AUXIN RESPONSE FACTOR17 Is Essential for Pollen Wall Pattern Formation in Arabidopsis

In angiosperms, pollen wall pattern formation is determined by primexine deposition on the microspores. Here, we show that AUXIN RESPONSE FACTOR17 (ARF17) is essential for primexine formation and pollen development in Arabidopsis (Arabidopsis thaliana). The arf17 mutant exhibited a male-sterile phenotype with normal vegetative growth. ARF17 was expressed in microsporocytes and microgametophytes from meiosis to the bicellular microspore stage. Transmission electron microscopy analysis showed that primexine was absent in the arf17 mutant, which leads to pollen wall-patterning defects and pollen degradation. Callose deposition was also significantly reduced in the arf17 mutant, and the expression of CALLOSE SYNTHASE5 (CalS5), the major gene for callose biosynthesis, was approximately 10% that of the wild type. Chromatin immunoprecipitation and electrophoretic mobility shift assays showed that ARF17 can directly bind to the CalS5 promoter. As indicated by the expression of DR5-driven green fluorescent protein, which is an synthetic auxin response reporter, auxin signaling appeared to be specifically impaired in arf17 anthers. Taken together, our results suggest that ARF17 is essential for pollen wall patterning in Arabidopsis by modulating primexine formation at least partially through direct regulation of CalS5 gene expression.In angiosperms, the pollen wall is the most complex plant cell wall. It consists of the inner wall, the intine, and the outer wall, the exine. The exine is further divided into sexine and nexine layers. The sculptured sexine includes three major parts: baculum, tectum, and tryphine (Heslop-Harrison, 1971; Piffanelli et al., 1998; Ariizumi and Toriyama, 2011; Fig. 1A). Production of a functional pollen wall requires the precise spatial and temporal cooperation of gametophytic and sporophytic tissues and metabolic events (Blackmore et al., 2007). The intine layer is controlled gametophytically, while the exine is regulated sporophytically. The sporophytic tapetum cells provide material for pollen wall formation, while primexine determines pollen wall patterning (Heslop-Harrison, 1968).Open in a separate windowFigure 1.Schematic representation of the pollen wall and primexine development. A, The innermost layer adjacent to the plasma membrane is the intine. The bacula (Ba), tectum (Te), and tryphine (T) make up the sexine layer. The nexine is located between the intine and the sexine layers. The exine includes the nexine and sexine layers. B, Primexine (Pr) appears between callose (Cl) and plasma membrane (Pm) at the early tetrad stage (left panel). Subsequently, the plasma membrane becomes undulated (middle panel) and sporopollenin deposits on the peak of the undulated plasma membrane to form bacula and tectum (right panel).After meiosis, four microspores were encased in callose to form a tetrad. Subsequently, the primexine develops between the callose layer and the microspore membrane (Fig. 1B), and the microspore plasma membrane becomes undulated (Fig. 1B; Fitzgerald and Knox, 1995; Southworth and Jernstedt, 1995). Sporopollenin precursors then accumulate on the peak of the undulated microspore membrane to form the bacula and tectum (Fig. 1B; Fitzgerald and Knox, 1995). After callose degradation, individual microspores are released from the tetrad, and the bacula and tectum continue to grow into exine with further sporopollenin deposition (Fitzgerald and Knox, 1995; Blackmore et al., 2007).The callose has been reported to affect primexine deposition and pollen wall pattern formation. The peripheral callose layer, secreted by the microsporocyte, acts as the mold for primexine (Waterkeyn and Bienfait, 1970; Heslop-Harrison, 1971). CALLOSE SYNTHASE5 (CalS5) is the major enzyme responsible for the biosynthesis of the callose peripheral of the tetrad (Dong et al., 2005; Nishikawa et al., 2005). Mutation of Cals5 and abnormal CalS5 pre-mRNA splicing resulted in defective peripheral callose deposition and primexine formation (Dong et al., 2005; Nishikawa et al., 2005; Huang et al., 2013). Besides CalS5, four membrane-associated proteins have also been reported to be involved in primexine formation: DEFECTIVE EXINE FORMATION1 (DEX1; Paxson-Sowders et al., 1997, 2001), NO EXINE FORMATION1 (NEF1; Ariizumi et al., 2004), RUPTURED POLLEN GRAIN1 (RPG1; Guan et al., 2008; Sun et al., 2013), and NO PRIMEXINE AND PLASMA MEMBRANE UNDULATION (NPU; Chang et al., 2012). Mutation of DEX1 results in delayed primexine formation (Paxson-Sowders et al., 2001). The primexine in nef1 is coarse compared with the wild type (Ariizumi et al., 2004). The loss-of-function rpg1 shows reduced primexine deposition (Guan et al., 2008; Sun et al., 2013), while the npu mutant does not deposit any primexine (Chang et al., 2012). Recently, it was reported that Arabidopsis (Arabidopsis thaliana) CYCLIN-DEPENDENT KINASE G1 (CDKG1) associates with the spliceosome to regulate the CalS5 pre-mRNA splicing for pollen wall formation (Huang et al., 2013). Clearly, disrupted primexine deposition leads to aberrant pollen wall patterning and ruptured pollen grains in these mutants.The plant hormone auxin has multiple roles in plant reproductive development (Aloni et al., 2006; Sundberg and Østergaard, 2009). Knocking out the two auxin biosynthesis genes, YUC2 and YUC6, caused an essentially sterile phenotype in Arabidopsis (Cheng et al., 2006). Auxin transport is essential for anther development; defects in auxin flow in anther filaments resulted in abnormal pollen mitosis and pollen development (Feng et al., 2006). Ding et al. (2012) showed that the endoplasmic reticulum-localized auxin transporter PIN8 regulates auxin homeostasis and male gametophyte development in Arabidopsis. Evidence for the localization, biosynthesis, and transport of auxin indicates that auxin regulates anther dehiscence, pollen maturation, and filament elongation during late anther development (Cecchetti et al., 2004, 2008). The role of auxin in pollen wall development has not been reported.The auxin signaling pathway requires the auxin response factor (ARF) family proteins (Quint and Gray, 2006; Guilfoyle and Hagen, 2007; Mockaitis and Estelle, 2008; Vanneste and Friml, 2009). ARF proteins can either activate or repress the expression of target genes by directly binding to auxin response elements (AuxRE; TGTCTC/GAGACA) in the promoters (Ulmasov et al., 1999; Tiwari et al., 2003). The Arabidopsis ARF family contains 23 members. A subgroup in the ARF family, ARF10, ARF16, and ARF17, are targets of miRNA160 (Okushima et al., 2005b; Wang et al., 2005). Plants expressing miR160-resistant ARF17 exhibited pleiotropic developmental defects, including abnormal stamen structure and reduced fertility (Mallory et al., 2005). This indicates a potential role for ARF17 in plant fertility, although the detailed function remains unknown. In addition, ARF17 was also proposed to negatively regulate adventitious root formation (Sorin et al., 2005; Gutierrez et al., 2009), although an ARF17 knockout mutant was not reported and its phenotype is unknown.In this work, we isolated and characterized a loss-of-function mutant of ARF17. Results from cytological observations suggest that ARF17 controls callose biosynthesis and primexine deposition. Consistent with this, the ARF17 protein is highly abundant in microsporocytes and tetrads. Furthermore, we demonstrate that the ARF17 protein is able to bind the promoter region of CalS5. Our results suggest that ARF17 regulates pollen wall pattern formation in Arabidopsis.     

Pharmacological targeting of guanosine monophosphate synthase suppresses melanoma cell invasion and tumorigenicity

Malignant melanoma possesses one of the highest metastatic potentials among human cancers. Acquisition of invasive phenotypes is a prerequisite for melanoma metastases. Elucidation of the molecular mechanisms underlying melanoma invasion will greatly enhance the design of novel agents for melanoma therapeutic intervention. Here, we report that guanosine monophosphate synthase (GMPS), an enzyme required for the de novo biosynthesis of GMP, has a major role in invasion and tumorigenicity of cells derived from either BRAF^V600E or NRAS^Q61R human metastatic melanomas. Moreover, GMPS levels are increased in metastatic human melanoma specimens compared with primary melanomas arguing that GMPS is an attractive candidate for anti-melanoma therapy. Accordingly, for the first time we demonstrate that angustmycin A, a nucleoside-analog inhibitor of GMPS produced by Streptomyces hygroscopius efficiently suppresses melanoma cell invasion in vitro and tumorigenicity in immunocompromised mice. Our data identify GMPS as a powerful driver of melanoma cell invasion and warrant further investigation of angustmycin A as a novel anti-melanoma agent.Malignant melanoma is one of the most aggressive types of human cancers. Its ability to metastasize in combination with resistance to conventional anticancer chemotherapy makes melanoma extremely difficult to cure, and the median survival of patients with metastatic melanoma is 8.5 months.^{1, 2, 3} A better understanding of the biology behind melanoma aggressiveness is imperative to facilitate the development of novel anti-melanoma strategies.Melanoma and other cancers cells have been shown to strongly rely on de novo nucleotide biosynthesis^{4, 5} and often overexpress several biosynthetic enzymes involved in these pathways.^{6, 7, 8, 9} Recently, we have identified a fundamental connection between melanoma invasion and biosynthesis of guanylates,⁸ suggesting that distortion of the guanylate metabolism facilitates melanoma progression.Guanosine monophosphate reductase (GMPR) reduces GMP to one of its precursors, inosine monophosphate (IMP), and depletes intracellular GTP pools (Figure 1a). We have recently demonstrated that GMPR suppresses melanoma cell invasion and growth of human melanoma cell xenografts. These findings tightly linked guanylate production to the invasive potential of melanoma cells.⁸Open in a separate windowFigure 1GMPS contributes to the invasive capability of melanoma cells. (a) Simplified schematic of the metabolic pathway for guanylates production. (b) SK-Mel-103 and SK-Mel-28 cells were transduced with a control vector or two independent shRNAs to GMPS and tested for invasion through Matrigel (left panel). Where indicated, cells were incubated with 100 μM guanosine for 24 h before the assay and guanosine supplementation was maintained throughout the experimental procedure. The data represent the average ± S.E.M. of at least two independent experiments. GMPS suppression was verified by immunoblotting (right panel). (c) Cells transduced as in (a) were plated on coverslips coated with FITC-conjugated gelatin. After 16 h cells were fixed with 4% PFA and stained for actin (rhodamine-conjugated phallodin) and nuclei (Hoechst). Where indicated, cells were incubated with 100 μM guanosine for 24 h before the assay and guanosine supplementation was maintained throughout the experimental procedure. At least 25 cells/sample were imaged to assess the number of cells with gelatin degradation. The data represent the average ± S.E.M. of two independent experiments. *P<0.05, **P<0.001 compared with control; ^#P<0.05, ^##P<0.001 compared with untreated cells. Statistics performed with Student''s t-Test. See also Supplementary Figure S1Of the several enzymes involved in guanylate biosynthesis, inositol monophosphate dehydrogenases 1 and 2 (IMPDH-1, -2), functional antagonists of GMPR (Figure 1a), have been targeted clinically with several drugs including the most specific one, mycophenolic acid (MPA) and its salt, mycophenolate mofetil (MMF).^{10, 11, 12, 13} Nonetheless, prior studies demonstrated that MPA possesses poor anti-tumor activity,^{14, 15} and it is primarily used as an immunosuppressing agent in organ transplantation.^{10, 11, 12}GMP synthase (GMPS) is the other functional antagonist of GMPR. GMPS catalyzes the amination of xanitol monophosphate (XMP) to GMP to promote GTP synthesis (Figure 1a).^{16, 17} Most of the studies on GMPS have been performed in bacteria, yeast, and insects, where GMPS has been shown to have a key role in sporulation, pathogenicity, and axon guidance, respectively.^{18, 19, 20} Mammalian GMPS has been the subject of several studies addressing its unconventional (GMP-unrelated) roles in the regulation of activity of ubiquitin-specific protease 7 (USP7).^{21, 22, 23, 24} However, because of the newly revealed importance of guanylate metabolism in control of melanoma cell invasion and tumorigenicity,⁸ GMPS emerges as an attractive target for anti-cancer therapy.In the late 1950s, a specific inhibitor of bacterial GMPS, angustmycin A (also known as decoyinine), has been isolated from Streptomyces hygroscopius as a potential antibiotic with sporulation-inducing activity in Bacillus subtilis.^{25, 26, 27, 28, 29} Its anti-tumor activity has never been experimentally explored. In the current study, we investigated the role of GMPS in regulation of melanoma invasion and tumorigenicity, and explored the possibility of targeting GMPS by angustmycin A as a novel anti-melanoma strategy.     

The Presence of Fucogalactoxyloglucan and Its Synthesis in Rice Indicates Conserved Functional Importance in Plants

The predominant structure of the hemicellulose xyloglucan (XyG) found in the cell walls of dicots is a fucogalactoXyG with an XXXG core motif, whereas in the Poaceae (grasses and cereals), the structure of XyG is less xylosylated (XXGG_n core motif) and lacks fucosyl residues. However, specialized tissues of rice (Oryza sativa) also contain fucogalactoXyG. Orthologous genes of the fucogalactoXyG biosynthetic machinery of Arabidopsis (Arabidopsis thaliana) are present in the rice genome. Expression of these rice genes, including fucosyl-, galactosyl-, and acetyltransferases, in the corresponding Arabidopsis mutants confirmed their activity and substrate specificity, indicating that plants in the Poaceae family have the ability to synthesize fucogalactoXyG in vivo. The data presented here provide support for a functional conservation of XyG structure in higher plants.The plant cell wall protects and structurally supports plant cells. The wall consists of a variety of polymers, including polysaccharides, the polyphenol lignin, and glycoproteins. One of the major polysaccharides present in the primary walls (i.e. walls of growing cells) in dicots is xyloglucan (XyG), which consists of a β-1,4-glucan backbone with xylosyl substituents. XyG binds noncovalently to cellulose microfibrils and thereby, is thought to act as a spacer molecule, hindering cellulose microfibrils to aggregate (Carpita and Gibeaut, 1993; Pauly et al., 1999a; Bootten et al., 2004; Cosgrove, 2005; Hayashi and Kaida, 2011; Park and Cosgrove, 2012).The side-chain substitutions on XyG can be structurally diverse depending on plant species, tissue type, and developmental stage of the tissue (Pauly et al., 2001; Hoffman et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009, 2012; Lampugnani et al., 2013; Schultink et al., 2014). A one-letter code nomenclature has been established to specify the XyG side-chain substitutions (Fry et al., 1993; Tuomivaara et al., 20145). According to this nomenclature, an unsubstituted glucosyl residue is indicated by a G, whereas a glucosyl residue substituted with a xylosyl moiety is shown as an X. In most dicots, the xylosyl residue can be further substituted with a galactosyl residue (L), which in turn, can be further decorated with a fucosyl residue (F) and/or an acetyl group (F/L). In some species, the xylosyl residue can be substituted with an arabinosyl moiety (S), and the backbone glucosyl residue can be O-acetylated (G; Jia et al., 2003; Hoffman et al., 2005).Numerous genes have been identified in Arabidopsis (Arabidopsis thaliana) that are involved in fucogalactoXyG biosynthesis (Fig. 1; Pauly et al., 2013; Schultink et al., 2014). The glucan backbone is thought to be synthesized by cellulose synthase-like C (CSLC) family proteins, such as AtCSLC4, as shown by in vitro activity data (Cocuron et al., 2007). Several xylosyltransferases (XXTs) from glycosyl transferase family 34 (GT34) are thought to be responsible for XyG xylosylation. Five of these XXTs in Arabidopsis seem to have XXT activity on XyG in vitro (Faik et al., 2002; Zabotina et al., 2008; Vuttipongchaikij et al., 2012; Mansoori et al., 2015). MURUS3 (MUR3) represents a galactosyltransferase that transfers galactosyl moieties specifically to xylosyl residues adjacent to an unsubstituted glucosyl residue on an XXXG unit, converting it to XXLG, whereas Xyloglucan L-side chain galactosyl Transferase2 (XLT2) was identified as another galactosyltransferase transferring a galactosyl moiety specifically to the second xylosyl residue, resulting in XLXG (Madson et al., 2003; Jensen et al., 2012). Both MUR3 and XLT2 belong to GT47 (Li et al., 2004). MUR2/FUCOSYLTRANSFERASE1 (FUT1) from GT37 was found to harbor fucosyltransferase activity, transferring Fuc from GDP-Fuc to a galactosyl residue adjacent to the unsubstituted glucosyl residue (i.e. onto XXLG but not onto XLXG; Perrin et al., 1999; Vanzin et al., 2002). O-acetylation of the galactosyl residue is mediated by Altered Xyloglucan4 (AXY4) and AXY4L, both of which belong to the Trichome Birefringence-Like (TBL) protein family (Bischoff et al., 2010; Gille et al., 2011; Gille and Pauly, 2012).Open in a separate windowFigure 1.Schematic structures of two types of XyGs and known biosynthetic proteins in Arabidopsis (Hsieh and Harris, 2009; Pauly et al., 2013). The corresponding one-letter code for XyG is shown below the pictograms (Fry et al., 1993; Tuomivaara et al., 2015).XyG found throughout land plants exhibits structural diversity with respect to side-chain substitution patterns (Schultink et al., 2014). Most dicots, such as Arabidopsis, and the noncommelinoid monocots possess a fucogalactoXyG of the XXXG-type XyG structure as shown in Figure 1. However, plant species in the Solanaceae and Poaceae as well as the moss Physcomitrella patens contain a different XyG structure with a reduced level of xylosylation, resulting in an XXGG_n core motif (York et al., 1996; Kato et al., 2004; Gibeaut et al., 2005; Jia et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009). In addition, the glucan backbone can be O-acetylated in plants of Solanaceae and Poaceae families (Gibeaut et al., 2005; Jia et al., 2005). XyG from Solanaceae with an XXGG core motif can be further arabinosylated and/or galactosylated (Jia et al., 2005). No XyGs with an XXGG_n motif backbone have been reported to be fucosylated.The function of structural diversity of XyG substitutions, such as fucosylation and/or altered xylosylation pattern, remains enigmatic. Removing the terminal fucosyl or acetyl moieties in the corresponding Arabidopsis mutants does not lead to any change in plant growth and development (Vanzin et al., 2002; Gille et al., 2011). However, removing galactosyl residues as well as fucosyl and acetyl moieties in the Arabidopsis xlt2 mur3.1 double mutant results in a dwarfed plant (Jensen et al., 2012; Kong et al., 2015). Replacing the galactosyl moiety with an arabinofuranosyl residue by, for example, expressing a tomato (Solanum lycopersicum) arabinosyltransferase in the Arabidopsis xlt2 mur3.1 mutant rescues the growth phenotype and restores wall biomechanics, indicating that galactosylation and arabinosylation in XyG have an equivalent function (Schultink et al., 2013). Recently, fucosylated XyG structures were found in the pollen tubes of tobacco (Nicotiana alata) and tomato, indicating that fucogalactoXyG is likely also present in other Solanaceae plants, albeit restricted to specific tissues (Lampugnani et al., 2013; Dardelle et al., 2015). Although there is circumstantial evidence that fucogalactoXyG is present in cell suspension cultures of rice (Oryza sativa) and cell suspension cultures of fescue (Festuca arundinaceae; McDougall and Fry, 1994; Peña et al., 2008), fucogalactoXyG has not been found in any physiologically relevant plant tissues of members of the Poaceae (Kato et al., 1982; Watanabe et al., 1984; Gibeaut et al., 2005; Hsieh and Harris, 2009; Brennan and Harris, 2011). Here, we provide chemical and genetic evidence that fucogalactoXyG is, indeed, present in plant tissues of a grass (rice) and prove that the rice genome harbors the genes that could be part of the synthetic machinery necessary to produce fucogalactoXyG.     

Hypoxia-cultured human adipose-derived mesenchymal stem cells are non-oncogenic and have enhanced viability,motility, and tropism to brain cancer

Adult human adipose-derived mesenchymal stem cells (hAMSCs) are multipotent cells, which are abundant, easily collected, and bypass the ethical concerns that plague embryonic stem cells. Their utility and accessibility have led to the rapid development of clinical investigations to explore their autologous and allogeneic cellular-based regenerative potential, tissue preservation capabilities, anti-inflammatory properties, and anticancer properties, among others. hAMSCs are typically cultured under ambient conditions with 21% oxygen. However, physiologically, hAMSCs exist in an environment of much lower oxygen tension. Furthermore, hAMSCs cultured in standard conditions have shown limited proliferative and migratory capabilities, as well as limited viability. This study investigated the effects hypoxic culture conditions have on primary intraoperatively derived hAMSCs. hAMSCs cultured under hypoxia (hAMSCs-H) remained multipotent, capable of differentiation into osteogenic, chondrogenic, and adipogenic lineages. In addition, hAMSCs-H grew faster and exhibited less cell death. Furthermore, hAMSCs-H had greater motility than normoxia-cultured hAMSCs and exhibited greater homing ability to glioblastoma (GBM) derived from brain tumor-initiating cells from our patients in vitro and in vivo. Importantly, hAMSCs-H did not transform into tumor-associated fibroblasts in vitro and were not tumorigenic in vivo. Rather, hAMSCs-H promoted the differentiation of brain cancer cells in vitro and in vivo. These findings suggest an alternative culturing technique that can enhance the function of hAMSCs, which may be necessary for their use in the treatment of various pathologies including stroke, myocardial infarction, amyotrophic lateral sclerosis, and GBM.Mesenchymal stem cells (MSCs) are multipotent cells, isolated from the bone marrow, adipose tissue, and muscle, among others. They are clonally expansive, with the capacity to differentiate into osteocytes, adipocytes, and chondrocytes.^{1, 2} MSCs are widely studied for their regenerative potential, tissue preservation capabilities, anti-inflammatory properties, and anticancer therapeutic potential.^{3, 4, 5} MSCs can serve as vehicles for delivering effective targeted therapy to primary brain cancer and metastatic cancer.^{6, 7, 8}Notwithstanding aggressive treatment of primary brain cancer (glioblastoma (GBM)) with surgical resection, chemotherapy, and radiotherapy, the median survival following diagnosis is 14.6 months.^{9, 10, 11, 12, 13, 14, 15} GBM-targeted therapy using neural stem cells and MSCs as vehicles for therapeutic agents is a promising strategy.¹⁶ MSCs seem to be the ideal stem cells, as they are autologous, easily collected, and easily re-implanted.^{17, 18} The most commonly used MSCs are bone marrow-derived MSCs (BM-MSCs) and human adipose-derived MSCs (hAMSCs). Compared with BM-MSCs, hAMSCs are easier to obtain.^{19, 20}Despite the potential utility of hAMSCs, their use is hampered by their low concentration within tissues.^{21, 22} Thus, in vitro expansion of hAMSCs is necessary. Compared with BM-MSCs, hAMSCs are more genetically and morphologically stable in long-term culture.^{19, 20} Nevertheless, current culturing conditions for both BM-MSCs and hAMSCs show a progressive decrease in viability and proliferative ability, and an increase in senescence ratio for these stem cells with time.^{23, 24, 25, 26, 27, 28, 29} Typically, hAMSCs are cultured under ambient conditions with 21% oxygen in vitro.³⁰ However, physiologically, hAMSCs exist at much lower oxygen tensions, between 1 and 14%.^{31, 32} As a result of the limitations of culturing under normoxia, we investigated the effect of hypoxia on intraoperatively obtained hAMSCs by assessing proliferation, survival, differentiation, tumor formation, tumor tropism, and migration in vitro and in vivo in a rodent model with a human brain cancer. hAMSCs have been reported to transform into tumor-associated fibroblasts (TAFs), which can potentially support tumor growth and promote malignant phenotypes.^{33, 34} Yet, no studies have reported on the changes that may occur in hypoxia-cultured hAMSCs after they are exposed to brain cancer, both in vitro and in vivo. An understanding of the effects of hypoxia on hAMSCs³⁵ is critical for its potential therapeutic applications including in the treatment of brain tumors, stroke, neuro-degenerative diseases such as multiple sclerosis, and dementia (Figure 1a).Open in a separate windowFigure 1Primary human adipose-derived cells cultured in hypoxia (hAMSCs-H) and normoxia (hAMSCs-N) are both MSCs but normoxia-cultured cells show increased signs of senescence, such as increased area and elongated morphology, compared with hypoxia-cultured cells. (a) hAMSCs were isolated from human fat tissue and cultured in hypoxic (1.5% oxygen) or normoxic (21% oxygen) conditions in vitro. The viability, mobility, tumor tropism, safety, and tumorigenic potential were subsequently compared in vitro and in vivo. (b) Differentiation assay. hAMSCs were cultured in control media and differentiation media for 3 weeks, 10 days after the second passage. Three different stains were performed to assess differentiation capabilities (scale bar, 100 μm). (c) Flow cytometric analysis was performed to confirm the absence of CD31-, CD34-, and CD45-positive cells in both cell cultures. In addition, primary hAMSC cultures expressed high levels of CD73, CD90, and CD105, both in hypoxic and normoxic culture conditions at day 10 after passage 2. (d) Representative images of cell morphologies of hAMSCs on 2D surface (scale bar, 200 μm). (e) Schematic of 3D-nanopatterned surface used to assess morphology and motility. (f) Images of cell morphologies of hAMSCs on 3D-nanopatterned surface (scale bar, 200 μm). (g–j) The length, width, area, and length-to-width ratio were measured and compared after cell aligned on the nanopattern surface. Error bars represent S.E.M. *P<0.05, **P<0.01, N.S., not significant     

p62/SQSTM1 upregulation constitutes a survival mechanism that occurs during granulocytic differentiation of acute myeloid leukemia cells

The p62/SQSTM1 adapter protein has an important role in the regulation of several key signaling pathways and helps transport ubiquitinated proteins to the autophagosomes and proteasome for degradation. Here, we investigate the regulation and roles of p62/SQSTM1 during acute myeloid leukemia (AML) cell maturation into granulocytes. Levels of p62/SQSTM1 mRNA and protein were both significantly increased during all-trans retinoic acid (ATRA)-induced differentiation of AML cells through a mechanism that depends on NF-κB activation. We show that this response constitutes a survival mechanism that prolongs the life span of mature AML cells and mitigates the effects of accumulation of aggregated proteins that occurs during granulocytic differentiation. Interestingly, ATRA-induced p62/SQSTM1 upregulation was impaired in maturation-resistant AML cells but was reactivated when differentiation was restored in these cells. Primary blast cells of AML patients and CD34⁺ progenitors exhibited significantly lower p62/SQSTM1 mRNA levels than did mature granulocytes from healthy donors. Our results demonstrate that p62/SQSTM1 expression is upregulated in mature compared with immature myeloid cells and reveal a pro-survival function of the NF-κB/SQSTM1 signaling axis during granulocytic differentiation of AML cells. These findings may help our understanding of neutrophil/granulocyte development and will guide the development of novel therapeutic strategies for refractory and relapsed AML patients with previous exposure to ATRA.p62 or sequestosome 1 (p62/SQSTM1) is a scaffold protein, implicated in a variety of biological processes including those that control cell death, inflammation, and metabolism.^{1, 2} Through its multi-domain structure, p62/SQSTM1 interacts specifically with key signaling proteins, including atypical PKC family members, NF-κB, and mTOR to control cellular responses.^{3, 4, 5, 6, 7} p62/SQSTM1 functions also as a key mediator of autophagy. Through its interaction with LC3, an essential protein involved in autophagy, p62/SQSTM1 selectively directs ubiquitinated substrates to autophagosomes leading to their subsequent degradation in lysosomes.^{8, 9} At the molecular level, p62/SQSTM1 acts as a pro-tumoral molecule by ensuring efficient and selective activation of cell signaling axes involved in cell survival, proliferation, and metabolism (i.e., NF-κB, mTOR, and Nrf-2 pathways).^{3, 5, 6, 7, 10, 11, 12, 13} p62/SQSTM1 can also signal anti-tumoral responses either by inactivating the pro-oncogenic signaling through BCR-ABL¹⁴ and Wnt pathways^{15, 16} or by inducing the activation of caspase 8, a pro-death protein.^{17, 18} Interestingly, in response to stress, autophagy promotes the degradation of p62, thus limits the activation of p62-regulatory pathways that control tumorigenesis.¹⁰ In addition, p62/SQSTM1 controls pathways that modulate differentiation of normal and cancerous cells. For example, p62/SQSTM1 has been shown to antagonize basal ERK activity and adipocyte differentiation.¹⁹ In contrast, p62/SQSTM1 favors differentiation of osteoclasts,²⁰ osteoblasts,²¹ neurons,²² megakaryocytes²³ and macrophages.²⁴ The role and regulation of p62/SQSTM1 during leukemia cell differentiation has been poorly documented.Acute myeloid leukemia (AML) is a hematological disease characterized by multiple deregulated pathways resulting in a blockade of myeloid precursors at different stages of maturation.^{25, 26} Acute promyelocyte leukemia (APL) is the M3 type of AML characterized by an arrest of the terminal differentiation of promyelocytes into granulocytes and frequently associated with the expression of the oncogenic PML-RAR alpha fusion gene.^{27, 28} All-trans retinoic acid (ATRA), a potent activator of cellular growth arrest, differentiation, and death of APL cells, has been shown to effectively promote complete clinical remission of APL when combined with chemotherapy.^{29, 30, 31} Despite the success of this treatment, some APL patients are refractory to ATRA treatment or relapse owing to the development of resistance to ATRA in leukemia cells.^{32, 33, 34}Our previous results revealed that autophagy flux is activated during granulocyte differentiation of myeloid leukemia cell lines induced by ATRA.³⁵ In the present study, we observed that p62/SQSTM1, an autophagic substrate, is markedly upregulated at both mRNA and protein levels during the granulocytic differentiation process. Here, we investigated the regulation and the function of p62/SQSTM1 during AML cells differentiation into neutrophils/granulocytes.     

Role of Retinoic Acid and Fibroblast Growth Factor 2 in Neural Differentiation from Cynomolgus Monkey (Macaca fascicularis) Embryonic Stem Cells

Retinoic acid is a widely used factor in both mouse and human embryonic stem cells. It suppresses differentiation to mesoderm and enhances differentiation to ectoderm. Fibroblast growth factor 2 (FGF2) is widely used to induce differentiation to neurons in mice, yet in primates, including humans, it maintains embryonic stem cells in the undifferentiated state. In this study, we established an FGF2 low-dose-dependent embryonic stem cell line from cynomolgus monkeys and then analyzed neural differentiation in cultures supplemented with retinoic acid and FGF2. When only retinoic acid was added to culture, neurons differentiated from FGF2 low-dose-dependent embryonic stem cells. When both retinoic acid and FGF2 were added, neurons and astrocytes differentiated from the same embryonic stem cell line. Thus, retinoic acid promotes the differentiation from embryonic stem cells to neuroectoderm. Although FGF2 seems to promote self-renewal in stem cells, its effects on the differentiation of stem cells are influenced by the presence or absence of supplemental retinoic acid.Abbreviations: EB, embryoid body; ES, embryonic stem; ESM, embryonic stem cell medium; FGF, fibroblast growth factor; GFAP, glial fibrillary acidic protein; LIF, leukemia inhibitory factor; MBP, myelin basic protein; RA, retinoic acid; SSEA, stage-specific embryonic antigen; TRA, tumor-related antigenPluripotent stem cells are potential sources of material for cell replacement therapy and are useful experimental tools for in vitro models of human disease and drug screening. Embryonic stem (ES) cells are capable of extensive proliferation and multilineage differentiation, and thus ES-derived cells are suitable for use in cell-replacement therapies.^18,23 Reported ES cell characteristics including tumorigenic potential, DNA methylation status, expression of imprinted genes, and chromatin structure were elucidated by using induced pluripotent stem cells.^2,11,17 Because the social expectations of regeneration medicine are growing, we must perform basic research with ES cells, which differ from induced pluripotent stem cells in terms of origin, differentiation ability, and epigenetic status.^2,8Several advances in research have been made by using mouse ES cells. Furthermore, primate ES cell lines have been established from rhesus monkeys (Macaca mulatta),²⁴ common marmosets (Callithrix jacchus),²⁵ cynomolgus monkeys (M. fascicularis),²⁰ and African green monkeys (Chlorocebus aethiops).¹⁹ Mouse and other mammalian ES cells differ markedly in their responses to the signaling pathways that support self-renewal.^8,28 Mouse ES cells require leukemia inhibitory factor (LIF)–STAT3 signaling.¹⁴ In contrast, primate ES cells do not respond to LIF. Fibroblast growth factor 2 (FGF2) appears to be the most upstream self-renewal factor in primate ES cells. FGF2 also exerts its effects through indirect mechanisms, such as the TGFβ–Activin–Nodal signaling pathway, in primate ES cells.²¹ In addition to the biologic similarities between monkeys and humans, ES cells derived from cynomolgus monkeys or human blastocysts have extensive similarities that are not apparent in mouse ES cells.^8,14,21,28 Numerous monkey ES cell lines are now available, and cynomolgus monkeys are an efficient model for developing strategies to investigate the efficacy of ES-cell–based medical treatments in humans.Several growth factors and chemical compounds, including retinoic acid (RA),^4,9,13,22,26 FGF2,^9,10,16,22 epidermal growth factor,^9,22 SB431542,^1,4,10 dorsomorphin,^10,27 sonic hedgehog,^{12,13,16,27,29} and noggin,^1,4,9,27 are essential for the differentiation and proliferation or maintenance of neural stem cells derived from primate ES cells. Of these factors, active RA signaling suppresses a mesodermal fate by inhibiting Wnt and Nodal signaling pathways during in vitro culture and leads to neuroectoderm differentiation in ES cells.^4,13,26 RA is an indispensable factor for the specialization to neural cells. FGF2 is important during nervous system development,¹² and FGF2 and RA both are believed to influence the differentiation to neural cells. The current study was done to clarify the mechanism of RA and FGF2 in the induction of differentiation along the neural lineage.We recently established a monkey ES cell line that does not need FGF2 supplementation for maintenance of the undifferentiated state. This ES cell line allowed us to study the role of differentiation to neural cells with RA and enabled us to compare ES cell differentiation in the context of supplementation with RA or FGF2 in culture. To this end, we established a novel cynomolgus monkey cell line derived from ES cells and maintained it in an undifferentiated state in the absence of FGF2 supplementation.     

Neuritin 1 promotes retinal ganglion cell survival and axonal regeneration following optic nerve crush

Neuritin 1 (Nrn1) is an extracellular glycophosphatidylinositol-linked protein that stimulates axonal plasticity, dendritic arborization and synapse maturation in the central nervous system (CNS). The purpose of this study was to evaluate the neuroprotective and axogenic properties of Nrn1 on axotomized retinal ganglion cells (RGCs) in vitro and on the in vivo optic nerve crush (ONC) mouse model. Axotomized cultured RGCs treated with recombinant hNRN1 significantly increased survival of RGCs by 21% (n=6–7, P<0.01) and neurite outgrowth in RGCs by 141% compared to controls (n=15, P<0.05). RGC transduction with AAV2-CAG–hNRN1 prior to ONC promoted RGC survival (450%, n=3–7, P<0.05) and significantly preserved RGC function by 70% until 28 days post crush (dpc) (n=6, P<0.05) compared with the control AAV2-CAG–green fluorescent protein transduction group. Significantly elevated levels of RGC marker, RNA binding protein with multiple splicing (Rbpms; 73%, n=5–8, P<0.001) and growth cone marker, growth-associated protein 43 (Gap43; 36%, n=3, P<0.01) were observed 28 dpc in the retinas of the treatment group compared with the control group. Significant increase in Gap43 (100%, n=5–6, P<0.05) expression was observed within the optic nerves of the AAV2–hNRN1 group compared to controls. In conclusion, Nrn1 exhibited neuroprotective, regenerative effects and preserved RGC function on axotomized RGCs in vitro and after axonal injury in vivo. Nrn1 is a potential therapeutic target for CNS neurodegenerative diseases.Central nervous system (CNS) trauma and neurodegenerative disorders trigger a cascade of intrinsic and extrinsic cellular events resulting in regenerative failure and subsequent damage to neurons.^{1, 2, 3, 4, 5} The intrinsic factors include deregulation in growth-promoting factors, apoptotic factors, intracellular signaling molecules and trophic factors.⁶ Similarly, the extrinsic factors correlate to growth inhibition due to inhibitory cues^{3, 7, 8, 9, 10, 11, 12, 13} that include myelin and myelin associated inhibitors, glial scarring,^{5, 14} slow clearance of axonal debris,⁷ incorrect development of neuronal projections⁶ and CNS inflammation.^{15, 16} Progressive degeneration of mature retinal ganglion cells (RGCs) has been associated with loss of trophic support,^{8, 9} detrimental inflammatory processes/immune regulation^{10, 11} and apoptotic effectors.^{9, 12, 13, 15, 17}After injury, mammalian RGC axons show only a short-lived sprouting response but no long-distance regeneration through the optic nerve (ON).¹⁶ Glial responses around the affected area are initiated by injured CNS axons.¹⁸ Axons undergoing Wallerian degeneration are surrounded by astrocytes that upregulate glial fibrillary acidic protein (Gfap) expression and these reactive astrocytes contribute to trauma-induced neurodegeneration.¹⁹ Glial scarring inhibits axonal transport after ON crush (ONC)^{5, 14} decreasing transport of proteins involved in neuroprotection and synaptic plasticity. Regenerative failure is a critical endpoint of these destructive triggers culminating in neuronal apoptosis^{3, 20, 21} and inhibition of functional recovery. Intrinsic factors affecting axonal regeneration after CNS injury are crucial for recovery and thus, dysregulation of genes involved in axonal plasticity and outgrowth can prove detrimental to the neuronal recovery.^{22, 23, 24}Current neuroprotection approaches include promoting survival of RGCs by intraocular injections of recombinant factors like ciliary neurotrophic factor (CNTF) and peripheral nerve (PN) transplantations in vitro²⁵ and in vivo after injury.²⁶ Studies performed with glial cell-line-derived neurotrophic factor and neurturin protect RGCs from axotomy-induced apoptosis.²⁷ Further, in the ON injury model, RGC survival was promoted after deletion of CCAAT/enhancer binding protein homologous protein²⁸ and enhanced regeneration observed with co-deletion of kruppel-like factor 4 (Klf4) and suppressor of cytokine signaling 3 (Socs3).²⁹ Intraocular administration of neurotrophin-4 (NT-4) and brain-derived neurotrophic factor (BDNF) after ON transection has also exerted neuroprotective effects on axotomized RGCs. In addition, PNs transplanted adjacent to ONs, ex vivo PN grafts with lenti-viral transduced Schwann cells, and stimulation of inflammatory processes have strong pro-regenerative effects on injured RGCs.^{26, 30, 31, 32, 33}In addition, using adeno-associated-virus (AAV) therapy, AAV mediated expression of CNTF in bcl2 overexpressing transgenic mice increases cell viability and axonal regeneration,³⁴ whereas BDNF promotes survival of RGCs.³⁵ Likewise, experiments with AAV–BDNF, –CNTF and –growth-associated protein 43 (GAP43) have shown that AAV–CNTF was the most crucial for promoting both long-term survival and regeneration.³⁶ The positive effects of CNTF are observed mainly through simultaneous deletion of both PTEN and SOCS3³⁷ and the concurrent activation of mTOR and STAT3 pathways.³⁸ Although CNTF shows robust increase and sustained axon regeneration in injured ONs of rodents, it causes axonal misguidance and aberrant growth.³⁹ Furthermore, it has been shown that CNTF acts as a chemoattractant. CNTF administration onto autologous PN grafts transplanted within transected ON increased regeneration, but these effects were significantly reduced after removal of macrophages from this site.⁴⁰ In addition, the effects of CNTF using PN grafts at ON transection sites are further subject to debate, as previously it has been shown that Ad-CNTF injections preserved RGC axons but did not induce regeneration of axotomized RGCs.⁴¹ Thus, other studies have addressed RGC survivability and axonal regeneration with CNTF and other growth factors,^{35, 36} but most trophic factors affect neuronal survival and regeneration differentially.Previous studies targeting neuronal apoptosis by overexpressing intrinsic growth factors, inhibiting apoptosis and enhancing regeneration in CNS trauma models have established that a multifactorial approach is required for successful and long-lasting therapeutic outcomes.^{6, 36} Current gaps still exist for a key gene that could effectively target neuroprotection, enhance neuron regeneration and sustain neuronal function.One key gene implicated in neuronal plasticity is Neuritin 1 (Nrn1), also known as candidate plasticity gene 15. It has multiple functions and was first identified and characterized when screening for candidate plasticity genes in the rat hippocampal dentate gyrus activated by kainate.^{42, 43, 44} Nrn1 is highly conserved across species⁴⁵ and translates to an extracellular, glycophosphatidylinositol-linked protein (GPI-linked protein), which can be secreted as a soluble form. Nrn1 stimulates axonal plasticity, dendritic arborization and synapse maturation in the CNS.⁴⁶ During early embryonic development, Nrn1 promotes the survival of neural progenitors and differentiated neurons,⁴⁷ while later in development it promotes axonal and dendritic growth and stabilization, allowing maturation and formation of synapses.^{43, 46, 48} In the adult brain, Nrn1 has been correlated with activity-dependent functional plasticity^{45, 49} and is expressed in post mitotic neurons.Nrn1 may be a crucial gene for neuroprotection and regeneration because growth factors such as nerve growth factor (NGF), BDNF and NT-3 as well as neuronal activity can potentiate the expression of Nrn1.^{44, 50} In addition, we reported that Nrn1 mRNA expression appears to be biphasic after ON axonal trauma, indicating a transient attempt by RGCs at neuroprotection/neuroregeneration in response to ONC injury.⁵¹ The dynamic regulation of Nrn1 coupled with neurotrophic effects may promote axonal regeneration in the CNS. To overcome CNS trauma, a new therapy geared towards neuroprotection and effective axonal regeneration is required to enhance a future multifactorial approach. The purpose of this study is to evaluate the therapeutic effects of Nrn1 in mouse RGC cultures as well as in the mouse ONC model. We have identified a distinct neuroprotective and regenerative strategy that prevents neurodegeneration after ON injury. AAV2–hNRN1 expression vectors partially rescued RGCs from apoptosis, maintained RGC function, and initiated regeneration of injured axons.     

Cell biology in development: Cell division and the maintenance of epithelial order

Epithelia are polarized layers of adherent cells that are the building blocks for organ and appendage structures throughout animals. To preserve tissue architecture and barrier function during both homeostasis and rapid growth, individual epithelial cells divide in a highly constrained manner. Building on decades of research focused on single cells, recent work is probing the mechanisms by which the dynamic process of mitosis is reconciled with the global maintenance of epithelial order during development. These studies reveal how symmetrically dividing cells both exploit and conform to tissue organization to orient their mitotic spindles during division and establish new adhesive junctions during cytokinesis.The association of large numbers of cells in tightly organized epithelial layers is a unique and defining feature of Metazoa. Although classical studies of development once labeled distinct embryonic regions as territories, fields, layers, placodes, and primordia, we now know many of these structures to be primarily constructed from epithelial sheets. Epithelial structure and function are critically dependent on cell polarization, which is coupled to the targeted assembly of adhesive junctions along the apicolateral membranes of adjacent cells (Tepass et al., 2001; Cavey and Lecuit, 2009). In brief, the plasma membrane of epithelial cells is polarized into apical and basolateral domains, each enriched with distinct lipid and protein components (Fig. 1; Rodriguez-Boulan et al., 2005; St Johnston and Ahringer, 2010). At the molecular level, E-cadherins are the major class of adhesion proteins that establish cell–cell connections through homophilic interaction across cell membranes (Takeichi, 1991, 2011; Halbleib and Nelson, 2006; Harris and Tepass, 2010). Whereas E-cadherin is apically enriched in invertebrate epithelia, it is localized along the lateral domain of vertebrate epithelial cells. In both cases, E-cadherin interacts with cytoplasmic actin filaments via the catenin class of adaptor proteins, thus coupling intercellular adhesive contacts to the cytoskeleton (Cavey and Lecuit, 2009; Harris and Tepass, 2010; Gomez et al., 2011). Within this framework, the maintenance of both polarity and cell–cell adhesion are essential for epithelial barrier function and tissue architecture during growth and morphogenesis (Papusheva and Heisenberg, 2010; Guillot and Lecuit, 2013b).Open in a separate windowFigure 1.Architectural implications of orthogonal and planar spindle orientations during epithelial cell division. (A) Programmed orthogonal orientation of the mitotic spindle can promote epithelial stratification, although the remodeling of adhesion and polarity complexes during this process remains an important area for further study. (B) Planar spindle orientation is coordinated with the overall cell polarity machinery and thus facilitates conservation of monolayer organization during rapid cell proliferation.During development, epithelia expand by the combined effects of cell growth (increase in cell size) and cell division (increase in cell numbers). Division events are typically oriented either parallel or orthogonal to the plane of the layer and less frequently at oblique angles (Gillies and Cabernard, 2011). When cells divide orthogonally (perpendicular to the plane of the epithelium), the two daughters will be at least initially nonequivalent with respect to position within the cell layer (Fig. 1 A). Under normal conditions, such programmed orthogonal divisions can be used to effect asymmetric segregation of cell fates or to establish distinct cell types, such as in the developing cortex (Fietz et al., 2010; Hansen et al., 2010) or during morphogenesis of stratified epithelia (Lechler and Fuchs, 2005; Williams et al., 2011). Conversely, when cells divide parallel to the plane of the epithelium (planar orientation; Fig. 1 B), both daughter cells are equivalent with respect to mother cell polarity and tightly integrated in the growing monolayer (Morin and Bellaïche, 2011).During planar division, epithelial cells typically round up, constrict in the middle to form the cytokinetic furrow, and divide symmetrically with respect to the apicobasal axis to produce two equal daughter cells. These daughters construct new cell–cell junctions at their nascent interface, thus integrating into the monolayer (Fig. 2, A–G). Although the intricate relationship between cell polarity and cell division has been explored for many years in the context of asymmetric cell division (Rhyu and Knoblich, 1995; Siller and Doe, 2009; Williams and Fuchs, 2013), recent studies have also begun to explore how epithelia maintain their morphology, integrity, and barrier function during continuous rounds of planar cell division and junction assembly. In this review, we highlight recent findings that provide new insights into the problem of symmetric planar cell division in diverse polarized epithelia, with a focus on two crucial mitotic events: (1) the orientation of cell division and (2) the formation of new cell junctions.Open in a separate windowFigure 2.Progression of planar cell division in an epithelial monolayer. Apical cross section (xy, top row) and longitudinal (xz, bottom row) view of a dividing cell (red). (A) At the level of apical junctions, cells are packed in a polygonal cell arrangement during interphase. (B) In prophase, the dividing nucleus begins to translocate apically as the cell rounds up and maintains a thin basal projection enriched with nonmuscle myosin II and actin (light blue). Notably, this type of nuclear migration is typically observed in pseudostratified columnar epithelia and does not occur in cuboidal and squamous epithelial tissues. (C) Localized molecular landmarks (apical complexes marked as gray bars on cell sides) direct orientation of the mitotic spindle to the plane of the epithelium (arrows). (D) Within the plane of the cell layer, the spindle can be further oriented (arrows) in response to molecular cues, global tissue tension, and local cell geometry. (E and F) After chromosome segregation during anaphase, the cell constricts in the middle and cleaves orthogonal to the plane of the monolayer. (G) After cytokinesis, daughter nuclei move basally and daughter cells form new junctions at their nascent interface (white) while elongating along the apicobasal axis.

Mitotic spindle position and orientation in epithelial cells

Planar orientation of epithelial cell division requires coordinated interaction between the cell polarization machinery and the mitotic spindle itself (Morin and Bellaïche, 2011). In animal cells, the spindle is organized by two symmetrically positioned poles or centrosomes, which nucleate three forms of microtubules (Tanaka, 2010): kinetochore microtubules that attach to the chromosomes, polar microtubules that overlap in an antiparallel fashion over the midplane, and astral microtubules that extend to the cell cortex, which is the actin-rich layer beneath the cell membrane (Lancaster and Baum, 2014). Work in Drosophila melanogaster and vertebrates reveals that at least three factors influence the orientation of this spindle machinery with respect to polarized epithelial architecture: cytoskeletal forces, localized cortical cues, and tissue tension.

Cytoskeletal forces position mitotic nuclei near the apical cell membrane.

In columnar and pseudostratified epithelia where cells elongate along their apicobasal axes, mitotic events are typically restricted to the apical domain of the epithelium (corresponding to the apical membrane of each cell; Fig. 2, C–F). How does the mitotic nucleus achieve the correct apical position? In Drosophila wing discs and zebrafish neuroepithelia, mitotic nuclei and the bulk of the cell cytoplasm are driven apically by actomyosin-dependent cortical contractility at prophase entry (Norden et al., 2009; Leung et al., 2011; Meyer et al., 2011). These events are fundamentally similar to mitotic cell rounding in tissue culture cells (Kunda and Baum, 2009; Lancaster et al., 2013; Lancaster and Baum, 2014). In many epithelia, as the cell rounds up and the nucleus translocates apically, a thin actin-rich projection maintains contact with the basal lamina (Fig. 2, B and C). It remains poorly understood how this structure behaves during cleavage and whether this basal process plays any role in the correct reintegration of the postmitotic daughter cells into the monolayer. Although actomyosin may be the primary driver of apical rounding in many cases, evidence also supports a role for microtubule-based mechanisms in the positioning of premitotic nuclei. In chicken neural tube and mouse cerebral cortex, nuclei migrate apically on microtubules before actomyosin-dependent rounding (Spear and Erickson, 2012a). Centrosomes provide directionality to the microtubules on which the nucleus migrates and organize the spindle once the mitotic chromatin reaches the apical domain (Peyre et al., 2011; Spear and Erickson, 2012a; Nakajima et al., 2013). Collectively, current evidence suggests that both actomyosin- and microtubule-dependent forces conspire to effect mitotic nuclear translocation in a highly context- and species-specific manner. One possibility is that the varying physical dimensions of epithelial cells require varying mechanisms for apical nuclear translocation. For example, highly elongated radial glial cells require active transport of the nucleus on microtubules before mitotic rounding, whereas cortical actomyosin contractility may be sufficient in less elongated cells (Spear and Erickson, 2012b). A major outstanding problem is how cortical contractility triggers cell rounding that is polarized along the apicobasal axis of the cell. Whereas centrosomes function as an apical landmark for nuclei moving on microtubules, it remains unclear what provides the directionality for the basal-to-apical actomyosin contraction. One hypothesis is that certain proteins can restrict the localization of nonmuscle myosin II at the basal domain of epithelial cells. The microcephaly protein Asp interacts with myosin II and regulates its polarized localization along the apicobasal axis in the fly optic lobe neuroepithelium. In asp mutant flies, myosin II is enriched apically instead of basally. Many dividing nuclei fail to reach the apical domain and are thus broadly distributed along the apicobasal axis of the epithelium, leading to a disorganized tissue (Rujano et al., 2013). Interestingly, Asp also interacts with microtubules, associates with spindle poles, and is essential for positioning the spindle in fly and vertebrate epithelia (Saunders et al., 1997; do Carmo Avides and Glover, 1999; Wakefield et al., 2001; Fish et al., 2006). Elucidating the function of proteins such as Asp at the interface of microtubules and actomyosin will be essential to our understanding of how the cytoskeleton drives apical mitotic rounding.

Localized molecular landmarks direct planar spindle orientation.

In most animal cells, the mitotic spindle is anchored to the cell cortex by astral microtubules (Fig. 2, C–E; Théry and Bornens, 2006). Translocation of the dynein–dynactin motor toward the astral microtubule minus ends provides a pulling force on centrosomes and is essential for spindle orientation and pole separation during cell division (Dujardin and Vallee, 2002; Kotak et al., 2012). Molecular cues embedded in the cortex can thus determine spindle orientation by anchoring the dynein–dynactin complex in restricted domains. In cultured MDCK and chick neuroepithelia cells, the Gαi–LGN–nuclear mitotic apparatus (NuMA) complex serves this function (Busson et al., 1998; Hao et al., 2010; Zheng et al., 2010; Peyre et al., 2011). Knockdown or mislocalization of these factors leads to spindle orientation defects that ultimately lead to removal of cell progenitors from the monolayer (Peyre et al., 2011). LGN (Pins in Drosophila) localizes to the lateral cell cortex by binding to the membrane-bound Gαi and enforces spindle orientation by recruiting NuMA (Mud in Drosophila), which binds directly to the dynein–dynactin motor. In certain epithelia, including MDCK cells and Drosophila wing discs, LGN is excluded from the apical domain by atypical PKC (aPKC) phosphorylation, thus restricting it at the lateral cell cortex (Konno et al., 2008; Hao et al., 2010; Zheng et al., 2010; Guilgur et al., 2012). In chick neuroepithelia, however, LGN is restricted at the lateral cortex independently of aPKC, suggesting that other cues control its localization (Morin et al., 2007; Peyre et al., 2011). In the mouse embryonic neocortex, the actin–membrane linkers ERM (ezrin/radixin/moesin) promote the association of LGN with NuMA (Machicoane et al., 2014), indicating that organized cortical actin is critical for correct LGN localization.Cell–cell junctions have been implicated in planar cell division in mammalian epithelia, suggesting a possible direct link between the polarity apparatus and the spindle machinery (Reinsch and Karsenti, 1994; den Elzen et al., 2009). Interfering with E-cadherin function or reducing E-cadherin levels abolishes junctional localization of APC (adenomatous polyposis coli), a microtubule-interacting protein that is required for planar spindle orientation and chromosome alignment (Green et al., 2005; den Elzen et al., 2009). However, spindle orientation may not directly depend on E-cadherin or adherens junctions (AJs) in all cases. In Drosophila follicle cells and imaginal discs as well as Xenopus laevis embryonic epithelia, mitotic spindles exhibit planar orientation but do not align with the AJs (Woolner and Papalopulu, 2012; Bergstralh et al., 2013; Nakajima et al., 2013). Moreover, disruption of AJs in Drosophila follicle cells does not affect spindle position (Bergstralh et al., 2013).In Drosophila wing discs, the spindle poles localize in close proximity to septate junctions, which are positioned immediately basal to AJs (Nakajima et al., 2013). Septate junctions are enriched with many proteins, including the neoplastic tumor suppressors SCRIB (Scribbled) and DLG1 (Discs large 1; Bilder and Perrimon, 2000; Bilder et al., 2000). In asymmetrically dividing cells, such as Drosophila sensory organ precursors and neuroblasts, DLG1 interacts with LGN at the cortex and is required for proper spindle orientation (Bellaïche et al., 2001; Siegrist and Doe, 2005; Johnston et al., 2009). Recent findings indicate that DLG1 is also essential for planar spindle orientation in the symmetric division of epithelial cells. In wing discs, knockdown of scrib or dlg1 leads to randomized spindle orientations. scrib knockdown wing discs exhibit diffuse DLG1 localization but no obvious apicobasal polarity defect, suggesting that epithelial disorganization could be a consequence of aberrant spindle orientation (Nakajima et al., 2013). However, it is not clear whether the septate junctions themselves are important. In Drosophila follicle epithelial cells where septate junctions do not form until relatively late in development (Oshima and Fehon, 2011), DLG1 is localized at the lateral cell cortex and is essential for planar spindle orientation (Bergstralh et al., 2013). Interestingly, dlg1 mutant follicle cells display misoriented divisions yet normal epithelial polarity and tissue organization. In this case, planar spindle orientation appears to be independent of junctions per se but still depends on a DLG1–LGN–NuMA complex, similar to asymmetrically dividing cells (Bergstralh et al., 2013).

Global stress and local cell geometry influence mitotic spindle orientation within the plane of the epithelium.

During planar divisions, the mitotic spindle aligns to the plane of the epithelium (xz; Fig. 2 C) and also within the plane of the cell layer (xy; Fig. 2 D). Studies in gastrulating zebrafish embryos revealed a role for the Wnt–Frizzled–planar cell polarity signaling pathway in orienting cell divisions (Concha and Adams, 1998; Gong et al., 2004). Similarly, the atypical cadherins Fat and Dachsous are involved in orienting cell divisions in the Drosophila wing and in developing mouse kidneys (Baena-López et al., 2005; Saburi et al., 2008). Although both of these pathways have been reviewed elsewhere (Morin and Bellaïche, 2011), recent studies also point to at least two other mechanisms that may independently influence spindle orientation within the plane of the monolayer: (1) global tissue stress and (2) local epithelial cell geometry.Epithelial cell shape and spindle orientation are modulated by global stress that accumulates during tissue growth. In Drosophila wing discs, cells in the center of the wing blade primordium proliferate at a faster rate than in the periphery. Consequently, cells in the periphery are mechanically stretched, and cells in the center are compressed. As a result of stretching, peripheral cells localize myosin II at their cortex and align their mitotic spindle with the stretch axis (LeGoff et al., 2013; Mao et al., 2013). Similarly, epithelial cells of the enveloping cell layer in gastrulating zebrafish embryos elongate and orient their spindle along the direction of tension generated by spreading during epiboly (Campinho et al., 2013). It is unclear whether myosin II directly conveys cell tension to the mitotic apparatus, and it will be necessary to dissect whether cell elongation alone or additional mechanosensing pathways signal cell tension to the mitotic spindle. Keratinocytes from the mammalian epidermis reorient their mitotic spindle in response to mechanical stretch in a NuMA-dependent manner. The mitotic spindle aligns with the cortical NuMA-localized crescent upon stretch and fails to orient when NuMA levels are reduced (Seldin et al., 2013). In summary, global tension generated by growth and cell spreading impact division orientation, suggesting that shape changes in proximity to dividing cells may also lead to a similar effect.Although variations certainly exist, the apical surfaces of proliferating epithelia tend to feature a consistent percentage of hexagonal, pentagonal, heptagonal, and octagonal cell shapes (Gibson et al., 2006; Farhadifar et al., 2007; Aegerter-Wilmsen et al., 2010). In Drosophila imaginal discs, these local patterns of cell packing may systematically influence spindle orientation, as mitotic cells are biased toward cleaving their common interfaces with subhexagonal neighbors (less than six sides) and avoid cleaving their interfaces with superhexagonal neighbors (more than six sides; Gibson et al., 2011). Although the mechanisms underlying the effect of local cell geometry remain elusive, cell packing influences mitotic cell shape and the distribution of adhesive cues, both of which could, in turn, bias spindle orientation. Indeed, dividing cells maintain contacts with their neighbors, which can influence the cell cortex and direct spindle orientation (Goldstein, 1995; Wang et al., 1997). The distribution of adhesions between epithelial cells may also alter the position or action of cortical force generators that interact with spindle microtubules in the mitotic cell. In support of this idea, when single cells are placed on micropatterned substrates, they orient their spindle relative to the geometry of their adhesion pattern and not their cell shape (Théry et al., 2005, 2007). Alternatively, neighbors of different polygonal shapes could stretch the mitotic cell, thus imposing a bias on its long axis. Indeed, sea urchin embryos orient their spindles to divide their longest axis (Hertwig, 1884) and can even sense complex cell geometries to orient their spindles accordingly (Minc et al., 2011). Still, precisely how the interphase morphology of epithelial cells might impinge on mitotic spindle orientation remains an open question.

Genesis of nascent junctions during epithelial cell division

After spindle orientation, the essential processes of cytokinesis and abscission are driven by the assembly and contraction of an actomyosin ring positioned in the cleavage plane (Fededa and Gerlich, 2012). In epithelia, ring contraction accompanied by membrane invagination ultimately gives rise to a new junctional interface between nascent daughter cells. Precisely how this new interface forms remains poorly understood. Recent studies in Drosophila epithelia reveal that, during cytokinesis, (a) E-cadherin levels are reduced at the interface between the cleavage furrow of dividing cells and their neighbors (Fig. 3), and (b) neighbor tension and midbody position guide establishment of new AJs in context with local epithelial geometry (Fig. 4).Open in a separate windowFigure 3.Cytokinetic membrane dynamics in epithelial cells. (A) Cytokinesis of a dividing epithelial cell (yellow) presents several unique structural considerations not addressed by the analysis of single cells. Recent studies (Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013, 2014) report a local reduction of E-cadherin levels in proximity to the contractile ring in the dividing cell and its neighbor (red). How cytokinesis is resolved from there may vary in a context-dependent manner. (B) In Drosophila embryos, ring contraction leads to E-cadherin disengagement, and a gap forms between the mitotic cell and its neighbor (Guillot and Lecuit, 2013a). (C) In the Drosophila pupal notum, the contractile ring pulls the neighbor cell plasma membrane into the cleavage furrow, perhaps enabled by uncoupling of the membrane and the cortex in the neighbor (Herszterg et al., 2013, 2014).Open in a separate windowFigure 4.New AJ formation in dividing epithelial cells. Apical cross section (xy, top row) and longitudinal (xz, bottom row) view of a dividing epithelial cell (red). (A) Opposing forces (black vertical arrows) develop between the contractile ring in the dividing cell and the two neighboring cells (orange) in proximity to the cleavage furrow. E-cadherin clusters are reduced at the furrow/neighbor interface. (B) Myosin II and tension build up in the neighboring cell, causing the nascent daughter cells to juxtapose their plasma membranes at the presumptive site of junction assembly (black horizontal arrows). (C) Arp2/3 and Rac1 drive actin polymerization at the daughter cell interface around the midbody, stabilizing the nascent junction as the neighboring cell membrane withdraws. (D) The new junction is complete and of suitable length in context with the local epithelial geometry.

Mitotic cells remodel their adhesion junctions during cytokinesis.

Two kinds of forces are at work during cytokinesis: an active force in the dividing cell caused by ring contraction and a reactive force in contacting neighbors caused by their resistance to pulling to maintain their shape (Fig. 3 and Fig. 4 A; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013). Recent results indicate that these opposing forces can lead to a transient and partial reduction of cell adhesion after mitotic exit. In Drosophila epithelia, E-cadherin levels are reduced at the interface between the cleavage furrow of the dividing cell and its neighbors (Fig. 3, B and C; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013; Morais-de-Sá and Sunkel, 2013). Specifically in embryonic epithelia, the local reduction of E-cadherin facilitates membrane separation, and a gap appears between the dividing cell and its neighbors (Fig. 3 B; Guillot and Lecuit, 2013a). In the dorsal thorax, in contrast, the neighbor cell plasma membrane detaches from the cortex and is drawn into the cleavage furrow (Fig. 3 C; Herszterg et al., 2013, 2014). What triggers E-cadherin modulation in cells after mitotic exit? The loss of overall cell polarity is one possible mechanism. During mitosis in Drosophila, follicular epithelial cells lose cortical enrichment of some apical polarity proteins (aPKC, Crumbs, and Bazooka/Par3; Bergstralh et al., 2013; Morais-de-Sá and Sunkel, 2013), and embryonic cells lose localization of lethal giant larvae, a basolateral cortical protein (Huang et al., 2009). Contrasting with these observations, however, MDCK cells and Drosophila embryonic and dorsal thorax epithelial cells appear to maintain apicobasal polarity as they divide (Reinsch and Karsenti, 1994; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013). Furthermore, E-cadherin reduction is limited to the furrow/neighbor interface and is not observed in other areas of cell contact. Therefore, an alternative mechanism that explains local E-cadherin modulation is mechanical tension that arises precisely at the area between the contractile ring and the neighboring cell membrane (Founounou et al., 2013; Guillot and Lecuit, 2013a).Does E-cadherin modulation serve a functional role in mitotic cells? In Drosophila embryonic and dorsal thorax epithelia, E-cadherin decrease leads to a local adhesion disengagement proposed to facilitate the formation of new AJs between daughter cells (Founounou et al., 2013; Guillot and Lecuit, 2013a). It has been previously reported that cells maintain their AJs throughout division. For example, intercellular junctions are maintained in dividing cells of human colonic mucosal crypt cells and basal keratinocytes (Baker and Garrod, 1993). Similarly, mitotic MDCK cultured cells maintain tight junctions apically and E-cadherin basolaterally (Reinsch and Karsenti, 1994). The E-cadherin loss in certain Drosophila epithelia may be either a tissue-specific phenomenon or a highly dynamic process only observable with the temporal resolution of live-cell imaging. Moreover, dividing cells in the Drosophila dorsal thorax show decreased levels of E-cadherin yet maintain their cohesiveness (Herszterg et al., 2013). Interestingly, E-cadherin is internalized in mitotic MDCK cells (Bauer et al., 1998). It will therefore be important to investigate whether loss of E-cadherin leads to adhesion disengagement in other epithelial tissues and whether tension alone or in combination with biochemical pathways is responsible for E-cadherin modulation.

Epithelial neighbors exert tension on daughter cell membranes to facilitate new AJ formation.

How new junctional contacts form during mitosis is a poorly understood problem at the heart of epithelial cell biology. In Drosophila, new membrane interfaces between nascent daughter cells initially show only a weak level of E-cadherin clusters (Guillot and Lecuit, 2013a; Herszterg et al., 2013). Subsequently, the daughter cells assemble their AJs de novo. How is the length of these new junctions determined with respect to cell geometry? Recent evidence indicates that AJ length is a function of local cell packing within the epithelium. In dividing cells of the Drosophila dorsal thorax, the contractile ring triggers tension and accumulation of myosin II in neighbors at the furrow/neighbor interface (Fig. 4 B; Founounou et al., 2013; Herszterg et al., 2013). Myosin II in the neighboring cells in turn contracts and creates tension at the furrowing membrane of the nascent daughter cells, keeping them tightly pressed against each other (Fig. 4, B and C). This local membrane juxtaposition facilitates AJ formation. To allow expansion of the daughter cell interface and maintain AJ length, branched actin polymerization via Rac1 and Arp2/3 is oriented to the midbody, which serves as a positional landmark for new AJs (Fig. 4 C; Herszterg et al., 2013). The midbody is a narrow intercellular bridge that remains after the contracted cytokinetic ring has driven membrane invagination, and it recruits the abscission factors that will eventually separate the daughter cells (Fededa and Gerlich, 2012). Interestingly, the midbody is positioned apically as a result of the presence of AJs. In Drosophila follicular epithelia, the midbody also provides cues for the formation of the apical daughter cell interface, suggesting that it plays a role in both AJ and epithelial cell polarity establishment and maintenance in dividing epithelial cells (Morais-de-Sá and Sunkel, 2013). Thus, examples from Drosophila epithelia show that cohesion between dividing cells and their neighbors together with the apically positioned midbody provides a spatial template and polarized positional cue for de novo AJ assembly (Herszterg et al., 2013; Morais-de-Sá and Sunkel, 2013). Further work on other epithelial tissues may provide alternative mechanisms of junction biogenesis.

Growth and order in the epithelium: Thinking outside the cell

During development, epithelial monolayers have the remarkable capacity to maintain specialized morphologies and barrier functions during rapid cell proliferation. Mitotic cells remain adherent to their neighbors throughout cell division. Cell cohesion enables local geometry and global tissue tension to instruct mitotic cells where to position their cleavage plane and how to assemble their junctions. However, local tension may also lead to a transient disengagement of dividing cells from their neighbors after mitotic exit. How is global and local tension conveyed to protein complexes in mitotic cells so that different outcomes take place? Moreover, it is unclear whether and how tissue tension instructs synchronously dividing epithelial cells how to divide and reestablish their junctions after division. Clearly, this is a fundamental problem for the maintenance of epithelial order and may be linked to the origin of epithelial cancers, in which cells undergo rapid proliferation but fail to remain integrated into the monolayer.The selected studies discussed here hint at the remarkable level of coordination that occurs during epithelial cell division, recasting mitosis as a truly multicellular process. Looking ahead, understanding the interface between cells, proteins, and mechanical forces that each operate on different scales will require creative multidisciplinary approaches in diverse organismal systems. Indeed, epithelial organization is widespread in nature and is encountered among even the most basal animals, including sponges and cnidarians as well as the fruiting body of the nonmetazoan social amoeba Dictyostelium discoideum (Wood, 1959; Ereskovsky et al., 2009; Houliston et al., 2010; Dickinson et al., 2011; Meyer et al., 2011). Combined, future interdisciplinary studies and a fresh look at diverse animal models should yield new insight into epithelial cell division for many years to come.     

The tumor-modulatory effects of Caspase-2 and Pidd1 do not require the scaffold protein Raidd

The receptor-interacting protein-associated ICH-1/CED-3 homologous protein with a death domain (RAIDD/CRADD) functions as a dual adaptor and is a constituent of different multi-protein complexes implicated in the regulation of inflammation and cell death. Within the PIDDosome complex, RAIDD connects the cell death-related protease, Caspase-2, with the p53-induced protein with a death domain 1 (PIDD1). As such, RAIDD has been implicated in DNA-damage-induced apoptosis as well as in tumorigenesis. As loss of Caspase-2 leads to an acceleration of tumor onset in the Eμ-Myc mouse lymphoma model, whereas loss of Pidd1 actually delays onset of this disease, we set out to interrogate the role of Raidd in cancer in more detail. Our data obtained analyzing Eμ-Myc/Raidd^−/− mice indicate that Raidd is unable to protect from c-Myc-driven lymphomagenesis. Similarly, we failed to observe a modulatory effect of Raidd deficiency on DNA-damage-driven cancer. The role of Caspase-2 as a tumor suppressor and that of Pidd1 as a tumor promoter can therefore be uncoupled from their ability to interact with the Raidd scaffold, pointing toward the existence of alternative signaling modules engaging these two proteins in this context.A number of mechanisms have evolved to trace and remove potentially dangerous cells. Deregulation of the induction of apoptosis upon oncogenic stress, for example, can facilitate the accumulation of cells prone to undergo malignant transformation. Cell death by apoptosis depends on the cascade-like activation of proteases of the Caspase family.¹ Among these, the evolutionarily most conserved protease, Caspase-2, turns out to be a potent tumor suppressor in mice^{2, 3, 4, 5, 6, 7} and correlative expression data support a conserved role in human cancer.^{8, 9, 10, 11, 12, 13}Early studies suggested that Caspase-2 interacts with other proteins for its activation (e.g., after genotoxic stress), but the protease seems also able to auto-activate cell death on its own when present in sufficiently high concentration.^{14, 15, 16, 17, 18} The most prominent Caspase-2-containing protein complex was dubbed the ‘PIDDosome'' and described to contain the p53-induced protein with a death domain (PIDD1) and receptor-interacting protein-associated ICH-1/CED-3 homologous protein with a death domain (RAIDD, also known as CRADD).¹⁹ Although the molecular details of the pro-apoptotic potential of Caspase-2 are still discussed and alternative roles in the DNA-damage response, cell cycle arrest or sensor of metabolic stress are mechanistically poorly understood, Caspase-2 clearly limits tumorigenesis in different settings. These include aberrant expression of c-Myc in B cells^{3, 4} or deletion of the DNA-damage response regulator, ataxia telangiectasia mutated kinase (ATM), both driving lymphomagenesis⁶ as well as overexpression of the Her2/ErbB2 oncogene in breast⁵ or that of mutated KRAS in the lung epithelium, driving carcinoma formation.⁷ One of these studies, addressing also the role of Pidd1 in c-Myc-driven lymphomagenesis, revealed an unexpected oncogenic role for Pidd1, thereby questioning the physiological relevance of the PIDDosome complex in Caspase-2-mediated cell death and tumor suppression.^{4, 20} However, the exact role of the scaffold protein Raidd within these processes remains unaddressed so far.Raidd, a bipartite adapter containing a death domain (DD) and a caspase-recruitment domain (CARD) was first described to bind to the DD-containing kinase RIPK1 and the C. elegans caspase CED-3,²¹ supporting a role in cell death initiation. Subsequently, the interaction of Caspase-2 and Raidd was biochemically proven²² and proposed to be required for Caspase-2 autoprocessing preceding its activation.¹⁹ More recent studies propose an anti-inflammatory role for Raidd through suppression of nuclear factor kappa-light-chain enhancer (NF-κB) activation and cytokine production upon T-cell receptor stimulation by negatively interfering with the Carma1/Malt1/Bcl-10 signaling complex.^{23, 24}First evidence for a potential role of RAIDD in human cancer was discovered in a biochemical screen using mantle cell lymphomas, which detected a downregulation of RAIDD by microarray analysis,¹⁰ whereas others reported on RAIDD-linked multidrug resistance in osteosarcoma cells.²⁵ Furthermore, tumor cell apoptosis induced by inhibitors of histone de-acetylases in treatment-resistant adult T-cell leukemia lines reportedly required Caspase-2 and Raidd.²⁶ It is also reported that the Caspase-2/Raidd axis is necessary after ER stress, for example, in the course of infection with the oncolytic maraba virus.²⁷Taken together, these studies support a role for RAIDD in drug-induced cancer cell death as well as in tumor suppression, most likely linked to its role as a direct activator of Caspase-2. Alternatively, RAIDD may negatively interfere with PIDD- or BCL10-regulated NF-κB signaling^{23, 24, 28} and thereby suppress pro-tumorigenic inflammation. To address the role of Raidd in tumorigenesis in more detail, we exploited different mouse models where we induced thymic lymphomas by γ-irradiation, fibrosarcomas by 3-methylcholanthrene (3-MC) injection or B-cell lymphomas by aberrant expression of the c-Myc proto-oncogene. Our results suggest that Raidd is not a suppressor of tumors in the mouse models tested.