3D in vitro cell culture models of tube formation

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Review

3D in vitro cell culture models of tube formation Mirjam M. Zegers ∗ Radboud University Medical Center, Radboud Institute for Molecular Life Sciences (RIMLS), Department of Cell Biology, P.O. Box 9101, 6500 HB Nijmegen, The Netherlands

a r t i c l e

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Article history: Available online xxx Keywords: Tubulogenesis Branching morphogenesis 3D cell culture In vitro models Extracellular matrix Organ development

a b s t r a c t Building the complex architecture of tubular organs is a highly dynamic process that involves cell migration, polarization, shape changes, adhesion to neighboring cells and the extracellular matrix, physicochemical characteristics of the extracellular matrix and reciprocal signaling with the mesenchyme. Understanding these processes in vivo has been challenging as they take place over extended time periods deep within the developing organism. Here, I will discuss 3D in vitro models that have been crucial to understand many of the molecular and cellular mechanisms and key concepts underlying branching morphogenesis in vivo. © 2014 Elsevier Ltd. All rights reserved.

Contents 1. 2.

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Overview of tube formation in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Mechanism of tube formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Tube architecture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Regulation of branching morphogenesis by growth factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Role of ECM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell culture models for tube formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Common cell lines for in vitro 3D models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Considerations on the choice of ECM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction The functional architecture of organs such as the lung, kidney, pancreas, mammary gland and salivary gland consists of threedimensional (3D) branching networks of epithelial tubes which transport fluids and serve as barriers between physiologically distinct compartments. All tubes are characterized by a central lumen enclosed by polarized epithelial or, in case of the vasculature, specialized epithelial cells called endothelial cells. Often, they end in spherical caps called acini (mammary gland) or alveoli (lung), or, in special cases such as the vasculature, form a continuous endothelial network. The main tubular organs form by branching

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morphogenesis, a highly dynamic and complex developmental process which involves coordinated cell migration, cell shape changes, apical-basal polarization, proliferation, differentiation, apoptosis and reciprocal interactions between the epithelial and the surrounding mesenchyme. Branching morphogenesis is regulated by many transcription factors, hormones, growth factors, and chemical and physical cues from the extracellular matrix (ECM), which often act in a highly localized fashion [1–6]. This complex regulation of branching morphogenesis has complicated the elucidation of cellular and molecular mechanisms underlying this process using in vivo models. In vitro 3D cell culture models of cells grown in ECM recapitulate many aspects of branching morphogenesis and have provided crucial insights in common mechanisms and key concepts involved in tube formation. As the various models currently in use each have their strengths and limitations, the appropriateness of the model depends on which aspects and cellular behaviors

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in branching morphogenesis are being analyzed. Here, I will discuss general design principles and signaling mechanisms involved in tube formation to provide context on factors that should be taken into account when choosing a model. Second, a number of commonly used models based on established cells lines will be discussed, along with strengths and weaknesses of commonly used types of ECM. 2. Overview of tube formation in vivo 2.1. Mechanism of tube formation As described in detail elsewhere [2,7,8], tubes form by different mechanisms, which can be subdivided into two types, depending on state of cell polarization during the process: (1) tubes that emerge from pre-existing polarized epithelial sheets and maintain epithelial polarity and lumen topology during the process, and (2) tubes that rely on de novo polarization during their formation. In processes called wrapping, budding or clefting, tubes form from pre-polarized epithelia. Wrapping involves cell shape changes mediated by apical constriction that cause parts of an epithelial sheet to move inward to form a groove, which eventually seals to form a tube parallel to the originating epithelium. An example of wrapping is neural tube formation. Budding also involves initial invagination of cells via apical constriction, and causes formation of a new tube outward, which subsequently extends by cell migration and/or cell division [2,7]. Examples of budding include branching morphogenesis of organs like the lung [9] and the kidney ureteric bud [4]. Clefting is a variant of budding that subdivides an epithelial bud into two new buds, and is involved in the formation of the mouse salivary gland [10]. Tube formation involving de novo apicalbasal polarization and lumen formation includes processes called cavitation, cord hollowing and cell hollowing. In cavitation, tubes form from a cluster of cells, in which the outer cells will polarize, and luminal space is created by eliminating the inner cells. An example of cavitation includes the clearance of the lumen by apoptosis in the terminal end bud (TEB) of the developing mammary gland [11,12]. In contrast, cord hollowing does not depend on cell death, but involves formation of apical surfaces and small lumens between opposing cells in cords with a diameter of a few cells. Merging of these small initial lumens subsequently separates cell layers and leads to a continuous lumen along the center of the cord. An example of cord hollowing is the formation of the zebrafish gut [13]. Finally, cell hollowing follows the same principle, but involves the merging of a large intracellular vacuolar compartment at adjoining areas of the plasma membrane, which leads to formation of a continuous central lumen extending through the middle of a chain of single cells. The best example of lumen formation by cell hollowing is formation of capillaries of the vasculature [14]. 2.2. Tube architecture The common final architecture of all mature tubes is a circular lumen enclosed by a layer of epithelial cells that are polarized with apical membrane surfaces facing the lumen, basal surfaces contacting the ECM or underlying tissue, and lateral surfaces that interact with adjacent epithelial cells through specialized cell–cell junctions. These junctions include adherens junctions, which physically link cells to each other, and tight junctions, which separate the apical and basolateral membrane domains and provide a tight seal between neighboring cells. Cell polarization is not only critical for the physiological functions of mature tubular organs, including barrier function and vectorial transport across luminal epithelia; it is also required for their initial development. Thus, when tubes form de novo, delivery of apical membrane precedes the formation

of lumens [7,15]. Furthermore, tubes arising from pre-polarized epithelia depend on polarization for apical constriction [16–18], maintenance of cell–cell adhesion and tube extension [19]. In addition, basal targeting of ECM-interacting proteins control adhesion, active remodeling and mechanochemical sensing of ECM, thereby reinforcing apical-basal polarization [20], and together with soluble signals locally control branching morphogenesis [21–23]. 2.3. Regulation of branching morphogenesis by growth factors As a main concept, branching morphogenesis involves locally induced initiation of branching at distal tip areas, coupled to inhibition of branching activity elsewhere [24]. Upon initiation, tubes elongate, which depending on the organ, relies on cell migration, combined with cell elongation, cell proliferation and recruitment of additional cells. Further patterning involves side-branching, or bifurcation, which splits a growing tube into two. Many growth factors, almost exclusively ligands for receptor tyrosine kinases (RTK), promote tube initiation and elongation [1]. For some tubular networks, like the Drosophila tracheal system or the vertebrate vasculature, branching relies on one or two dedicated tip cells, which exhibit protruding filopodia and actively migrate to guide the elongating tube [6]. Tip cells form in response to locally secreted ligands, including fibroblast growth factor (FGF)/Branchless in the trachea and vascular endothelial growth factor (VEGF) in vascular endothelial cells. Cells with the highest receptor activity will become tip cells, which are genetically and morphologically very distinct from stalk cells that are located behind, and linked to tip cells via cell–cell adhesions. Tip cells inhibit induction of additional tips and lateral branching at the stalk via Delta-Notch signaling [6]. Branching of organs and glands that form by budding, such as the lung, kidney ureteric bud and the mammary gland is less well understood. Here, morphologically discernible tip cells are absent, but high branching activity and proliferation rates, and distinct gene expression patterns in distal bud areas compared to regions behind the bud indicate that these branches have functionally distinct tip and stalk or trunk regions as well [25–27]. Branch initiation and elongation of budding tubes depends on inductive and reciprocal interactions with cells and ECM of the surrounding stroma or mesenchyme. The mesenchymal RTK ligands that induce branching [3,5,22] often involve members of the FGF family (lung, kidney, mammary- and salivary glands) [1], but also include glial cell neurotrophic factor (GDNF) (kidney), hepatocyte growth factor (HGF) (lung, mammary gland) [28,29] and members of the epithelial growth factor (EGF) family (mammary gland) [30]. Inhibitory signaling is still poorly understood, but involves complex reciprocal signaling with the mesenchyme. For example, mesenchymally secreted FGF10 initiates outgrowth of lung epithelial buds by FGF receptor 2 (FGFR2)-mediated proliferation and migration of distal tip cells. Negative feedback signaling involves paracrine inhibitory signaling in which FGFR2-induced secretion of Sonic hedgehog (SHH) by distal lung buds inhibits production of FGF10 by the mesenchyme [31]. In addition, FGFR2-induced expression of the RTK antagonist sprouty2 provides an autoinhibitory feedback loop in the lung epithelial buds [9,32]. Reciprocal signaling may also promote positive paracrine feedback signaling, for example during branching of the kidney ureteric bud. Here, branching is mainly induced via the activated Ret receptor in response to secretion of its ligand GDNF by the metanephric mesenchyme [3,33]. Signaling is amplified by Ret-mediated induction of Wnt11 secretion, which increases GDNF expression in the mesenchyme [34]. Intracellular negative regulation includes GDNF-Ret-induced epithelial expression of sprouty1, which inhibits Ret activity [35]. Tubules of the mammary gland mostly form postnatally and involve a bilayered trunk region of a layer of luminal cells covered by a layer of myoepithelial cells, and a distal multilayered TEB, which controls

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branching into the pre-existing stroma of the fat pad. Branching is initiated by many different growth factors, including IGF, HGF, EGF, FGF and is additionally controlled by hormones [30]. Contrary to lung and kidney, the initiation and patterning of mammary branching is not hard-wired. Instead, expansion of branching tubules to fill the fat pad may depend on self-avoidance, possibly via secretion of transforming growth factor ␤ (TGF␤) [24,36,37], which, along with other members of the TGF␤ superfamily are potent inhibitors of branching morphogenesis in many other organs, including kidney [38] and lung [5]. 2.4. Role of ECM The ECM, a complex fibrillar meshwork of collagens, glycoproteins and proteoglycans [39], plays critical roles in branching morphogenesis of many tubular organs beyond its role as a structural scaffold [21,23,40]. Similar to mesenchymal cells, dynamic and reciprocal interactions of the ECM with the budding epithelium [41] and constant remodeling of the ECM by epithelial cells provides inhibitory and stimulatory cues during branching morphogenesis [23,42]. Typical for ECM molecules are their highly localized functions [21]. For example, accumulation of fibronectin at sites of clefting induces local transcriptional downregulation of adherens junctions which drives cleft progression in the salivary gland and lung [43,44]. In particular, spatial regulation of the basement membrane, a thin layer of specialized ECM at the interphase of cells with the mesenchyme consisting of laminins, collagen IV, glycoproteins and proteoglycans, critically regulates branching morphogenesis, and remodeling of basement membrane, mostly via metalloproteases of the MMP and ADAM families, is most dynamic at distal tip buds [23,42,45,46]. The overlapping chemical and physical functions of ECM molecules include their role as ligands for adhesion receptors, mostly integrins and syndecans [40], which activate signaling cascades controlling cell migration, polarization, proliferation and cell survival [47] via pathways that often crosstalk with those downstream of RTK [48]. The ECM also binds and acts as a sink for growth factors and reservoir for bioactive ECM cleavage products, allowing for controlled release by proteolytic enzymes, presentation of these factors, and the generation of haptotactic gradients [39,40]. A key aspect of ECM in morphogenesis are its physical properties, in particular its stiffness, which depends on the organization and crosslinking of collagen I fibers [49]. Stiffness is sensed by epithelial cells by integrins, which link to intracellular contractile actomyosin. Increasing ECM stiffness is counterbalanced by increased intracellular contractile forces, which, via a cascade of mechanotransduction processes that also involve supracellular force transmissions via cell–cell adhesion, collectively control cell migration, proliferation and apical-basal polarization [23,40]. Finally, dense ECM may provide a barrier inhibiting directional tube growth and possibly provide tracks to guide migration. For instance, local accumulation of collagen stabilizes tissue during clefting of the salivary gland [21], whereas thickening of ECM along the trunk area of the mammary tube may prevent side budding and propel the moving TEB forward [50]. Thus, rather than a passive structural scaffold, the ECM cooperates with epithelia to build the tubular architecture of organs. 3. Cell culture models for tube formation As discussed above, tube formation involves many dynamic cellular processes, depends on the chemicophysical characteristics of the ECM, and takes place over extended time periods deep within the developing organism. These factors combined make it difficult to understand the molecular and cellular mechanisms and key concepts underlying branching morphogenesis using in vivo models.

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Traditional cell culture studies in cell culture plastic on the other hand, lack the influence of the ECM and are inheritably 2D, and are therefore insufficient to understand branching morphogenesis. To bridge this gap, 3D cell culture models have been developed. These models include organs grown ex vivo, epithelial tissue fragments stripped from mesenchyme, often called organoids [2], and simplified models of established cell lines, the latter of which will be discussed here. Several 3D in vitro models of cell lines grown in different types of purified ECM, most commonly collagen I, or reconstituted basement membrane extract (BME) have been established (Table 1, Section 3.2). While none of these models faithfully recapitulate the development of these organs, they have been extraordinary useful to delineate common molecular pathways underlying specific aspects of branching morphogenesis, including initiation of branching, apical-basal cell polarization, and the different mechanisms leading to lumen formation. However, no single in vitro model will be able to incorporate all relevant parameters and the choice of cells, ECM and other mesenchymal factors will greatly determine outcome. Table 1 summarizes several widely used model cell lines and summarizes their ability to form lumens and their response to branching initiating- and inhibiting mesenchymal factors in the context of different types of ECM. 3.1. Common cell lines for in vitro 3D models Cell lines used for in vitro 3D tubulogenesis models are often from the mammary gland and the kidney (Table 1), of which the MCF10A and MDCK cell lines are the most widely used. MCF10A is a spontaneously immortalized, nontransformed human cell line that exhibits features of both luminal and myoepithelial mammary cells [51]. When grown on top of a layer of BME, covered with medium supplemented with a low concentration of BME (2.5D), they form uniformly sized, growth-arrested spheres, called acini, of a single layer of cells enclosing a lumen [51]. These acini develop in ∼15 days via an initial proliferative stage, followed by lumen formation by cavitation, which depends on apoptosis [52] and autophagy [53]. While lumen formation in general is intrinsically linked to apical-basal polarization, this relation is unclear in MCF10A, which, despite having a polarized cytoplasm and secrete ECM basally, lack tight junctions and thus incompletely polarize [54,55]. Perhaps as a result, MCF10A do not form discernible lumens in collagen [56] (Zegers, unpublished data). Whereas MCF10A do not branch in BME, in 3D collagen they form branching cords with distal tips that merge upon contact with other cell clusters or branches [57] (Zegers, unpublished data), likely mediated by direct transmission of traction force along collagen fibers [57]. Therefore, MCF10A is particularly useful as model for cavitation in the context of a BME matrix, but not for tubular outgrowth. Alternatives are the spontaneously immortalized mouse mammary cell lines HC11 [58] and, in particular, EpH4. EpH4 cells are well polarized, form lumens, and branch into lumen-containing tubules in both collagen and BME upon addition of growth factors. However, despite being clonally derived, EpH4 show considerable morphogenetic heterogeneity [59], which complicates analyses of genetic gain- or loss of function approaches and comparison of results from different laboratories. Thus, taken together, while available mammary cell lines are well suited to analyze cell polarization and lumen formation in vitro (Table 1), analyses of cell behaviors associated with tubular outgrowth have been more problematic. This may relate to the use of a single cell type, whereas mammary gland branching involves both luminal and myoepithelial cells [22,60]. As alternative, organoid branching models, consisting of epithelial organ fragments embedded in ECM, may hold better promise, although they come with their own set of caveats, discussed elsewhere [2]. Nevertheless, the organoid model has shown to be very useful to understand cell dynamics in mammary branching, as was shown recently in

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Cell line (species)

Lumens in collagen

Lumens in/on BME

Branching in collagen

Branching in BME

Mesenchymal/stromal inducer of branching

Mesenchymal/stromal inhibitors of branching

Refs.

Kidney

MDCK (canine)

Yes

Yes

Yes

Only initial tip cells

Col: HGF, fibronectin BME: HGF (only tip cells)

Tgf␤, IL-8 Collagen IV, TIMP-2 vitronectin, HSPG, ephrinA1

[63,64,66,70– 74,79,80,82,96]

Kidney

mIMCD3 (mouse)

Yes

Yes

Yes

Yes (in BME/Coll mix)

Col: 5% FCS, HGF, TGF␣, EGF BME/Coll: HGF

TBF␤ KC (mouse IL8 homolog)

[97–99]

Lung

VA10 (human)

Yes

BME: Co-culturing HUVECs, FGFR-dependent

[100]

Mammary gland

MCF10A (human)

Col: HGF, mechanical force,

[51,56,101]

Yes

No

Yes

Yes

[54,102,103]

Mammary gland

HMT-3522-S1 (human)

No

Yes

Mammary gland

Eph4 (mouse)

Mostly no. Heterogeneous

Yes, Heterogeneous

Yes

Mammary gland

NuMuG (mouse)

Yes

Yes

Mammary gland

HC11 (mouse)

No

Yes

Yes

Yes

Col: HGF, EGF, FGF2, epimorphin, TGF␤ (low concentration) BME: HGF

TGF␤, TIMP-2

[36,46,59,60, 104–106]

Col: Spontaneous, HGF

[56,107]

BME: Constitutive FGFR1 activation

[58], *

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Table 1 Commonly used in vitro 3D models. The capability of these lines to form lumens and branching tubules in collagen I and BME matrices is indicated, along with extracellular mesenchymal factors inducing or inhibiting branching. BME, basement membrane extract; Col, collagen I; HSPG, heparan sulfate proteoglycan; HUVECs, Human vascular endothelial cells; KC, Keratinocyte chemoattractant; * (Zegers, unpublished data); “empty”, unknown.

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a study using long-term, live cell imaging, which revealed intricate details on cell polarization, migration and lumen during mammary branching [45]. The MDCK cell line has properties of the kidney distal tubule and collecting duct. It is the most versatile in vitro model for tubulogenesis, which, depending on the culture conditions, exhibits distinct mechanisms driving lumen formation and branching morphogenesis, including cavitation, hollowing, tip cell formation and budding (Figs. 1 and 2). Single cells, embedded into a BME or collagen I matrix will proliferate, polarize de novo and form cysts of highly polarized cells enclosing a single lumen (Fig. 1) [61]. A defining difference between collagen and BME is the presence of laminin in BME, which primes assembly of a basement membrane around the surface of the developing cyst, which in turn orients and reinforces apical-basal polarization [62]. In collagen, cells depend on their own secreted laminin to assemble a basement membrane via a process that involves ␤1-integrin-mediated stimulation of Rac1 [20,62] and downregulation of Rho-ROCK-dependent actomyosin contraction [63]. Cells therefore polarize rapidly in BME and form a single lumen in 2–3 days, whereas polarization and lumen formation in collagen I is slower, and takes 10–12 days [64]. These differences also affect how lumens form, which occurs through hollowing with no associated apoptosis in BME, and by apoptosis-dependent cavitation in collagen I [64] (Fig. 1). Spatial cues provided by the extracellular environment control apicalbasal polarization. One cue is provided by the ECM, which, via pathways that critically rely on ␤1-integrins, cause formation and orientation of an apical surface opposite to the cell–ECM contact [20,62,63,65]. A second spatial cue is provided by neighboring cells, likely transduced by components of cadherin-based adherens junctions [19,65–68]. Downstream of these cues, polarity complexes and regulators of vesicular traffic act together to target specific apical and basolateral membrane proteins to the appropriate membrane surfaces. In addition, pathways downstream of cdc42 control the orientation of the mitotic spindle perpendicular to the apical-basal axis, thereby promoting and maintaining formation of lumens. Details on the regulation of these processes have been covered in many recent excellent reviews [15,68,69]. In regular growth medium with serum, MDCK cysts with single lumens form reliably and robustly in either ECM and comprise >95% of cellular structures, without any branching activity [20]. Addition of HGF, or conditioned growth medium from the embryonic mesenchymal cell line MRC-5, which contains high levels of HGF, induces elaborate branching from cysts in collagen [70,71]. Branching occurs over a time course of several days in distinct stages, named extensions, chains, cords and tubules [61,71], and is controlled by molecular pathways that are beginning to emerge (Fig. 2). Extensions are basolateral protrusions from cells that retain an apical surface continuous with the cyst wall. They exhibit dynamic extension and retraction behavior, display additional migratory front–rear polarization [72], and may be the functional equivalent of tip cells. Extensions are promoted by the presence of fibronectin in the ECM [73], and by intracellular signaling molecules associated with PI3K and Rac1 signaling, including Pak1, ␤PIX, Scribble and ARF6 [72,74–76], by transcription factors, including TSN4 and Stat3 [77] and by ERK signaling [76,78]. Branching is inhibited by extracellular factors that include collagen IV [73], TGF␤ [73], IL8 [79] and ephrinA1 [80]. Next, some extensions will mature into chains of single cells and subsequent cords of 2–3 cells in diameter that extend into the collagen. As compared to extensions, chains and cords appear at a ∼10 fold lower rate and typically range 2–5 per cyst. Cells in cords loose apical polarity while maintaining cell–cell contacts via adherens and tight junctions [71]. Cord and chain formation depends on proliferation and active migration and is promoted by PI3K, ERK [72,78], Stat1 and Stat3 [81] and requires ECM remodeling via MMPs [78,82]. The mechanisms controlling

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whether an extension retracts or matures is currently not clear, but likely involves Rho-ROCK-dependent signaling, since inhibition of ROCK highly increases extension formation, which subsequently fail to mature [72,80]. Extracellular factors inhibiting chain formation include the discoidin domain receptor 1 (DDR1), an RTK for collagen ligands, which inhibits Stat1/3 activation downstream of ␣2␤1 integrin [81]. In the final stages of tube formation, cells within cords re-polarize leading to the formation of small apical surfaces by hollowing, which merge and eventually become continuous with the central lumen, at which time apical-basolateral polarization is entirely restored. This process, which depends on Stat1 [83], is currently not well understood. Since cords are surrounded by assembled laminin (Zegers, unpublished data) and repolarize in 1–2 days, it is tempting to speculate and worth investigating whether cord hollowing is regulated by the same mechanisms that regulate cyst hollowing of cells of cysts in BME. Alternatively, tubes can form by budding (Fig. 2), with apicalbasolateral polarization retained, either in response to HGF [84] in 3D culture or in a modified 2.5 D configuration, on top of a thick collagen layer containing HGF-secreting mesenchymal cells [77]. In the absence of HGF, budding is induced by the Rac effector Pak1 and ␤PIX [75] or by downregulation of the kidney-specific cadherin-6 [66]. Tube formation by either budding or cord hollowing appears to be mutually exclusive, as experimental conditions that promote tube formation by a combination of both processes have not been reported. This indicates that the mechanisms driving either form of tube formation are distinct and robust, and may depend on one or more critical factors that remain to be identified. Parameters acting upstream of these distinct mechanisms may include relative abundance of signaling pathways initiated by cell–cell and cell–ECM contacts, which is controlled by Pak1 and cadherin-6 [66,85]. 3.2. Considerations on the choice of ECM The choice of ECM greatly affects outcome in 3D models. This is mostly apparent when comparing collagen I and BME, with many models relying on BME to polarize and form lumens (Table 1, Section 3.1). BME is generally prepared from the high amounts of basement membrane proteins produced by the Engelbreth–Holm–Swarm tumor, available as a commercial preparation called Matrigel. It mainly consists of laminins, collagen IV, heparan sulfate proteoglycan (HSPG) and entactin/nidogen [86]. Despite the high levels of these proteins, BME lacks the normal architecture of a basement membrane and displays poor mechanical properties [87]. Most likely, laminins in BME are permissive for initial cell polarization, but cellular assembly of native basement membrane is required to enable full polarization [55,62]. Considering that ECM acts as a sink for many stromal factors, it is not surprising to find these in BME. Indeed, many growth factors (FGF, EGF, PDGF, IGF, TGF␤), proteases (MMP-2, MMP-9, urokinase, tissue type plasminogen activator) and other factors have been in reported in BME, including available growth-factor reduced versions [88,89]. Another concern is variability between different lots, exemplified by recent proteomic analyses of Matrigel preparations, which showed the presence of a daunting ∼1500 unique proteins and only a 53% lot-to-lot consistency [88]. Additional disadvantages of BME are its tumor origin, thereby possibly promoting aberrant growth characteristics and its considerable expense. In contrast to the complex protein composition of BME, matrices from polymerized collagen I are relatively pure [87]. However, both the source material and polymerization conditions can cause considerable variation in ECM architecture. The most widely used types of collagen I commercially available are from either rat tails or bovine skin, for which the extraction and purification differs substantially [49,87]. A main distinction between rat tail collagen and bovine skin collagen is that the latter is more highly crosslinked

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Fig. 1. MDCK cysts form lumens by cavitation or hollowing. Top, left to right: single MDCK cells, embedded in a collagen I matrix, secrete and assemble laminin into a basement membrane (blue), which is required for the formation of small lumens lined by apical membrane domains (pink) and subsequent formation of a single lumen by cavitation, involving inner cell death. In contrast, cells plated in laminin-rich BME (bottom, left to right), polarize much faster and form lumens by hollowing.

Fig. 2. Branching morphogenesis of MDCK cysts. Polarized MDCK cysts can form branching tubes by budding after addition of HGF, or in the absence of HGF by downregulation of cadherin-6 (top row). Mostly however, HGF induces tubulogenesis by tip cell selection, partial depolarization, and cord hollowing and repolarization. Apical membrane surfaces are indicated in pink. See text for details.

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and requires pepsin treatment for extraction. As a consequence, when compared to non-pepsinized rat tail collagen, polymerization of pepsinized bovine skin collagen into a 3D fibrillar gel at equal concentrations is slower, generates longer and thicker fibrils, and generates a softer gel with larger pores [49,87]. Additional variables affecting polymerization include concentration, pH, ionic strength, temperature and the age of the source-organism [49,87]. The resulting variations in architecture of the collagen I matrix strongly affect cell migration and tubulogenesis [90,91]. For instance, the porosity of the ECM determines whether cells employ MMP activity to degrade ECM in order to migrate into the ECM [90], which in turn may also affect chemotactic and haptotactic signaling. In addition, the architecture of collagen fibers globally impacts tubulogenesis via mechanotransduction, as it was shown that bidirectional force transmission between cells and collagen I, and resulting collagen fiber alignment is primary factor inducing tubulogenesis in the MCF10A model [57]. Thus, whereas variation in BME mostly lies is its chemical composition, variations in the physical structure of different collagen I matrices are equally important. 4. Conclusion 3D models of a single cell type in a simple biologically derived ECM like collagen I, optionally supplemented with a mesenchymally derived factor to induce branching, recapitulate mechanisms involved in cell polarization, lumen formation and initial branching to a degree that is surprisingly similar to more complex ex vivo and in vivo models, such as mammary and kidney organoids [12,45,60,67] and the zebrafish gut [13]. These models, however, are not well suited to assess roles of spatial physicochemical heterogeneity of ECM and reciprocal interactions with mesenchymal cells, and the associated exact spatial patterning of branching. Furthermore, these models do not allow analysis of roles of fluid flow mediated shear forces that play important roles in the development of tubular development of organs like the heart [92] and nephron [93]. Recent advances in microfabrication techniques capable of positioning cells and ECM at micrometer scales, hold great promise for analyzing the roles of the physicochemical environment in organ branching, particularly when combined with microfluidics allowing for perfusion of microengineered tubes or “organ-onchips” [94]. Furthermore, additional multidisciplinary approaches, including in silico modeling, for instance to distinguish relative importance of cell behaviors in branching [95] will be necessary to improve on currently available 3D models. Acknowledgements I want to thank Martin ter Beest for critically reading this manuscript and for his help in preparing the figures. MZ is supported by the European Commission FP7-PEOPLE-2011-IIF (302067). References [1] Lu P, Werb Z. Patterning mechanisms of branched organs. Science 2008;322:1506–9. [2] Andrew DJ, Ewald AJ. Morphogenesis of epithelial tubes: insights into tube formation, elongation, and elaboration. Dev Biol 2010;341:34–55. [3] Costantini F, Kopan R. Patterning a complex organ: branching morphogenesis and nephron segmentation in kidney development. Dev Cell 2010;18:698–712. [4] Dressler GR. Advances in early kidney specification, development and patterning. Development 2009;136:3863–74. [5] Morrisey EE, Hogan BL. Preparing for the first breath: genetic and cellular mechanisms in lung development. Dev Cell 2010;18:8–23. [6] Ochoa-Espinosa A, Affolter M. Branching morphogenesis: from cells to organs and back. Cold Spring Harb Perspect Biol 2012:4. [7] Lubarsky B, Krasnow MA. Tube morphogenesis: making and shaping biological tubes. Cell 2003;112:19–28.

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Please cite this article in press as: Zegers MM. 3D in vitro cell culture models of tube formation. Semin Cell Dev Biol (2014), http://dx.doi.org/10.1016/j.semcdb.2014.02.016