The dynamics of tubulogenesis in development and disease

ABSTRACT

Tubes are crucial for the function of many organs in animals given their fundamental roles in transporting and exchanging substances to maintain homeostasis within an organism. Therefore, the development and maintenance of these tube-like structures within organs is a vital process. Tubes can form in diverse ways, and advances in our understanding of the molecular and cellular mechanisms underpinning these different modes of tubulogenesis have significant impacts in many biological contexts, including development and disease. This Review discusses recent progress in understanding developmental mechanisms underlying tube formation.

Summary: This Review provides a comprehensive overview of the mechanisms underlying the assembly of cells into functional tubes and how failures in these processes can lead to disease.

Introduction

Tubulogenesis is the process of creating biological tubes. Tubes consist of cells that encircle a central cavity called a lumen (an internal space within the tube), extend and, in specific tissues, branch out to form functional organs. Tubes have essential functions in facilitating the transport and exchange of nutrients, gases and waste, and selective transport is crucial for regulating the movement of substances into and out of the body, as well as between tissues (Lubarsky and Krasnow, 2003). The terms ‘tube’ and ‘tubule’ both refer to hollow, cylindrical structures, but their usage depends on the context. ‘Tube’ is a general term used in broader contexts, whereas ‘tubule’ usually refers to a small tube. However, the naming of tubes and tubules varies among organs and organisms, making it challenging to definitively establish a scale for distinguishing tubes from tubules across different tissues. Therefore, in this Review, we use ‘tube’ and ‘tubule’ interchangeably, adapting these terms to the specific tissue context as required.

Understanding the mechanisms that drive tubulogenesis is crucial for better comprehending the physiological, mechanical and structural functions of organ systems. Indeed, tubulogenesis is a key aspect of organogenesis (Shaye and Soto, 2021). Numerous studies have examined tubulogenesis using 3D cell culture and vertebrate tissues such as the kidney, lung, pancreas, vasculature and gut (Martin-Belmonte et al., 2008; Bryant et al., 2010; Hiremath et al., 2023; Metzger et al., 2008; Kesavan et al., 2009; Herwig et al., 2011; Alvers et al., 2014). Additionally, invertebrate systems, such as the excretory system in Caenorhabditis elegans and tubular structures in Drosophila, can serve as tractable in vivo models for understanding how cells organize into tubular shapes (Armenti et al., 2014; Roper, 2012). Emergent animal systems, such as the hydro-vascular organ in sea stars and the notochord in ascidians, have also facilitated our understanding of the evolutionary origins of tube formation (Denker and Jiang, 2012; Perillo et al., 2023).

The coordination of cellular processes, such as cell–cell and cell–matrix interactions, cell polarization, cytoskeletal regulation, trafficking events and mechanical forces, allows the organized assembly of cells into tube structures (Loganathan et al., 2020; Levic and Bagnat, 2023; Camelo and Luschnig, 2021). Each of these mechanisms plays a significant role in lumen formation, tube elongation and tube maintenance. The formation of a single enclosed lumen is a common and conserved process described in metazoans (Lubarsky and Krasnow, 2003; Andrew and Ewald, 2010). However, the exact mechanisms underlying tubulogenesis vary across tissues and species. For example, some tubes have lumens enclosed by a single cell, while others consist of multiple cells surrounding a central lumen (Marciano, 2017). In this Review, we discuss common modes of tubulogenesis across various organisms and tissues, including the respiratory, circulatory, urogenital and nervous systems of invertebrate and vertebrate models (Table 1). Previous articles have reviewed the molecular mechanisms that are common in lumen formation (Camelo and Luschnig, 2021; Levic and Bagnat, 2023). In this Review, we extend this knowledge and provide a comprehensive overview of the processes and cellular mechanisms that allow cells to assemble into tubes, the mechanisms underlying tubule network expansion, and how failures in these processes can result in disease (Table 2, Box 1). We also provide a perspective on the current understanding of how mechanical forces influence the development of biological tubes.

Table 1.

Models of tubulogenesis in vivo

Mode of lumen formation	Mechanism	Organism	Tissues	References
From epithelial layers	Wrapping	Mouse and Xenopus	Neural tube	Kowalczyk et al., 2021; Ossipova et al., 2014; McShane et al., 2015; Lee et al., 2007
	Budding	Drosophila	Tracheal system	Kondo and Hayashi, 2013; Hayashi and Kondo, 2018
			Salivary glands	Roper, 2012; Sanchez-Corrales et al., 2018, 2021; Sidor et al., 2020
		Mouse	Lungs	Metzger et al., 2008
		Mouse	Ureteric bud	Chi et al., 2009; Riccio et al., 2016
De novo	Entrapment	Drosophila	Heart	Santiago-Martínez et al., 2008; Medioni et al., 2008
	Cavitation	Mouse	Mammary glands	McCaffrey and Macara, 2009; Elias et al., 2015; Mailleux et al., 2007
	Cord hollowing	Mouse	Aorta	Strilic et al., 2009, 2010
		Chick	Second neurulation	Gonzalez-Gobartt et al., 2021; Dady et al., 2014
		Zebrafish	Vasculature	Herwig et al., 2011; Wang et al., 2010
			Intestine	Bagnat et al., 2007; Horne-Badovinac et al., 2001; Alvers et al., 2014; Rodriguez-Fraticelli et al., 2015
		Mouse	Pro-amniotic cavity	Bedzhov and Zernicka-Goetz, 2014; Christodoulou et al., 2018
		Mouse	Pancreas	Kesavan et al., 2009; Villasenor et al., 2010; Azizoglu et al., 2017
			Renal vesicles	Yang et al., 2013; Hiremath et al., 2023; Karner et al., 2009
	Cell hollowing	Zebrafish	Vasculature	Kamei et al., 2006
		C. elegans	Excretory system	Armenti et al., 2014; Khan et al., 2013; Kolotuev et al., 2013; Berry et al., 2003
		Drosophila	Dorsal fusion branch	Kakihara et al., 2008; Gervais and Casanova, 2010; Gervais et al., 2012
From an existing tube	Tube splitting	Mouse and Xenopus	Trachea and esophagus	Nasr et al., 2019; Que, 2015

Table 2.

Birth defects associated with tubulogenesis

Disease/ condition	Mode of tubulogenesis	Gene mutations	Species	Phenotype	Mechanism and signaling pathways	References
Neural tube defects	Wrapping	MTHFR	Human	NTD	Folate metabolism	Daly et al., 1995
		Vangl2	Mouse	Craniorachischisis	PCP signaling	Kibar et al., 2001
		Pax3	Mouse	NTD	Canonical Wnt signaling	Palmer et al., 2021
		PAX3	Human	NTD	Canonical Wnt signaling	Agopian et al., 2013
		Grhl3	Mouse	NTD	Canonical Wnt signaling	Gustavsson et al., 2007
		Ptc1 (Ptch1)	Mouse	NTD	SHH signaling	Zhang et al., 2001
		Bmp4	Mouse	NTD	BMP signaling	Felder et al., 2002
		Nog
		Raldh2 (Aldh1a2)	Mouse	NTD	Retinoid signaling	Niederreither et al., 1999
		Hes1	Mouse	Exencephaly	Notch signaling	Ishibashi et al., 1995
Tracheoes-ophageal fistula	Budding; splitting	Foxf1	Mouse	EA, TF, and lung anomalies	Disruption of endothelial AFs	Mahlapuu et al., 2001
		Sox2	Mouse	EA and TF	Dorsal ventral patterning, epithelium organization	Que et al., 2007
		Shh	Mouse	EA and TF	SHH signaling	Litingtung et al., 1998
		Gli2−/−	Mouse	Absent esophagus, trachea, and lungs	SHH signaling	Motoyama et al., 1998
		Gli3+/−
Congenital pulmonary airway malformation	Budding	Spry2	Mouse	Abnormal bronchial branch formation	FGF10 signaling*	Mailleux et al., 2001; Scott et al., 2010
		Bmp4
Congenital heart defects	Entrapment; cord hollowing	NKX2-5	Human	ASD	BMP and Notch signaling	Schott et al., 1998
		GATA4	Human	VSD and BAV	SHH signaling	Li et al., 2018
		TBX1	Human	CTD	Cell proliferation	Gao et al., 2015
Polycystic kidney disease	Tubule expansion	Pkd1	Mouse	Kidney cysts	Lengthening of cilia	Kurbegovic et al., 2010
		Pkd2	Mouse	Kidney cysts	Loss and shortening of cilia	Kim et al., 2009
		Phdhd1	Mouse	Kidney cysts and liver fibrosis	Increased branching and shortening of cilia	Woollard et al., 2007

ASD, atrial septal defect; AFs, adherens junctions; BAV, bicuspid aortic valve; BMP, bone morphogenetic protein; CTD, conotruncal defect; EA, esophageal atresia; FGF10, fibroblast growth factor 10; NTD, neural tube defects; PCP, planar cell polarity; SHH, sonic hedgehog; TF, tracheoesophageal fistula; VSD, ventricular septal defect.

Box 1. Disrupted tubulogenesis in birth defects.

Abnormal development of organs originating from tube structures can result in various congenital malformations. Some examples are neural tube defects, tracheoesophageal fistula, congenital heart defects and polycystic kidney disease (Table 2).

The brain and spinal cord are formed by a two-step neurulation process. In mice, primary neurulation is achieved by the creation of neural folds by bending and fusing the neural plate. The spinal cord is then formed during secondary neurulation by the hollowing of neuroectodermal cells (McShane et al., 2015). Failure of primary neurulation results in open neural tube defects, such as spina bifida.

Tracheoesophageal fistula is a congenital anomaly resulting from the abnormal separation of the trachea and esophagus, leading to aberrant connections between these two major organs. In several mouse mutant models, defective separation of primitive foregut by abnormal outgrowth or septation formation between respiratory and gastrointestinal tracts results in similar phenotypes to human with incomplete separation of lung and esophagus (van Lennep et al., 2019).

Congenital pulmonary airway malformation (CPAM) arises from abnormal branching morphogenesis in lung development. This condition is characterized by cyst-like structures in the lung tissues, which result from improper budding and excessive growth and proliferation of lung branches. Although the precise cause of CPAM remains unclear, several molecular factors have been identified that may inhibit normal budding, as detailed in Table 2 (Caldeira et al., 2021).

Gross heart structure is formed by correct folding, looping, septation, and differentiation of cells of the primitive heart tube. Disruption of each process can result in various types of congenital heart disease. In addition, failure of complete septation formation between chambers results in abnormal blood flow and heart failure (Goenezen et al., 2012).

Polycystic kidney disease occurs due to the abnormal formation of primary cilia in the kidney tubules. Major mutations in Pkd1, Pkd2, or Pkhd1 disrupt the differentiation of kidney epithelium in mouse by causing abnormal signaling cascades (Olson et al., 2019), which lead to increased cell proliferation and apoptosis. This results in the formation of cysts and a decline in kidney function.

Modes of tubulogenesis

Tube formation

The fundamental structure of a tube consists of a highly polarized epithelium (or endothelium, in vascular tissues) surrounding a central luminal space (Andrew and Ewald, 2010). The apical surface of the cells lines the lumen and is in contact with the transport medium, while the basal surface interacts with other tissues or a basement membrane, and the lateral surfaces connect adjacent cells through specialized junctions (Andrew and Ewald, 2010; Lubarsky and Krasnow, 2003). This cellular organization can be achieved by remodeling an existing sheet of epithelial cells (in the case of wrapping and budding) or by the polarization of precursor cells that then create a luminal space de novo (in the case of entrapment, cavitation and hollowing) (Table 1).

During wrapping, an epithelial sheet folds to form a tube (Fig. 1A). This sheet typically folds inwards along ‘hinges’ until the opposite edges meet (Lubarsky and Krasnow, 2003; Iruela-Arispe and Davis, 2009). These edges come into contact with each other along the midline, and the new hollow tube is eventually separated from the surrounding non-tube epithelia (Andrew and Ewald, 2010). The most prominent example of wrapping is neural tube development. Neural tube closure occurs in chordates and involves bending the neural plate along the midline so that neural folds from the opposite sides are brought together at the dorsal midline to fuse, creating a hollow tube (Moon and Xiong, 2022). Failure of this process causes neural tube defects (Table 2, Box 1), affecting approximately one in every 2758 newborns in the USA (Mai et al., 2019).

Fig. 1.

Modes of tubulogenesis. (A,B) Tubes can be formed from existing sheets of epithelial cells. During wrapping (A), an epithelial sheet folds to form a tube along ‘hinges’ until the opposite edges meet. This results in a new, hollow tube being separated from the surrounding non-tube epithelia. During budding (B), cells initiate tube formation as an outgrowth from an epithelial sheet. As the new tube elongates, further lateral budding can lead to the formation of a new, single tube, while bifurcation events create two identical tubes. (C-E) Tubes can also be formed de novo, from unpolarized cells. During entrapment (C), cells migrate (blue arrows) to the future location of the lumen and organize into two parallel rows that meet at the dorsal midline. The cells then establish polarity along the dorsal-ventral axis. As the cells meet at the midline, they form a tube by bringing together their dorsal regions. Finally, their ventral regions come together to enclose a central lumen. During cavitation (D), apoptosis in the central cells of a cell mass facilitates the formation of a luminal space. Tubes can also be formed via two different modes of hollowing (E). During cord hollowing, a lumen is created between two or more cells that were initially tightly connected. During cell hollowing, a single cell forms a lumen within itself (autocellular tubes). (F) During splitting tubulogenesis, the tube wall extends into the lumen, creating a transient septum that is progressively resolved by epithelial remodeling, localized extracellular matrix degradation, and mesenchymal cell invasion around the lateral constriction points. This results in the original tube being split into two tubes.

During budding, cells initiate de novo tube formation as an outgrowth of an existing epithelial sheet (Fig. 1B). This process occurs during the development of the tracheal system and salivary gland in fruit fly embryos, as well as in the formation of the mammalian kidney and lung (Riccio et al., 2016; Metzger et al., 2008; Sanchez-Corrales et al., 2018).

By contrast, entrapment, cavitation and hollowing create a luminal space de novo using unpolarized precursor cells (Andrew and Ewald, 2010). Entrapment occurs when migrating cells converge to form a lumen. A notable example is observed during Drosophila heart morphogenesis (Fig. 1C). In this process, myoendothelial cardioblasts align into two parallel rows that converge at the dorsal midline (Medioni et al., 2008; Santiago-Martínez et al., 2008). The cells establish polarity along the dorsal-ventral axis as they meet at the midline (Medioni et al., 2008). Lumen formation begins when the dorsal regions of the cardioblasts come together, followed by their ventral regions, forming a tube (Fig. 1C) (Medioni et al., 2008). Cavitation involves the death of cells in the central region of a multicellular mass, with these cells being eliminated through apoptotic mechanisms when they no longer contact the surrounding cells (Debnath and Brugge, 2005) (Fig. 1D). The external cells polarize and create a lumen that matures and expands over time (Debnath and Brugge, 2005). Cavitation occurs during the development of the mammary gland, where the gland forms a tree-like structure consisting of ducts with hollowed lumens (Pfannenstein and Macara, 2023). The mammary ducts branch out, while apoptosis creates the lumens by clearing body cells through the proapoptotic factor BIM (also known as BCL2L11) (Mailleux et al., 2007). Reducing the levels of Bim in the MCF10A breast epithelial cell line or in developing mouse mammary glands results in decreased apoptosis and impaired lumen formation (Schmelzle et al., 2007; Mailleux et al., 2007). Finally, hollowing tubulogenesis involves the appearance of a luminal space between two or more tightly connected, neighboring cells, or within a single cell; these processes are called cord hollowing and cell hollowing, respectively (Fig. 1E). Cell hollowing occurs during excretory organ development in C. elegans (Kolotuev et al., 2013; Khan et al., 2013) and in the specialized fusion and tracheal cells that facilitate maturation of the primary branches in the Drosophila tracheal system (Gervais et al., 2012). Cell and cord hollowing events both seem to occur during the formation of the zebrafish vasculature (Herwig et al., 2011; Kamei et al., 2006).

Tube network expansion

Expanding the tube network to increase the number of tubes can improve organ function. For example, the lungs are specialized for efficient oxygen exchange due to their large surface area, and the kidneys utilize thousands of nephrons to achieve the surface area required for efficient blood filtration. Many organs have, therefore, evolved a means to branch their tubes, meaning that new tubules can also form by altering existing tubes.

Budding

In addition to forming new tubules from an epithelial sheet, budding tubulogenesis can occur when a new tubule buds away from an existing parental tubule (Fig. 1B). Lateral (side) budding leads to the formation of a single tube, while bifurcation events create two identical tubes (Fig. 1B).

The mammalian lungs employ both lateral budding and bifurcation to form their branched architecture (Fig. 2) (Metzger et al., 2008), and aberrations in pulmonary branching morphogenesis are thought to be the underlying cause of congenital pulmonary airway malformation (CPAM), the most common congenital malformation detected in the neonatal lungs (Table 2) (Swarr et al., 2018). In mice, the lung forms a tree-like structure via three budding modes, which have been classified as domain branching, planar bifurcation and orthogonal bifurcation (Metzger et al., 2008). In domain branching, lateral daughter branches regularly form along a single circumferential position through lateral budding. The tip of each row expands and bifurcates to form a pair of tertiary branches, which then bifurcate again in a similar orientation to form four quaternary branches. When these bifurcation events occur in the same plane, it is called planar bifurcation. If the end of a branch splits into two branches, but the division is made in a plane rotated 90° with respect to the plane of the previous branching, this event is called orthogonal bifurcation (Metzger et al., 2008). These reproducible branching events also occur in the mammalian kidney (Fig. 2) (Yu et al., 2019).

Fig. 2.

Tubule branching geometries in different mammalian organs. Schematics showing the branching geometries of the mouse kidney, lung and mammary gland during embryogenesis. The images depict initiation of organ formation (top), formation of the lateral branches (middle) and organ maturation (bottom). These organs undergo a mixture of tube elongation (yellow) and tube budding (brown) to build different organs. In the kidney, the ureteric bud undergoes branching to form a tree-like structure, which eventually becomes the collecting ducts (Costantini and Shakya, 2006). This process is regulated by signaling between the ureteric bud and the surrounding mesenchyme (Costantini and Shakya, 2006). In the lung, growth factors and genetic signals guide repeated budding to create the airway tree (Metzger et al., 2008). FGF10 and SHH signaling play a key role in promoting branching morphogenesis in the mouse lung (Goodwin and Nelson, 2020). In the mammary gland, duct formation is regulated by hormones including estrogen and growth hormone (Gjorevski and Nelson, 2011). For example, growth hormone stimulates stromal cells in the mammary fat pad to release insulin-like growth factor 1 (IGF1), which interacts with the IGF1 receptor (IGF1R) on epithelial cells to promote the growth and branching of the mammary ducts (Gjorevski and Nelson, 2011).

By contrast, during the expansion of the tube network in the mouse mammary gland, tube elongation is more prominent than tube bifurcation (Fig. 2) (Huebner and Ewald, 2014). During puberty (3 weeks of age in mice), tube-like structures called ducts form, and their elongation is driven by groups of stem cells at their tips, known as cap cells, that are present in the terminal end buds (Sreekumar et al., 2017).

Splitting

The tube network can also be expanded via tube splitting, whereby an existing tube is divided into two new tubes (Fig. 1F) (Que, 2015; Nasr et al., 2019). During angiogenesis, tubule splitting begins with the formation of a tissue pillar within a blood vessel (Mentzer and Konerding, 2014). This pillar then expands and remodels, effectively dividing the vessel into two distinct channels, each maturing into a separate, fully functional blood vessel. By contrast, the separation of the trachea and esophagus in Xenopus and mouse embryos is thought to occur through the action of tightly packed cells (mesenchymal condensations) positioned on the sides of the tube where the boundary between the esophagus and trachea will form (Fig. 1F) (Nasr et al., 2019). These cells press against the tissue at the constriction points, forming a transient septum that is progressively resolved by epithelial remodeling, localized extracellular matrix (ECM) degradation, and mesenchymal cell invasion, leading to the formation of two distinct tubes: the esophagus and the trachea (Nasr et al., 2019). Splitting tubulogenesis has wide-ranging implications for human health, as detailed in Table 2 and Box 1. For example, in the context of angiogenesis, splitting appears to function as a compensatory mechanism for remodeling blood vessels in diseased lungs (Mentzer et al., 2022), while disruption of tracheoesophageal separation is suspected to be an underlying cause of tracheoesophageal birth defects (Table 2), occurring in approximately one in every 3500 newborns (Shaw-Smith, 2006).

Molecular mechanisms underpinning tubulogenesis

As discussed above, tubes can form either from epithelial sheets that already exhibit apicobasal polarity, or from cells that polarize de novo. In the latter case, unpolarized precursors can undergo a mesenchymal-to-epithelial transition, during which mesenchymal cells gradually acquire apicobasal polarity (Pei et al., 2019; Gredler and Zallen, 2023; Jackson et al., 2017). Therefore, organization of the proteins that establish and maintain epithelial cell polarity is crucial for tube formation.

Cell polarity refers to the asymmetric organization of cell structures, including the cell surface, organelles and cytoskeleton (Bryant and Mostov, 2008). In various epithelial tissues, specific polarity protein complexes establish the apical and basolateral domains, defining their identity and boundaries (Buckley and St Johnston, 2022). The Par (Par6, aPKC and Cdc42) and CRUMBS (Crumbs, Pals1 and Patj) complexes specify the apical domain, whereas the Scribble complex (Scrib, Lgl and Dgl) defines the basolateral domain (Assemat et al., 2008; Jewett and Prekeris, 2018; Blasky et al., 2015). These complexes comprise scaffolding proteins that attach transmembrane proteins to specific membrane domains and control cell shape and function by regulating the actomyosin cytoskeleton, which is responsible for cell movement and contraction (Campanale et al., 2017). The polarity complexes also regulate the secretion of specific cargoes (growth factors, ion channels, receptors and ECM proteins) to the different membrane domains and control the distribution and stability of adherens junctions (AJs), which hold cells together laterally, as well as tight junctions (TJs), which act as a barrier to paracellular diffusion (Campanale et al., 2017; Mira-Osuna and Le Borgne, 2024).

Many proteins that regulate cell polarity also control junctional components in tubular structures (Flasse et al., 2020; Krneta-Stankic et al., 2021). During tubulogenesis, cells need be flexible to reorganize their positions and change their shapes to form a tube (Pfannenstein and Macara, 2023; Symonds et al., 2020; Bruser and Bogdan, 2017). At the same time, they must remain cohesive to maintain tissue structure (Guo et al., 2018). The balance between cellular plasticity and cohesion is achieved through the regulation of cell–cell adhesion (Bruser and Bogdan, 2017). For example, endothelial cells bud out from existing blood vessels to produce a hierarchically branched vascular network (Chappell et al., 2011). This process involves dynamic cell intercalations and coordinated changes in cell shape, which require cell–cell contacts to be continuously remodeled (Bentley et al., 2014). In mouse, gain- and loss-of-function studies have demonstrated that the endocytosis of VE-cadherin (cadherin 5)-mediated AJs plays a crucial role in both cell migration and tissue integrity during blood vessel formation (Grimsley-Myers et al., 2020). Additionally, studies in the mouse mammary gland suggest that both AJs and TJs are essential in driving cell intercalation in the terminal end buds, which in turn is required for duct elongation (Pfannenstein and Macara, 2023; Kurley et al., 2012; Daniel et al., 1995; Shamir et al., 2014).

Cell polarity is initiated by interactions with the ECM, which are primarily mediated by integrins (Lee and Streuli, 2014). In Madin–Darby canine kidney (MDCK) 3D cysts (spherical structures with a single central lumen), β1-integrins sense collagen in the ECM and initiate polarity (Yu et al., 2005). When laminin, the β1 integrin receptor, or downstream signaling of the Rac1 GTPase is lost, the polarity of these epithelial cells can be reversed either partially or completely, meaning that the ECM-adjacent membranes exhibit reduced levels of basolateral proteins and are enriched in apical markers (Yu et al., 2005). Depleting β₁ integrin in the nascent mouse endothelium perturbs both arterial endothelial cell polarity and lumen formation (Zovein et al., 2010). Integrins also play an important role during tubulogenesis in invertebrate systems. For example, in the Drosophila tracheal system, key laminin receptors, such as dystroglycan (Dg) and the αPS3 integrin (Scb), control cell shape and modulate the cortical cytoskeleton during tube elongation (Klussmann-Fricke et al., 2022).

The establishment and maintenance of apicobasal polarity plays an important role in all the modes of tubulogenesis discussed in this Review. In the following sections, we focus on examples of tubulogenesis that occur via wrapping, budding and hollowing, as well as tube network expansion, to explore mechanisms that are specific to these different modes.

Mechanisms involved in wrapping tubulogenesis

During neurulation, the neural plate develops into the neural tube, the precursor of the central nervous system. In amphibians, this process forms the entire neural tube, while in birds and mammals, it forms the anterior portion (Andrew and Ewald, 2010). Primary neurulation employs the wrapping mode of tubulogenesis, where the flat neural plate folds into a closed tube. This occurs through a fusion point that moves directionally, progressively joining the two edges of the epithelial sheet in a process called ‘epithelial zippering’ (Mole et al., 2020).

The exact sequence of primary neurulation events may vary across different species, but, in most vertebrates, it involves convergent extension, whereby the tissue becomes narrower along the mediolateral axis while extending along the anterior-posterior axis (Nikolopoulou et al., 2017). This process shapes the neural plate before its closure. Meanwhile, processes such as apical constriction (a reduction in the apical surface area of a cell) and localized cell proliferation in both neural and non-neural ectoderm produce hinge points in the neural plate, causing the neural plate to fold into a tube (McShane et al., 2015). The processes of convergent extension, apical constriction and cell proliferation all rely on signals from the ECM, detected by receptors on the cell surface (Davidson et al., 2004; Huebner and Wallingford, 2018; Au et al., 2021). For instance, activation of mechanosensors on primary cilia causes the cytoskeleton to reorganize, guiding the migration of neural and non-neural ectodermal cells in specific directions (Au et al., 2021).

Analysis of mouse mutants has revealed that more than 300 genes are involved in the regulation of neural tube closure, with disruptions in any of these genes potentially leading to neural tube defects (Wilde et al., 2014). These include genes that encode structural and functional components of cilia and ECM, as well as genes involved in the WNT signaling pathway, actomyosin cytoskeleton remodeling, cell migration, the non-canonical Wnt/PCP pathway, Shh/BMP signaling, and the transcription factors Pax3, Cdx2, Zic2 and Grhl3 (Nikolopoulou et al., 2017; Wilde et al., 2014). The non-canonical Wnt/PCP signaling pathway regulates cytoskeleton dynamics and drives neural tube closure and body axis elongation by controlling convergent extension (Greene and Copp, 2014). It also regulates the transcription factors Pax3 and Cdx2 during spinal closure (Zhao et al., 2014). During neural tube closure, Pax3 is expressed in dorsal neuroepithelial cells along the entire length of the anterior-posterior axis (Zhao et al., 2014), and PAX3 mutations have been linked to spina bifida in humans (Agopian et al., 2013). Meanwhile, Cdx2 is essential for caudal body axis elongation during neurulation (Young et al., 2009). In mice, as neurulation begins, Zic2 is expressed in the developing neural tube and migrating neural crest but becomes restricted to the dorsal neural tube once closure is complete (Elms et al., 2003). Mutant embryos lacking Zic2 exhibit reduced levels of BMP antagonists in the dorsal neural plate, which appears to cause a loss of dorsolateral hinge points, thus resulting in severe spina bifida (Nagai et al., 2000; Ybot-Gonzalez et al., 2007). In the curly-tail mouse, which serves as a model for spina bifida, reduced expression of Grhl3 in the caudal region is associated with spinal neural tube defects (Gustavsson et al., 2007). This reduced expression leads to decreased cell growth in the hindgut, causing excessive bending that hinders neural fold closure and can lead to spina bifida or tail flexion defects (Gustavsson et al., 2007).

Mechanisms involved in budding tubulogenesis

The Drosophila tracheal system, which provides oxygen to the organs of the fly, develops through budding tubulogenesis (Hayashi and Kondo, 2018). It comprises a network of sacs with polarized cells surrounding a central lumen (Kerman et al., 2006). The tracheal system originates from ten pairs of tracheal placodes, which form in the dorsal anterior regions of the epidermis in each segment by embryonic stage 10 (Kondo and Hayashi, 2019; Hayashi and Kondo, 2018). These tracheal placodes first appear as cells expressing trachealess (trh), a key regulator of tracheal development (Chung et al., 2011). Next, epidermal growth factor receptor (EGFR) signaling and the rounding up of mitotic cells synergistically drive the budding of these tracheal placodes by creating inward pressure and causing the epithelial sheet to buckle (Kondo and Hayashi, 2013). Finally, fibroblast growth factor (FGF) signaling triggers tracheal branching (Du et al., 2017; Sutherland et al., 1996). During embryonic stage 13, FGF signaling drives the fusion of tracheal placodes, forming a continuous network of tube-like structures resembling the vertebrate blood vessels (Caviglia and Luschnig, 2014).

Drosophila salivary glands provide another example of how a tube is formed through budding. The salivary gland tubes form from a flat, nearly circular epithelial primordium known as the salivary gland placode (Fig. 3A) (Sanchez-Corrales et al., 2021). Budding in this placode is driven by a combination of isotropic apical constriction (in the region known as the ‘pit’, which is where the bud will form) and cell intercalation (away from the pit, in the radial direction), which promote apical cell wedging in the pit (Fig. 3A,B) (Sanchez-Corrales et al., 2021). Apical constriction is mediated by dynamic pools of apical-medial actomyosin, while cell intercalation is mediated by pools of apical-junctional actomyosin (Fig. 3A) (Sanchez-Corrales et al., 2018; Sidor et al., 2020). In addition, a prominent actomyosin cable surrounds the salivary gland placode, aiding tissue budding by generating tension during tube formation (Roper, 2012) (Fig. 3A). It has been suggested that the assembly of the actomyosin cable is directed by the anisotropic localization of the transmembrane protein Crumbs (Crb) (Sidor et al., 2020; Roper, 2012). Crb is highly expressed in placode cells and can form homophilic interactions between neighboring cells in the subapical region, where epithelial cells make contact (Letizia et al., 2013, 2011). These interactions result in Crb being distributed unevenly, especially at the apical side of placode cells near the placode boundary (Roper, 2012). Crb appears to play a dual role: maintaining the apical-basal polarity of epidermal cells and influencing cytoskeletal behavior in the apical domain (Sidor et al., 2020). In fact, changes in Crb levels often occur alongside actomyosin accumulation at these boundaries (Roper, 2012), suggesting that Crb is a key factor in determining the position of the actomyosin cable at the placode boundary (Roper, 2012).

Fig. 3.

Tubulogenesis occurs via budding in Drosophila salivary glands. (A) Left: A schematic of a Drosophila embryo in embryonic stage 11 (A, anterior; D, dorsal; P, posterior; V, ventral). The salivary gland placode (green) is shown below in lateral view. Right: A ventral view of the salivary gland placode showing the flat epithelial sheet. Two cell behaviors drive the isotropic constriction of cells and result in budding of the placode. Isotropic constriction is driven by apical-medial actomyosin (green) in the nascent invagination pit (pale orange), while cell intercalation (inset) occurs away from the pit and is driven by a polarized accumulation of junctional actomyosin (dark blue). Image analyses have shown that active apical intercalation (red arrows) and isotropic constriction (gray arrows in the pit) cause strong wedging of cells near the pit (Sanchez-Corrales et al., 2018). Additionally, a supracellular actomyosin cable forms at the placode boundary (purple), maintaining tension and acting as a tissue-level ratchet that supports the constriction and stabilization of the apical domain within the placode. (B) Cells near the pit have a wedge-like shape, providing a ‘pushing’ force for invagination (red arrows). As the invaginating pit forms, cells located anterior and ventral to the pit constrict dynamically, moving inward into the embryo without active migration (Girdler and Roper, 2014). Cells farther from the pit tilt strongly toward it. Once tissue bending begins at the pit, active apical constriction in and around the pit combines with circumferential intercalations to support the elongation of the pit tube (Sanchez-Corrales et al., 2018). (C) Tube extension and positioning are achieved through the convergent extension of proximal lumen cells (blue arrows), a dynamic process whereby tissue narrows along one axis and elongates along a perpendicular axis. This results in a reduction of the number of cells around the proximal lumen perimeter, where the cells become more cuboidal (blue). Meanwhile, active collective epithelial migration of distal cells (red arrows) further contributes to tube formation. The distal gland cells eventually reach the circular visceral mesoderm (CVM), at which point they initiate a turn so that the elongating tube begins to migrate posteriorly (Bradley et al., 2003; Vining et al., 2005).

Salivary gland tube extension and positioning are achieved through the convergent extension of proximal lumen cells, which is driven by their mediolateral intercalation (Fig. 3C), resulting in a reduction of the number of cells around the proximal lumen perimeter (Xu et al., 2011), where the cells also become more cuboidal (Xu et al., 2008). Meanwhile, active collective epithelial migration of the distal cells of the growing tube further contributes to tube formation (Bradley et al., 2003). The distal gland cells eventually reach the circular visceral mesoderm, at which point they initiate a turn so that the elongating tube begins to migrate posteriorly (Cheshire et al., 2008). This process has been reviewed in detail by Girdler and Roper (2014).

Mechanisms involved in hollowing tubulogenesis

The fundamental cellular and molecular mechanisms of de novo lumen formation via cord hollowing have been extensively researched using MDCK 3D cysts (Bryant and Mostov, 2008). Individual non-polarized MDCK cells have apical and basolateral proteins randomly distributed across their plasma membrane (Fig. 4A) (Bryant et al., 2010). Cell division begins and produces daughter cells that remain in cell–cell contact with each other (Bryant et al., 2010). These cell doublets have two distinct plasma membrane domains: a cell–cell contact surface and an ECM-facing surface (Fig. 4B). The presence of ECM triggers collagen signaling through integrins to Rac1, leading to the assembly of laminin and the loss of apical factors from the outer cell membrane (Liu et al., 2007) (Fig. 4B). The intercellular proteins E-cadherin (cadherin 1) and occludin are located along the cell–cell adhesion zone, while Par-3 (Pard3) is enriched at the periphery (Apodaca, 2010; Bryant et al., 2010) (Fig. 4B). At the end of cytokinesis, the two daughter cells are connected by a transient structure called the midbody (Schluter et al., 2009; Li et al., 2014). Around this area, a region called the apical membrane initiation site (AMIS) forms, where apical proteins are actively transported via Rab11a-positive recycling endosomes (Desclozeaux et al., 2008; Bryant et al., 2010; Schluter et al., 2009) (Fig. 4C). As these apical proteins fuse with the apical membrane, Par-3 is displaced and the pre-apical patch forms (Bryant et al., 2010). The pre-apical patch contains separate apical and junctional domains, which allows factors important for lumen formation to be secreted from the apical side of the cell (Ferrari et al., 2008) (Fig. 4D). Meanwhile, the AMIS matures into TJs, which form a boundary separating the apical and basolateral domains (Fig. 4D) (Bryant et al., 2010; Li et al., 2014).

Fig. 4.

De novo lumen formation in 3D mammalian cell culture. Madin–Darby canine kidney (MDCK) cells cultivated in a three-dimensional extracellular matrix form cysts, consisting of a single layer of epithelial cells surrounding an inner lumen. (A) In non-polarized MDCK cells, apical proteins (yellow) are randomly distributed throughout the cell membrane. (B) Cells sense the signaling mediated by the extracellular matrix and enter mitosis to produce two daughter cells that remain in contact via their cell membranes. Proteins including E-cadherin and occludin (orange) are found along this cell–cell contact zone. Transmembrane proteins such as Crumbs-3a and podocalyxin are internalized into Rab11a endosomes, and the protein Par-3 (green) is localized at the point of contact between the two daughter cells. (C) Rab11a endosomes (yellow) transport apical proteins around the midbody and establish the apical membrane initiation site (AMIS). (D) A new lumen begins to form through the addition of apical proteins (yellow), which displace Par-3 (green) and create a pre-apical patch where opposing plasma membranes are separated. At this stage, the apical membrane marker podocalyxin is present but does not yet form an optically resolvable lumen (Bryant et al., 2010). (E) The secretion of luminal determinants opens the lumen. This is followed by further cell division and an increase in hydrostatic pressure, leading to lumen expansion. The enlarged cross-section on the right shows the proteins responsible for establishing and maintaining cell polarity. The Par complex is crucial for efficiently transporting apical proteins such as Crumbs. Establishing apical-basal polarity involves the assembly and positioning of adherens junctions at the apical side of cells, which link neighboring cells to each other and to the actin cytoskeleton, while tight junctions function as a barrier to prevent diffusion between cells. Afadin and nectin are crucial in regulating adherens junctions and are involved in the timely formation and positioning of the lumen. ECM, extracellular matrix; ER, endoplasmic reticulum; TJ, tight junction.

The location of the AMIS, the orientation of the apical-basal polarity axis and the direction of apical vesicle trafficking are determined during cytokinesis, which acts as a symmetry-breaking event to ultimately determine where the lumen will form (Overeem et al., 2015). In subsequent division rounds, mitotic spindles are oriented to be perpendicular to the apical-basal axis and parallel to the basal lamina (Overeem et al., 2015). The position of the midbody appears to direct Rab11a vesicles containing apical proteins to the AMIS, thus ensuring that the apical surface forms in the correct place (Fig. 4E) (Bryant et al., 2010). If Rab11a expression is disrupted, intracellular accumulation of E-cadherin reduces junctional integrity and inhibits cyst formation (Desclozeaux et al., 2008). The transport of vesicles is also regulated by Cdc42, a Rho-GTPase family member that is a master regulator of cytoskeletal dynamics and cell polarity (Qin et al., 2010). Depletion of specific proteins controlling the mitotic spindle, such as Cdc42 and its guanine nucleotide exchange factors and effectors, leads to defects in apical membrane traffic and misorientation of the spindle (Rodriguez-Fraticelli et al., 2011). This disrupts cleavage furrow orientation and midbody positioning during cytokinesis, resulting in the formation of multiple lumens at the apical surfaces of cells (Rodriguez-Fraticelli et al., 2015; Durgan et al., 2011). The cord-hollowing tubulogenesis factors identified in the MDCK cysts have also been explored in vivo. For instance, in the mouse intestine, Rab11a knockout perturbs apical protein trafficking (Sobajima et al., 2014).

De novo lumen formation by cord hollowing in vivo occurs in mammalian glandular organs, including the pancreas, mammary glands, prostate and salivary glands (Hogan and Kolodziej, 2002). In the developing mouse pancreas, the first sign of lumen formation occurs at embryonic day 11.5 with small, randomly placed microlumens appearing throughout the epithelium (Kesavan et al., 2009). These microlumens are formed by clusters of epithelial cells that share a common surface facing the lumen (Kesavan et al., 2009). They expand as cell polarization spreads, leading to the fusion and rearrangement of microlumens into a connected tube network (Kesavan et al., 2009). Later, this network of cells grows and branches extensively, creating a complex, tree-like structure of pancreatic ducts (Sznurkowska et al., 2018). Cdc42 plays a crucial role in lumen formation in the pancreas; loss of Cdc42 affects the positioning of the multipotent pancreas progenitors, leading to a fragmented pancreatic epithelium and the formation of cellular aggregates without tubules, and a tamoxifen-inducible model for timed ablation of Cdc42 within the pancreatic epithelium showed that Cdc42 is required for maintaining apical polarity in fully polarized tubular epithelial cells (Kesavan et al., 2009).

Mechanisms involved in tube network expansion

The mechanisms underlying tube network expansion in different organs share similarities. For instance, tyrosine kinase pathways, such as FGF and glial cell line-derived neurotrophic factor (GDNF) signaling, play vital roles in the induction of branching in the lung, kidney and mammary glands (Pond et al., 2013; Bates, 2011; Walker et al., 2016; Zhang et al., 2014). The first step of mammalian kidney development is ureteric bud growth, which is induced by GDNF/Ret signaling (Michos et al., 2010; Costantini and Shakya, 2006). GDNF/Ret signaling promotes cell movements in the nephric duct, leading to ureteric bud formation and stimulation of branching that invades the metanephric mesenchyme (Costantini and Shakya, 2006). The absence of upstream regulators of GDNF or Ret causes renal agenesis (Costantini and Kopan, 2010). In the mouse mammary epithelium, the major FGF receptors are expressed during ductal morphogenesis, and mice lacking the FGF receptor number 2 (FGFR2) do not develop mammary buds (Parsa et al., 2008). Studies have shown that functional FGFR signaling is necessary for the maintenance of mammary stem cells and for normal mammary gland development (Parsa et al., 2008; Pond et al., 2013; Zhang et al., 2014).

The role of mechanical forces in tubulogenesis

During early embryonic development, organ identity and positioning are determined by genetic programming, but can be altered by external factors such as mechanical forces. These forces are generated by the intrinsic physical properties of the cells and their environment, and they act as mechanical signals to influence cell polarization and the contractility of the cytoskeleton. It is challenging to measure and visualize physical forces in vivo, and how mechanical forces at the cellular and tissue level can influence the genetic program in early development is an active field of research. Here, we summarize recent work that explores how mechanical forces promote lumen expansion during tubulogenesis.

Hydrostatic pressure promotes lumen expansion

As the lumen forms, it is filled by substances that generate the internal pressure required for lumen expansion (Fig. 5) (Bagnat et al., 2022). These substances can include ions, fluids, solids and gases that are often produced, secreted, or transported by the cells lining the lumen (Bagnat et al., 2022). Substances can accumulate in the lumen via secretory vesicles (Caviglia et al., 2016; Ryan et al., 2019) and by paracellular transport via TJs (Alvers et al., 2014; Bagnat et al., 2007). Typically, these substances are related to the function of the new organ; for instance, in vascular tubulogenesis, fluid shear stress from blood flow and the stretch of vessel walls due to blood pressure are important factors that regulate the development and function of blood vessels (Hahn and Schwartz, 2009) and the secreted components of a chitin-rich ECM that lines the lumen of Drosophila tracheal tubes help regulate the elongation of these tubes along their axis (Ozturk-Colak et al., 2016). Proper regulation of transport into the lumen is essential. For example, in the zebrafish brain, impaired ion transport by the Na⁺/K⁺ ATPase leads to enlarged ventricles as a result of disrupted osmotic pressure (Lowery and Sive, 2005).

Fig. 5.

Physical forces regulate lumen expansion. In the process of de novo lumen formation (such as through cord hollowing), the secretion of cargo via vesicles (yellow circles), paracellular transport (Pt) and transcellular transport (Tt) leads to an increase in the concentration of solutes inside the emerging lumen (Cin). When Cin is higher than the concentration of solutes outside (Cout), an osmotic imbalance is created. This imbalance leads to fluid influx, which generates pressure inside the lumen (Pin) that becomes greater than the pressure outside (Pout). As a result, there is a change in hydrostatic pressure (ΔP). As the lumen grows, Pin increases while Pout remains constant, resulting in increased hydrostatic pressure, which results in further lumen expansion (black arrows) and generates significant wall stress inside the lumen (S) that opposes further influx. As a result of the stretch caused by increased luminal pressure, epithelial thinning (Et; green arrows) occurs. Cell divisions (cd) can relax tension, leading to reduced hydrostatic pressure and increased fluid influx. Various tissues create hydrostatic pressure through fluid or cargo secretion. The mechanisms regulating fluid secretion differ, and the dynamics of lumen formation and maturation depend on the balance between extracellular fluids and cytoskeletal forces.

As hydrostatic pressure increases, fluid transport into the lumen slows down, creating a negative-feedback loop between pressure and transport (Fig. 5). This was shown in the zebrafish inner ear, where imaging and pressure measurements confirmed that hydrostatic pressure and fluid transport work together to control lumen size (Mosaliganti et al., 2019).

ECM properties can regulate lumen expansion and tube elongation

The integrity, length and width of the tubes in the Drosophila tracheal system is regulated by the dynamics of the apical extracellular matrix (aECM), a viscoelastic material inside the lumen that attaches to the apical membrane (Luschnig et al., 2006; Dong et al., 2014). An important component of the aECM is chitin, which begins to accumulate in the tracheal system lumens at stage 14 (Tonning et al., 2005). The accumulation of chitin and other secreted components is thought to generate the force that drives lumen expansion (Luschnig et al., 2006; Dong et al., 2014). In response to secretory activity, F-actin accumulates on the apical plasma membrane facing the lumen, forming parallel cables oriented circumferentially that can exert constrictive forces on the tube (Hannezo et al., 2015; Hayashi and Kondo, 2018; Ozturk-Colak et al., 2016; Kondo et al., 2007). This feedback between F-actin and the aECM prevents excessive expansion of the luminal diameter (Ozturk-Colak et al., 2016; Hayashi and Kondo, 2018). This coupling is also important for regulation of tube elongation, since the degree of luminal diameter expansion influences tube length (Dong et al., 2014).

Mechanistically, changes in the aECM influence actin rearrangement and phosphorylated Src42A levels at cell junctions, a factor necessary for regulation of both tube diameter and tube length (Nelson et al., 2012; Forster and Luschnig, 2012). This suggests that the physical properties of the chitinous ECM regulate the apical cytoskeleton, shaping lumen growth through a feedback loop between the aECM and apical F-actin in tracheal cells (Ozturk-Colak et al., 2016; Nelson et al., 2012; Forster and Luschnig, 2012). Mutations in aECM or septate junction components result in over-elongated tubes, indicating their role in restricting axial elongation (Nelson et al., 2012; Wang et al., 2006; Luschnig et al., 2006; Dong et al., 2014). At embryonic stage 17, just before hatching, the aECM is broken down and absorbed by tracheal cells, clearing most large molecules from the lumen (Tonning et al., 2005). Subsequently, internally produced gas spreads throughout the tracheal lumen, completely filling the tube (Hayashi and Kondo, 2018). In addition to ECM dynamics, cell–cell junctions (Wang et al., 2006) and the apical polarity protein Crb also play crucial roles in tracheal tube elongation and lumen stability (Laprise et al., 2010; Skouloudaki et al., 2019).

Forces generated by flow can modulate tubulogenesis

During vertebrate cardiovascular system development, vasculogenesis forms new arteries from specialized endothelial cells (Jones et al., 2006). This process creates the heart and the initial primitive vascular network inside the embryo and its surrounding membranes (Hahn and Schwartz, 2009). Cells in the cardiovascular system are regularly exposed to hemodynamic forces due to the pressure from the heart's contractions and the resulting blood flow through the circulations (Hahn and Schwartz, 2009). These forces include pressure, which results in circumferential stretching of the vessel wall, and shear stress, which is exerted longitudinally in the direction of blood flow (Hahn and Schwartz, 2009). The importance of these forces was first suggested a century ago, when surgical manipulation of blood flow in avian embryos was shown to induce broad spectrum of heart and blood vessel abnormalities (Jones et al., 2006). Subsequently, experiments in mice demonstrated that disrupting heart function and blood flow can perturb vascular remodeling of the yolk sac and thus lead to embryonic lethality (Wakimoto et al., 2000), and perturbations to shear stress in developing zebrafish result in severe cardiac malformations (Hove et al., 2003).

In mammals, endothelial cells form a large vascular network that wraps around the yolk sac in a branching pattern (Ross and Boroviak, 2020). Once the heart starts beating and blood cells enter circulation, the network reorganizes into a hierarchical, tree-like structure, comprising large arteries, smaller arteries and capillaries, which come together to form veins (Hahn and Schwartz, 2009). This reorganization appears to be promoted by fluid shear stress (Lucitti et al., 2007). For example, reducing shear stress in the developing mouse circulatory system causes issues with atrial contraction, ventricle filling and vascular remodeling (Lucitti et al., 2007). These mice have defects in embryonic circulation, leading to abnormal movement of plasma and blood cells (Lucitti et al., 2007).

YAP (also known as YAP1), a key molecule in Hippo signaling, acts as a mechanotransducer that responds to various mechanical forces affecting the actin cytoskeleton (Piccolo et al., 2014). In vivo experiments in zebrafish and in vitro studies with cultured human endothelial cells showed that mechanical flow signals trigger the movement of YAP into the cell nucleus through changes in the actin cytoskeleton, and this translocation seems to be important for blood vessel stability (Nakajima et al., 2017). In YAP-deficient mice, blood vessel development in the yolk sac is disrupted (Morin-Kensicki et al., 2006). These studies indicate that YAP may play a role in maintaining lumen structure by responding to blood flow (Nakajima et al., 2017).

Flow generated by motile cilia can also influence tubulogenesis. When cilia-dependent fluid movement is disrupted, it can lead to kidney cysts and hydrocephalus in zebrafish (Kramer-Zucker et al., 2005). The cystic fibrosis transmembrane conductance regulator (CFTR) channel, a chloride and bicarbonate channel in vertebrates, works alongside the Na⁺/K⁺ ATPase to drive fluid secretion (Levic and Bagnat, 2023). Beyond its known role in human diseases, CFTR-dependent fluid transport is essential for controlling lumen size during development (Levic and Bagnat, 2023). Mutations in cftr can hinder lumen expansion without affecting cilia formation or movement in the zebrafish Kupffer's vesicle, leading to left-right asymmetry defects (Navis et al., 2013).

Cells can detect mechanical flow through primary cilium signaling (Battle et al., 2015). The primary cilium, a specialized membrane compartment, transduces both mechanical and chemical extracellular signals (Battle et al., 2015). In kidney epithelial cells, primary cilia play a crucial role in sensing flow (Wilson, 2011). Disruptions in primary cilia signaling in mice can result in polycystic kidney disease (PKD), a condition characterized by increased cell proliferation, ectopic cyst formation, and abnormal luminal fluid secretion (Wilson, 2011; Pazour et al., 2002). PKD1 and PKD2 are mutated in the human autosomal dominant form of PKD and encode the proteins polycystin 1 and polycystin 2, respectively (Stayner and Zhou, 2001; Qian et al., 1996). These proteins are expressed in the primary cilia of kidney epithelial cells and in the left-right organizer in vertebrates (Yoshiba et al., 2012; Nauli et al., 2003). Both proteins function together in the same mechanotransduction pathway, sensing fluid flow through the primary cilium (Hilgendorf et al., 2024). It has been proposed that loss of function of these proteins disrupts the ability of cells to detect mechanical cues essential for regulating tissue morphogenesis, leading to the development of PKD (Nauli et al., 2003). The primary mechanism initially proposed for the activation of cilia signaling in response to dynamic fluid movement was an elevation of calcium levels in response to shear stress within the cilia (Nauli et al., 2003). However, it has been shown that mechanical stimulation of primary cilia in kidney epithelia and the left-right organizer does not promote calcium influx (Delling et al., 2016). Although primary cilia are not direct calcium-responsive mechanosensors in the kidney epithelium (Delling et al., 2016), their mechanical stimulation can induce intraciliary calcium signals (Katoh et al., 2023; Djenoune et al., 2023). Furthermore, repeated mechanical stimulation triggers calcium influx, suggesting that polycystin channels are indirectly mechanically gated through chemical cues modulated by mechanical forces (Djenoune et al., 2023; Katoh et al., 2023). This emerging mechanistic understanding of cilia function was outlined in a recent review (Hilgendorf et al., 2024).

Conclusion

Proper formation and maintenance of tubes is vital for the survival of multicellular organisms. Extensive research, both in vitro and in vivo, has uncovered molecular players that perform crucial roles in these processes (Jackson et al., 2017; Lienkamp et al., 2012; Krneta-Stankic et al., 2021; Metzger et al., 2008). Recent studies have shown that physical forces, in conjunction with genetic programming, are essential for the development of functional tubes (Balaghi et al., 2023; Jackson et al., 2017; Menshykau et al., 2019; Yang et al., 2021a,b). However, many unanswered questions remain about how biological tubes are formed. A central question is how cells integrate various mechanical stimuli to form tubular structures with varying shapes across different body plans. Therefore, variations in tubulogenic mechanisms among different groups of organisms can help us distinguish the genetic and mechanical complexities involved in the formation and function of tubes. In addition, it is crucial to understand how cells are assembled into tubes via different modes of tubulogenesis, and how these cells detect their location and respond to various chemical signals and mechanical forces to modify their genetic program and ultimately promote the formation of tubes. Many of the molecular players involved in tube formation have been identified, but how they interact is still largely unknown. Therefore, more research is necessary to comprehend tubulogenesis in living animals and determine how to replicate this process more effectively in laboratory conditions.

Tubulogenesis has evolved to create different types of tubes that are adapted to various functions and environments (Perillo et al., 2023). For example, aquatic species such as fish and frogs do not have branched lungs or kidneys, whereas terrestrial vertebrates have evolved branched lungs that improve oxygen transfer and additional kidney segments that improve the retention of water (Cupello et al., 2022; Wiener, 2023). Comparative research could further enhance our understanding of the evolution of tubulogenic processes (Cain et al., 2020; Corkins et al., 2023).

In addition, interdisciplinary approaches that involve advancements in microscopy, biomechanical analysis, spatial single-cell transcriptomics, organoid culture and mathematical modeling are crucial for improving our understanding of tubulogenesis. For instance, optogenetics allows the coupling of light-sensitive probes to study the spatiotemporal activity of a specific protein within single cells at a subcellular location, thereby identifying how changes in individual cells affect tissue architecture during morphogenesis (Martinez-Ara et al., 2022). Developing methods to study how localized mechanical forces regulate morphogenesis and maintain tubular structures is crucial for advancing our understanding of these processes in vivo. Significant efforts combining optical tweezers, light-sheet microscopy and deep-learning analysis have advanced our understanding of mechanical transduction across some tissues (Katoh et al., 2023; Djenoune et al., 2023). Applying these new technologies could provide valuable insights into how physical forces at the level of single cells and organelles contribute to the shaping and maintenance of tubular structures.

Finally, dysregulation of tubulogenesis can significantly impact morphogenesis and lead to human disease. In this Review, we explored how tubulogenesis has been linked to multiple congenital defects that impact the formation of organs including the vasculature, heart, kidney and neural tube. Tubulogenesis research is therefore crucial in addressing these pressing health issues.