.C. performed the experiments. K.Z., E.Y., E.C., B.C., E.Y.C. and P.-T.C analyzed the data. K.Z., E.Y., and P.-T.C. wrote the manuscript. P.-T.C. supervised the research.

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Correspondence to Pao-Tien Chuang.

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Zhang, K., Yao, E., Chuang, E. et al. mTORC1 signaling facilitates differential stem cell differentiation to shape the developing murine lung and is associated with mitochondrial capacity. Nat Commun 13, 7252 (2022). https://doi.org/10.1038/s41467-022-34763-y

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Received: 02 January 2022
Accepted: 07 November 2022
Published: 25 November 2022
DOI: https://doi.org/10.1038/s41467-022-34763-y

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