Tissue patterning during development requires that cells interact with each other by means of adhesion complexes to orient cell polarity, organize the cytoarchitecture and to coordinate and adapt cell behavior (Mateus et al.,2009). Once polarity is established, cells are compartmentalized to generate distinct apical and basal regions that carry out specialized functions, a process dependent upon cytoskeletal re-organization (Salas et al.,1997; Quinlan and Hyatt,1999; Ameen et al.,2001; Kim and Coulombe,2007). This allows progenitors and more developmentally restricted cell populations to respond in a coordinated manner to external signals (e.g., secreted, extracellular matrix (ECM) or mechanical signals) such that differentiation is regulated and normal morphogenesis can occur.
During placental development, villi undergo extensive branching morphogenesis to increase the surface area for the exchange of nutrients and wastes necessary to support fetal growth and survival (Cross et al.,2006). In mice, the villi of the labyrinth layer contain three trophoblast cell types: the mononuclear sinusoidal trophoblast giant cells (S-TGCs) that surround the maternal blood spaces, and two types of syncytiotrophoblast (SynT) cells, one of which (SynT-II) directly abuts the fetal vascular endothelial cells while the other (SynT-I) is sandwiched between the SynT-II cells and S-TGCs (reviewed by Watson and Cross,2005). Recently, genetic marker analysis has given us clues regarding the developmental origins of these labyrinth trophoblast cell types suggesting that they arise from distinct precursor populations in the chorion before branching morphogenesis begins (Simmons et al.,2008). Multipotent trophoblast stem (TS) cells are present in the extraembryonic ectoderm of the chorion (Uy et al.,2002) and become developmentally restricted after embryonic day (E) 7.5, as indicated by the decrease in ability to derive TS cell cultures from this tissue (Uy et al.,2002) and the developmental restriction of TS cell-specific genes Cdx2, Esrrb, and Eomes (Beck et al.,1995; Pettersson et al.,1996; Luo et al.,1997; Hancock et al.,1999). By the time of chorioallantoic attachment at embryonic day (E) 8.5, the chorion has become patterned and divided into at least three distinct cell layers, each of which is hypothesized to contain precursor populations that correspond to the three differentiated cell types within the mature labyrinth (Simmons et al.,2008). Briefly, cells in the apical-most layer of the chorion are likely the precursors to S-TGCs, as they reside in direct contact with developing maternal sinusoids and express the Hand1 gene; the cell layer situated immediately below expresses Syna and is anticipated to give rise to SynT-I cells, whereas the basal-most layer of the chorion contains clusters of cells that express the genes Gcm1, Synb and Cebpa, which mark branchpoint initiation sites and give rise to SynT-II cells (Simmons et al.,2008). Of interest, the basal layer of the chorion also contains distinct clusters of proliferating Rhox4b+/Gcm1− cells suggesting a population of less restricted labyrinth progenitor cells since Rhox4b expression colocalizes with the TS cell markers Esrrb and Cdx2 at early postimplantation stages and persists in cells with similar morphology even after Esrrb and Cdx2 expression declines (A. Davies, E. Maríusdóttir, D.N., D.S., J.C., in preparation). How these labyrinth trophoblast precursor populations are patterned and regulated is not well understood.
We previously showed that the Mrj co-chaperone (also known as Dnajb6) is necessary for chorioallantoic attachment (Hunter et al.,1999), a key developmental milestone required for mouse placenta development. In mice, Mrj is widely expressed in adult and embryonic tissue, although its expression in the trophoblast lineage of the placenta is necessary for embryo survival past E10.5 (Hunter et al.,1999; Seki et al.,1999; Chuang et al.,2002). Mrj is a member of the Hsp40 family of co-chaperones, which convey substrate specificity to the chaperone Hsp70 in a tissue dependent manner (Chuang et al.,2002; Fan et al.,2003). Recently, a large number of Mrj-substrate interactions have been identified suggesting a wide range of cytoplasmic and nuclear functions for Mrj (Izawa et al.,2000; Chuang et al.,2002; Dai et al.,2005; Hurst et al.,2006; Cheng et al.,2008; Li et al.,2008; Mitra et al.,2008; Pan et al.,2008; Zhang et al.,2008; Bhowmick et al.,2009; Dey et al.,2009; Durrenberger et al.,2009; Edo De Bock et al.,2010; Mitra et al.,2010). In the context of placenta development, the interaction between Mrj and its substrate keratin (K) 18 is particularly important (Izawa et al.,2000; Watson et al.,2007). Mrj deficiency inhibits proteasome degradation of K18-containing filaments leading to the collapse of the keratin cytoskeleton into large aggregates within the chorion trophoblast cells and TGCs. These keratin aggregates inhibit normal cell function because removing them by generating K18/Mrj double mutants restores allantoic attachment in Mrj−/− conceptuses (Watson et al.,2007). However, the mechanism by which the aggregates disrupt chorion trophoblast cell function has not yet been determined. We show here that Mrj−/− trophoblast cells lack normal cell behavior and have altered cell–cell and cell–matrix interactions. These phenotypes are associated with a profound effect of Mrj deficiency on patterning of the chorion and syncytiotrophoblast progenitor differentiation.
Disorganization of Mrj−/− Chorion Trophoblast Cells Is Associated With Erratic TS Cell Behaviour In Vitro
We previously reported that the formation of keratin aggregates in Mrj−/− trophoblast cells was associated with fewer desmosome adhesion complexes and reduced desmoplakin expression (Watson et al.,2007) suggesting that the adhesive properties of Mrj−/− cells were altered. Accordingly, we re-examined the morphology of Mrj−/− chorions at E8.25, just prior to chorioallantoic attachment, using electron microscopy. As was previously observed (Watson et al.,2007), instead of a columnar arrangement of trophoblast cells forming organized layers as in wild-type chorions (Fig. 1A,C), the trophoblast cells of Mrj−/− chorions were amorphous and disorganized (Fig. 1B,D) indicating that chorion trophoblast cell adhesion was altered. Furthermore, trophoblast stem (TS) cell cultures, an established model of studying chorionic trophoblast cells in vitro (Tanaka et al.,1998; Uy et al.,2002), recapitulated this phenotype as Mrj−/− TS cells were rounded and less adherent to each other (Fig. 1E,F) as indicated by the increased level of light refraction between each cell compared to wild-type.
To understand how Mrj deficiency affected trophoblast cell behavior, we used time-lapse video microscopy on TS cell cultures over the span of 120 min. Upon attaching to the culture plate, wild-type TS cells remained clustered together (Fig. 1G). After approximately 80 min, some cells sent out lamellipodia-like projections but continued to be tightly adherent to the rest of the cell colony, with little to no migration (Fig. 1G and Supp. Data Movie File 1) (Supp. movies are available online). The behavior of Mrj−/− TS cells was dramatically different. Immediately after Mrj−/− cells contacted the culture plate, they formed lamellipodia and migrated away from each other, though keeping in contact with neighboring cells via long, thin projections (Fig. 1H). The paths of these highly migratory cells were random and often converged, but instead of adhering upon contact, they moved in a repulsive manner in opposite directions (Fig. 1H and Supp. Data Movie File 2). Cytoplasmic “trails” were also frequently observed in the wake of Mrj-deficient cells (Fig. 1H), a phenomenon not observed in wild-type cultures. Most of these “trails” were retracted back into the cells over time (Supp. Data Movie File 2). The unstable and inadequate cell-cell interactions along with the dynamic cell behavior of Mrj-deficient trophoblast cells suggests that, without Mrj, proper communication between cells may be hindered.
Mrj−/− Trophoblast Stem Cells Remain in a Progenitor State
To test the developmental potential of Mrj−/− chorion trophoblast cells, we compared the proliferation and differentiation capabilities of two independently-derived Mrj−/− TS cell lines to wild-type TS cells. Even when cultured in the absence of fibroblast growth factor 4 (FGF4) for 5 days, both Mrj−/− cell lines continued to proliferate at the same rate as mutant cells cultured with FGF4 and revealed a four to eightfold increase in total cell number compared to wild-type cultures (P < 0.0001) (Fig. 2A). Furthermore, morphological assessment of cells after FGF4 withdrawal showed that the vast majority of wild-type TS cells differentiated into TGCs or syncytiotrophoblast over the span of 5 days (Fig. 2B,C), as determined by the presence of cells with expansive cytoplasm and large or multiple nuclei. By comparison, significantly fewer Mrj−/− TS cells appeared to differentiate (P < 0.0001) (Fig. 2B,C). It is possible that the intrinsic cytoskeletal defect caused by Mrj deficiency (Watson et al.,2007; see below) prevented the stereotypical cytoplasmic spreading that occurs during normal differentiation resulting in atypically small and rounded cells (Fig. 2B). A few morphologically normal cells were present in the mutant cultures in differentiation conditions (Fig. 2B,C) likely due to the fact that Mrj is not expressed in all TGCs (Watson et al.,2007).
To further assess the capacity of Mrj−/− TS cells to differentiate into specific trophoblast cell types, we performed northern blot analysis on RNA collected from wild-type and Mrj−/− TS cells cultured in the presence of FGF4 (day 0) and 2, 4, and 6 days after FGF4 was withdrawn. No differences between wild-type and Mrj−/− TS cells were apparent in the expression patterns of early TS cell markers Esrrb and Cdx2 (Fig. 2D). This suggested that multipotent cells were present in Mrj−/− TS cell cultures and that, similar to wild-type, these cells were able to differentiate when growth factors were removed. Although by morphology they did not appear to differentiate as frequently, Mrj−/− TS cells expressed the genes Prl3d1 (also known as Pl1) and Tpbpa, markers of parietal TGCs and ectoplacental cone/spongiotrophoblast cells, respectively, both before and after the withdrawal of FGF4. Wild-type TS cells, on the other hand, only expressed Prl3d1 and Tpbpa after 6 days of differentiation (Fig. 2D) indicating that Mrj deficiency may cause the premature differentiation of some TS cells into TGCs and ectoplacental cone/spongiotrophoblast cells.
Trophoblast subtypes of the placental labyrinth layer were distinctly affected by Mrj deficiency. Even in differentiating conditions, the syncytiotrophoblast progenitor population was maintained in Mrj−/− TS cultures as revealed by a significant up-regulation of Rhox4b expression (Fig. 2D). Correspondingly, Gcm1, Synb and Syna expression was undetectable or markedly reduced in Mrj−/− TS cells even after withdrawal of FGF4 (Fig. 2D) suggesting that fewer mature syncytiotrophoblast cells were able to form in the absence of Mrj. Despite this, higher levels of Ctsq and Hand1 mRNA were detected indicating that more Mrj-deficient TS cells differentiated into S-TGCs compared to wild-type TS cells (Fig. 2D). Together, these data suggested that Mrj-deficient TS cells have a reduced ability to differentiate into syncytiotrophoblast cells. Instead, these cells remain as syncytiotrophoblast progenitors or, alternatively, may differentiate into ectoplacental cone/spongiotrophoblast cells, S-TGCs or parietal TGCs.
Aggregation of Actin and Its Associated Cell Adhesion Molecules in Mrj−/− TS Cells
Normally, the keratin cytoskeleton forms a filamentous network throughout the cytoplasm, incorporating with the actin cytoskeleton (Kim and Coulombe,2007). Therefore, we hypothesized that the presence of keratin aggregates observed in Mrj−/− trophoblast cells (Watson et al.,2007) may adversely affect the actin cytoskeleton and associated cell adhesion molecules. Using rhodamine-conjugated phalloidin to visualize F-actin, we found that the filaments in wild-type TS cells were located at the cell periphery (Fig. 3A,B). In contrast, the majority of actin filaments in Mrj−/− TS cells formed large perinuclear inclusions (Fig. 3C,D), which colocalized with the K18-containing aggregates (data not shown). Actin filaments attach to the cell membrane at adherens junctions, protein complexes that contain β-catenin and E-cadherin and are involved in cell adhesion in a Ca2+-dependent manner (Pokutta and Weis,2002). In most wild-type TS cells, β-catenin (Fig. 3E,F) and E-cadherin (Fig. 3I,J) expression was located at the cell membrane. Independent cells not in contact with other cells had low to undetectable expression of both proteins (data not shown). By contrast, both β-catenin (Fig. 3G,H) and E-cadherin (Fig. 3K,L) appeared up-regulated in Mrj−/− TS cells including unusually robust expression in cells without cell contacts. However, in both cases, these proteins were not restricted to the membrane and localized to large aggregates (Fig. 3G,H,K,L). Together, these data imply that the keratin aggregates resulting from Mrj deficiency may cause the actin cytoskeleton collapse providing insufficient means to establish and/or stabilize adhesion complexes.
E-cadherin Loss-of-Function Is Not the Primary Cause of the Mrj Differentiation Defect
The nonadherent phenotype and the large proportion of aggregated E-cadherin protein observed in Mrj−/− TS cells grown with (Fig. 3L) or without (Fig. 3M) FGF4 demonstrated that inadequate levels of functional E-cadherin protein may be present on the cell membrane. As a result, we hypothesized that the differentiation phenotype of Mrj−/− TS cell might be attributed to the loss of adherens junctions. To test this, wild-type TS cells were treated with small interfering RNA (siRNA) against Cdh1, the gene that encodes for E-cadherin, to down-regulate E-cadherin expression. Treatment of TS cells with a control GFP-labeled nonsense siRNA indicated that the transfection efficiency was between 17.9 and 62.0% (data not shown). Cdh1 siRNA-treatment of TS cells cultured in the presence of FGF4 did not recapitulate the morphological phenotype of Mrj−/− TS cells as they remained adherent and similar in shape compared to control TS cells treated with nonsense siRNA or no siRNA (Fig. 3N). However, while the control cells became typically large with sizeable nuclei, expansive cytoplasm and indistinguishable cell borders upon FGF4 withdrawal, the majority of Cdh1 siRNA-treated cells remained small, were rounded and lacked normal adherent behavior, reminiscent of Mrj-deficiency (Fig. 3N).
Of interest, knockdown of E-cadherin expression did not affect the capacity of TS cells to differentiate into labyrinth trophoblast cell types, based on gene expression analysis as assessed by quantitative reverse transcription-polymerase chain reaction (qRT-PCR). Cdh1 knockdown was validated in siRNA-treated TS cells cultured with FGF4 (9.7-fold decrease; P < 0.001) or without FGF4 (1.25-fold decrease; P < 0.01) (Fig. 3O). In the presence of FGF4, Cdh1 knockdown resulted in an up-regulation of Esrrb mRNA by 2.2-fold (P < 0.001) and a down-regulation of Rhox4b and Gcm1 mRNA expression by 2.5-fold (P < 0.001) compared with control nonsense siRNA-treated cells (Fig. 3O). Alternatively, Cdh1 siRNA-treated TS cells cultured in the absence of FGF4 for 5 days resulted in a down-regulation of Rhox4b (1.2-fold; P < 0.05) and an up-regulation of Gcm1 (2.1-fold; P < 0.01) and Syna (1.5-fold; P < 0.001) compared to control cells (Fig. 3O), indicating that E-cadherin is not required for the differentiation of TS cells into syncytiotrophoblast cells and, in fact, in the absence of FGF4, its down-regulation may promote syncytiotrophoblast differentiation. Collectively, these data imply that the failure of Mrj−/− TS cells to differentiate into syncytiotrophoblast cells is not primarily caused by the misexpression of E-cadherin.
Extracellular Matrix Organization Is Disrupted in Mrj-deficient Chorions
Alterations in ECM deposition and organization can also influence tissue organization and differentiation (Xu et al.,2009). Therefore, we examined electron micrographs of the ECM between the trophoblast and mesothelial layers of the chorion in wild-type and Mrj−/− littermate conceptuses at E8.25. Although an ECM was detected on the basal membrane of the chorionic trophoblast cell layer in Mrj mutants (Fig. 4C,D), it appeared to be thicker and less organized compared to wild-type (Fig. 4A,B).
Laminin is a key component of the trophoblast basement membrane (Klaffky et al.,2001). In particular, the laminin isoform α5 (Ln α5; contributes to laminin-511) is normally expressed between the chorion trophoblast and mesothelial cell layers from E6.5 to E8.5 (Miner et al.,1998; Klaffky et al.,2001; Alpy et al.,2005; Aumailley et al.,2005). To address whether the expanded ECM layer observed in Mrj−/− chorions was due to changes in laminin-511 deposition, we immunostained Mrj−/− trophoblast cells in vivo and in vitro with an antibody against Ln α5. Compared to wild-type chorions at E8.5 (Fig. 4E,F), the distribution of Ln α5 expression appeared thicker in Mrj−/− chorions and it was unclear whether the protein was properly presented on the cell membrane of the trophoblast cells (Fig. 4G,H). Mrj−/− TS cells cultured in the presence of FGF4 appeared more intensely stained for Ln α5 than wild-type cells (Fig. 4I–L), although western blot analysis did not show an obvious difference in expression levels (Fig. 4M). The apparent discrepancy between the qualitative (immunofluorescence) and quantitative (Western blot) Ln α5 observations may be the result of differences in protein localization between wild-type and Mrj−/− TS cells, suggesting that Ln α5 protein may not be confined to the basal membrane of the mutant cells. Furthermore, the western blot analysis revealed a failure of Mrj−/− TS cells to down-regulate Ln α5 following FGF4 withdrawal (Fig. 4M). These data suggest that the adverse effects of Mrj deficiency may arise from the mislocalization and/or the disorganization of the ECM at the trophoblast cell surface.
Exogenous Laminin-511 Normalizes Behaviour But Not Gene Expression of Mrj−/− TS Cells
To elucidate whether a defective ECM was responsible for the erratic cell behavior and limited developmental potential caused by Mrj deficiency, we provided wild-type and Mrj−/− TS cells with exogenous laminin-511. As previously reported (Klaffky et al.,2006), laminin-511 caused a general increase in wild-type TS cell circumference and the formation of large lamellipodia-like projections at their leading edges (Fig. 5A,B). Of interest, we observed that culturing Mrj-deficient cells on exogenous laminin-511 (Fig. 5D) resulted in a morphology that was more similar to wild-type TS cells on laminin-511 (Fig. 5B) than to Mrj−/− TS cells without a substrate (Figs. 1F, 5C). Even though Mrj−/− TS cells plated either without a substrate or on laminin-511 had a large cell circumference and frequently formed lamellipodia-like projections, exogenous laminin-511 enabled Mrj-deficient TS cells to form attachments to neighboring cells (Fig. 5D), which rarely occurred when cultured on plastic (Figs. 1H, 5C). Immunostaining with rhodamine-conjugated phalloidin (Fig. 5E,F,K,L) or antibodies against β-catenin (Fig. 5G,H,M,N) and E-cadherin (Fig. 5I,J,O,P) reinforced this observation and implied that adherens junctions in Mrj-deficient TS cells may be more stable in the presence of exogenous laminin-511 though some actin, β-catenin and E-cadherin aggregates were still present in these cells. Using time-lapse imaging, we observed further that providing exogenous laminin-511 to Mrj−/− TS cells reduced the amount of migration (Fig. 5D, data not shown) compared to cells on plastic (Fig. 1H).
To determine whether exogenous laminin-511 normalized the developmental potential and differentiation of Mrj−/− TS cells, we performed gene expression analysis. Despite stabilizing the cell behavior phenotype, northern blot analysis revealed that exogenous laminin-511 did not normalize the gene expression profiles in Mrj−/− TS cells in vitro (Fig. 5Q). Therefore, abnormal ECM organization is not likely the primary cause of the differentiation defect observed in Mrj−/− trophoblast cells in vitro.
Mrj-Deficient Chorions Lack the Establishment of Early Patterning
To address whether the developmental organization of Mrj−/− chorion trophoblast cells was disrupted in vivo, we analyzed the expression of several chorion layer-specific genes at E8.25. At this time, the chorion is divided into distinct layers of trophoblast cells, all of which express Mrj (Fig. 6A,B). Normally, the basal trophoblast layer of the chorion (closest to the site of allantoic attachment) contains clusters of proliferating Rhox4b-expressing cells (Fig. 6C,D), which are hypothesized to be the progenitors of syncytiotrophoblast cells (A. Davies, E. Maríusdóttir, D.N., D.S., and J.C., in preparation). These cell clusters are interspersed with branchpoint initiation sites containing nonproliferating, Rhox4b-negative cells that express the genes Gcm1, Cebpα, and Synb (Fig. 6G–I) and will eventually differentiate into SynT-II cells (Simmons et al.,2008). The apical layer of the chorion, on the other hand, contains precursors of SynT-I cells (Simmons et al.,2008) and express genes such as Syna and Snai1 (Fig. 6M,N). Remarkably, Rhox4b expression was not restricted in Mrj−/− chorions but instead was detected throughout the entire basal layer of the chorion with expansion into the layers above (Fig. 6E,F). Gcm1, Cebpα, and Synb gene expression was down-regulated or undetectable in the mutant chorions (Fig. 6J–L) suggesting that sites of branchpoint initiation were either not established or maintained in Mrj−/− chorions. Syna and Snai1 expression was also undetectable in Mrj−/− chorions (Fig. 6P,Q). Taken together, these data suggest that Mrj deficiency impedes the specification of both types of syncytiotrophoblast cells resulting in cells that are maintained in an undifferentiated, progenitor state. Interestingly, the uppermost layer of the chorion, which is also considered to be the base of the ectoplacental cone and is hypothesized to give rise to S-TGCs of the mature labyrinth (Simmons et al.,2008), appeared to be normal in Mrj−/− chorions as Hand1, a marker for these cells, was similarly expressed in wild-type and Mrj−/− chorions at E8.25 (Fig. 6O,R). This indicates that Mrj deficiency does not perturb all patterning within the chorion and that specification of S-TGCs may be cell-autonomous.
Patterning of tissue primordia requires that cells interact via adhesion complexes and the basement membrane to orient polarity, coordinate cell behavior and organize the cytoskeleton (Mateus et al.,2009). The establishment of cell polarity results in compartmentalization of cellular components such that distinct apical and basal regions of the cell are able to carry out separate functions. We have shown here that Mrj-deficient chorion trophoblast cells have altered cell behavior and, in addition to keratin inclusion bodies previously identified (Watson et al.,2007), mutant cells also show actin cytoskeleton instability and a lack normal cell–cell and cell–ECM interactions. These finding are associated with a failure to correctly pattern the chorion given that the cells are maintained in a Rhox4b+ progenitor-state and do not differentiate.
Mrj Deficiency Disrupts Both Keratin and Actin Cytoskeleton As Well As Cell–Cell Adhesion
Both the keratin and actin cytoskeletons are required to establish and maintain cell polarity (Salas et al.,1997; Quinlan and Hyatt,1999; Ameen et al.,2001). The Mrj co-chaperone was previously shown to directly interact with K18-containing filaments (Izawa et al.,2000), promoting their degradation through the proteasome machinery (Watson et al.,2007). Actin, β-catenin and E-cadherin are not among the known substrates for Mrj. However, Mrj may only need to interact with one of these proteins for all three to be affected since actin disorganization can affect adherens junction stability and vice versa (Peifer,1993; Quinlan and Hyatt,1999). On the other hand, keratin and actin networks are structurally interconnected (Kim and Coulombe,2007) and therefore, the effects of Mrj deficiency on actin architecture are likely secondary to the collapse and aggregation of the keratin cytoskeleton. This is supported by the fact that genetic removal of keratin aggregates from Mrj−/− chorions, as in Mrj−/−;K18−/− double mutant conceptuses, rescues the chorioallantoic attachment defect in Mrj mutants (Watson et al.,2007).
Theoretically, the formation of actin, β-catenin and E-cadherin aggregates in Mrj−/− trophoblast cells may be a consequence of keratin inclusions acting as a sink for other proteins. Precedence for this occurs in cirrhosis of the liver where proteins such as β-catenin, 14-3-3ζ and ZO-1 have been shown to adhere to keratin aggregates in diseased hepatocytes (Hanada et al.,2007). We have observed colocalization of actin and keratin aggregates in Mrj−/− TGCs in vitro (E.W. and J.C., unpublished). Alternatively, un-degraded keratin may overload the proteasome, preventing it from degrading other proteins, thus causing their aggregation. For example, amyloid-containing aggregates in Alzheimer's disease have been shown to overload the proteasome and affect its normal function (Upadhya and Hegde,2007). Regardless of the mechanism, aggregate formation likely interferes with the localization of key proteins required for the establishment of adhesion and cell–cell interactions. In the end, it is possible that this may prevent the developmental organization of Mrj−/− chorions.
Trophoblast Cell Differentiation and Patterning of the Chorion Is Affected by Mrj Deficiency
The major developmental consequence of Mrj deficiency is that trophoblast cells in the chorion fail to differentiate and establish the normal pattern of distinct cell layers. We have found that Mrj−/− TS cells are able to initiate differentiation by down-regulating the expression of genes associated with multipotency (Err2, Cdx2), but they continue to proliferate and lack the ability to progress beyond the Rhox4b+ stage to form more restricted syncytiotrophoblast precursors. The expanded Rhox4b expression domain in Mrj−/− chorions occurs at the expense of both the Gcm1/Synb/Cebpa-positive and the Syna-positive layers implying that these cells are maintained in a progenitor state. To date, we only have a few insights into how patterning in the chorion is established. It is important to note that neither the induction nor the maintenance of Syna and Hand1 expression patterns is dependent upon the presence of Gcm1/Synb/Cebpa+ cells because both are properly specified in Gcm1−/− and Cebpa−/−;Cebpb−/− mutant placentas (Simmons et al.,2008). Similarly, Syna−/− labyrinths are able to form SynT-II cells that express Gcm1 and Synb (Dupressoir et al.,2009). Furthermore, the fact that Hand1+ cells are appropriately positioned at the top layer of Mrj−/− chorions implies that S-TGC precursors are able to form independently of the syncytiotrophoblast progenitor cells and that Hand1+ cells cannot induce Rhox4b+ cells to form syncytiotrophoblast cells. As a result, signaling between different layers of the chorion is not necessary for their induction yet nonautonomous signaling within each cell layer may be required. To fully understand these interactions, it will be important to assess other mutants with chorion trophoblast defects for the establishment of chorionic patterning (e.g., Grb2 hypomorph, CtBP1−/−; CtBP2−/−, CyclinF−/− and Wnt7b−/−) (Parr et al.,2001; Saxton et al.,2001; Hildebrand and Soriano,2002; Tetzlaff et al.,2004).
Mrj mutants show both defects in chorioallantoic attachment and chorion patterning but whether there is a relationship between these two developmental events is not well studied. It is likely, though, that these processes are independent. For instance, Gcm1 expression is detected as early as E7.5 (Basyuk et al.,1999), one full day before chorioallantoic attachment (Downs,2002) suggesting that patterning of the chorion occurs independently of allantoic contact (Cross et al.,2006). In addition, Gcm1−/− and Syna−/− chorions undergo chorioallantoic attachment (Anson-Cartwright et al.,2000; Dupressoir et al.,2009) suggesting that neither the Gcm1/Synb+ nor the Syna+ layer of the chorion needs to be patterned for the allantois to attach. However, it is possible that the establishment of proper cell–cell interactions within the chorion supersedes these two developmental processes because Mrj−/− chorions, which have cytoskeleton and cell adhesion defects, are associated with phenotypes affecting chorioallantoic attachment (Hunter et al.,1999; Watson et al.,2007) and patterning (this study).
Potential Mechanisms Underlying the Patterning Defects in Mrj Mutants
Our studies have revealed several defects in Mrj−/− trophoblast cells that shed some light on the mechanism(s) underlying the patterning defect in the chorion. The most obvious change is that both the ECM and cell–cell interactions are abnormal in Mrj mutant trophoblast cells. We detected changes in ECM organization and found that providing Mrj-deficient TS cells with exogenous laminin-511 normalized their migratory behavior and adhesive stability. Laminin-511, itself, is unnecessary for chorioallantoic attachment or patterning of the chorion since Lama5−/− conceptuses undergo successful allantoic attachment and subsequent branching morphogenesis (Miner et al.,1998). Of interest, the exposure of TS cells to distinct substrates (e.g., laminin-511, laminin-111, or matrigel) results in behavioral differences (Klaffky et al.,2006; Lei et al.,2007). For example, TS cells plated on exogenous laminin-111 are rounded with decreased cell–cell contacts and are more migratory compared to those plated on laminin-511 or without a substrate (Klaffky et al.,2006). We have observed that the persistent migration of Mrj−/− TS cells, their inability to adhere to one another and their abnormal gene expression profiles were retained even in the presence of laminin-111 (E.W., M.H., A.S., and J.C., data not shown) suggesting that Mrj−/− cells are responsive to some substrates but not others. Importantly, while cell behavior was improved by providing exogenous laminin-511 to Mrj−/− TS cells, their developmental potential remained altered as revealed by gene expression. It may be that the localization of receptors that organize the basement membrane or bind the cell to it to ultimately change gene expression may be affected by Mrj deficiency (e.g., dystroglycan or α6β1 integrin, respectively; Cohn et al.,2002; Olsen and Ffrench-Constant,2005). With respect to normal cell–cell adhesion, chorion trophoblast cells and cultured TS cells are tightly adherent to each other through cell adhesion complexes that include E-cadherin. It is known that human syncytiotrophoblast progenitors (cytotrophoblast cells) express E-cadherin, which is down-regulated upon their differentiation into syncytiotrophoblast cells (Brown et al.,2005; James et al.,2005). In our studies, however, down-regulation of E-cadherin expression in cultured TS cells did not modify gene expression in ways similar to that observed in Mrj mutants. Collectively these results imply that while cell–cell and ECM interactions are abnormal in Mrj mutants, these defects on their own do not appear to explain the differentiation and chorion patterning defects.
A second general mechanism by which Mrj may function in chorion trophoblast cells is related to cell signaling. Apart from its involvement in cell adhesion, β-catenin also regulates tissue patterning and differentiation via the Wnt signaling pathway (reviewed by Miller and McCrea, 2010). It was recently shown that exogenous expression of MRJ in human carcinoma cells induced proteasome degradation of β-catenin through the up-regulation of DKK1, a secreted inhibitor of Wnt signaling (Mitra et al.,2010). We observed that Mrj deficiency may lead to an up-regulation of nuclear β-catenin expression in TS cells (Figs. 3, 5), though confocal analysis is necessary to understand the extent to which this occurs. Presumably, an increase in β-catenin nuclear translocation would initiate Tcf/Lef-1 transcriptional activation of genes downstream of Wnt signaling. Whether the up-regulation of genes in Mrj−/− TS cells, such as Rhox4b or Hand1, is a direct consequence of increased nuclear β-catenin or whether other factors caused by Mrj deficiency affect gene expression is yet to be determined. Similar to Mrj mutants, knockout mice for specific genes involved in the Wnt pathway (e.g., Wnt7b−/−, LPP3−/−, and Tcf−/−; Lef1−/−) do not undergo chorioallantoic attachment (Galceran et al.,1999; Parr et al.,2001; Escalante-Alcalde et al.,2003) but further work is needed to assess whether chorion patterning is also affected.
A third possible mechanism for Mrj function relates to its role in cell division. Altered cell polarity associated with keratin and actin cytoskeleton aggregates in Mrj−/− chorion trophoblast cells may affect asymmetric stem cell division, a mechanism required by some progenitors for the asymmetric partitioning of components that determine cell fate or the asymmetric placement of daughter cells with respect to extrinsic cues (Morrison and Kimble,2006). The choice between symmetric division (producing two progenitor cells) versus asymmetric division (producing one progenitor and one differentiated cell) is dependent upon the positioning of the microtubule organizing centre (Morrison and Kimble,2006), the location of which has been shown to rely on keratin filaments (Oriolo et al.,2007). Therefore, the collapse of the keratin cytoskeleton in Mrj−/− trophoblast cells may alter placement of the microtubule organizing centre resulting in symmetric rather than asymmetric partitioning of cell fate factors. Ectopic expression of MRJ in human carcinoma cells causes the cells to polarize and become more epithelial-like (Mitra et al.,2010). Furthermore, actin reorganization during trophoblast differentiation in vitro is correlated with changes in small Rho GTPase activity (Parast et al.,2001), which, in turn, can alter gene expression (Coso et al.,1995; Hill et al.,1995; Minden et al.,1995; Perona et al.,1997). Aside from a potential role in asymmetric division, Mrj interacts with other substrates that promote cell cycle arrest (Pei,1999; Hurst et al.,2006; Li et al.,2008; Mitra et al.,2008; Zhang et al.,2008; Dey et al.,2009) or the maintenance of progenitor populations (Watson et al.,2009) in other contexts. Therefore, it is possible that Mrj acts in concert with one or more of these cell cycle regulators to control whether Rhox4b+ progenitors continue to proliferate or whether these cells undergo cell cycle arrest to start down the path of differentiation.
Mrj+/− mice were originally generated by 6AD1 β-geo gene-trap insertion and maintained in a CD-1 genetic background (Hunter et al.,1999). Mrj−/− conceptuses were derived from Mrj+/− intercrosses. Pregnant females were dissected at E8.25 or E8.5 (noon of the day that the vaginal plug was detected was considered E0.5). PCR genotyping was performed using DNA from yolk sac tissue as previously described (Watson et al.,2007). Experiments were performed in accordance with the Canadian Council on Animal Care and the University of Calgary Committee on Animal Care (Protocol No. M06045).
E8.25 implantation sites from Mrj+/− matings were fixed, dehydrated, resin embedded and sectioned as previously described (Watson et al.,2007). Yolk sacs were removed for genotyping prior to fixation. Electron micrographs were taken on a Hitachi 7000 transmission electron microscope.
Wild-type (line 6-3) and Mrj−/− (lines 1-4 and 4-1) TS cell lines were derived according to previous descriptions (Watson et al.,2007). These lines and the Rosa26 TS cell line (kindly provided by Dr J. Rossant, Hospital for Sick Children, Toronto, Canada) (Tanaka et al.,1998) were maintained in 25 ng/ml basic fibroblast growth factor (FGF4), 1 μg/ml heparin in TS cell medium, 70% of which was preconditioned by embryonic fibroblasts (CM), at 37°C under 95% humidity and 5% CO2. Cells were differentiated by withdrawing FGF4, heparin and CM from the media (Tanaka et al.,1998). For laminin-511 substrate (EMD Biosciences) coating of culture dishes, 1.25 ml of a 10 μg/mL solution in PBS without calcium and magnesium-CMF-PBS was used and blocked with 1% BSA (Sigma) as previously described (Klaffky et al.,2006).
TS cells were cultured on coverslips in the presence of growth factors for 4 days, fixed in 4% PFA/1× PBS for 10 min, treated with 0.1% Triton X-100 (Fisher Biotech) in 1× PBS for 10 min and blocked in 1% BSA/1× PBS for 30 min. Primary antibody dilutions were made in 5% host serum, 1% BSA in 1× PBS, and 1-hr incubations at room temperature (RT) were used. Primary antibodies and dilutions used include: rhodamine conjugated anti-phalloidin (1:1,000; Sigma Aldrich), anti-β-catenin (1:100; BD Biosciences) and anti-E-cadherin (1:100; Zymed Laboratories Inc). Secondary antibodies were diluted to 1:300 in 5% host serum, 1% BSA in 1× PBS and incubated for 1 hr at RT. These included: goat anti-mouse Cy3 and donkey anti-goat Cy3 (Jackson ImmunoResearch Laboratories). DNA was counterstained with 1:10,000 bisbenzimide (Sigma) in 1× PBS and coverslips were mounted onto slides in 50% glycerol/1× PBS. Alternatively, staining TS cells using anti-laminin α5 (G-20, Santa Cruz) required that the cells be fixed with 4% PFA/1× PBS for 30 min and subsequently with −20°C methanol for 10 min on ice. Cells were blocked with 0.8% BSA/1× PBS and exposed to anti-laminin α5 (1:30 in 5% host serum, 1% BSA in 1× PBS) for 1 hr at RT. Similar conditions for secondary antibodies, counterstaining and coverslip mounting were used as above. For histological sections, rehydrated paraffin sections of E8.5 conceptuses were treated with 3% H2O2/1× PBS for 30 min and then with 1 mg/ml Trypsin (Sigma) for 10 min. Sections were blocked with 1% BSA/1× PBS for 30 min and exposed to the Ln α5 antibody (G-20, Santa Cruz) diluted 1:30 in 1% BSA/1× PBS for 1 hr followed by donkey anti-goat HRP-conjugated secondary antibody (1:300; Santa Cruz) in 1% BSA/1× PBS for 1 hr and exposure to DAB chromagen (Dako) for 2 min. Sections were counterstained in nuclear fast red and mounted in Cytoseal (Richard-Allen Scientific).
Time-lapse Video Microscopy
Time-lapse microscopy was carried out as previously described (Klaffky et al.,2006). Briefly, TS cells were plated on 35 mm Greiner tissue culture dishes with or without laminin-511 substrate coating (as above) and cultured in TS cell medium with FGF4, heparin and CM, overlaid with mineral oil. The cells were incubated in a PMDI-2 stage incubator (Harvard Apparatus) in 5% CO2 on an Olympus IX70 microscope equipped with phase optics. Digital images were collected every 2 min over a period of 4 hr. Images were captured using a Hamamatsu Orca camera driven by Openlab 2.0 software on a Macintosh G3 computer.
The cDNA probes used were previously described: Cebpα (Simmons et al.,2008), Esrrb (Luo et al.,1997), Gcm1 (Basyuk et al.,1999), Hand1 (Cross et al.,1995), Rhox4b (also known as Ehox) (Jackson et al.,2003), Snai1 (Smith et al.,1992), Synb and Syna (Simmons et al.,2008), Tpbpα (Lescisin et al.,1988), Ctsq (Simmons et al.,2007), Cdx2 (Tanaka et al.,1998), Prl3d1 (also known as Pl1) (Colosi et al.,1987).
Northern Blot Analysis
Total RNA from TS cell cultured with or without laminin substrate was collected at days 0, 2, 4 and 6 of differentiation and was isolated using QIAshredder and RNeasy columns (Qiagen, Inc.) following the manufacturer's instructions. Ten micrograms of total RNA was separated on a 1.1% formaldehyde agarose gel, blotted onto GeneScreen nylon membrane (Perkins Elmer) and UV cross-linked. Random-primed DNA labeling of cDNA probes was carried out with 25 μCi 32P-dCTP and probes were isolated on Sephadex G-50 columns (Amersham Biosciences Inc.). Hybridizations were done at 60°C overnight in hybridization buffer as previously described (Church and Gilbert,1984). Following posthybridization washes, signals were detected by exposure to BioMaz MR film (Kodak) at −80°C.
Knockdown of Cdh1 mRNA expression in TS cell cultures was conducted by the transfection of small interfering (si) RNAs. Commercially available siRNAs against Cdh1 were obtained from Qiagen (FlexiTube, #GS12550) and transfected into TS cell cultures using Lipofectamine 2000 (Invitrogen), according to manufacturers' instructions. A cocktail of four individual siRNAs designed to target Cdh1 were utilized at a final concentration of 10 μM (2.5 μM each). To control for transfection efficiency and off-target effects, a nonsense, GFP-labeled siRNA was included in all experiments and subsequent analyses (AllStars Negative Control siRNA, #SI03650318, Qiagen).
Quantitative RT-PCR Analysis
mRNA expression was assessed by quantitative reverse transcription (RT)-PCR using the SYBR green method. Briefly, total RNA from TS cells treated with siRNA and cultured with or without FGF4 was collected by lysis in RLT buffer and Qiashredder column (Qiagen) and processed on RNeasy columns (Qiagen) as outlined in the manufacturer's instructions. For each sample analyzed, 1 μg of total RNA was reverse transcribed using the QuantiTect Reverse Transcription Kit for SYBR green quantitative PCR (#205311, Qiagen). PCR reactions were then prepared using the QantiTect SYBR green PCR kit (#204143, Qiagen), according to the manufacturers' instructions, and thermocycling was conducted on an MRJ Research, DNA Engine, Opticon2 thermocycler. Primer sequences were as follows: Esrrb-F, 5′-AAC AGC CCC TAC CTG AAC CT, Esrrb-R, 5′-CTC ATC TGG TCC CCA AGT GT, Gcm1-F, 5′-CAT CTA CAG CTC GGA CGA CA, Gcm1-R, 5′-CCT TCC TCT GTG GAG CAG TC, Syna-F, 5′-TTG CAA TCA CAC CTT TCA GC, Syna-R, 5′-TGG TGT CCA CAG ACA GGG TA, Gapd-F, 5′-CCA GGA GCG AGA CCC CAC TAA CA, Gapd-R, 5′-TCG GCA GAA GGG GCG GAG. Primers for Rhox4b and Cdh1 were ordered from Qiagen (Quantitect Primer Assays, QT00121163 and QT00169743, respectively). Quantitative PCR reactions were conducted in triplicate for each gene and data collected from thermocycling was compiled and analyzed significant changes in gene expression using the Relative Expression Software Tool (REST) (Pfaffl et al.,2002).
Western Blot Analysis
Whole cell protein lysates were collected from wild-type (Rosa26) and Mrj−/− (line 1-4) TS cells cultured with growth factors for 2 days or without growth factors for 5 days. Using standard procedures, 60 μg of protein/well was electrophoresed on a 10% polyacrylamide gel and transferred onto an Immobilon-PSQ membrane (Millipore, cat no. ISEQ15150). The membrane was blocked in 3% donkey serum in TBST (Tris-buffered saline plus 0.1% tween-20) for 1 hr at RT and then incubated in 1:500 goat polyclonal anti-laminin α5 (G-20, Santa Cruz) diluted in 0.1% donkey serum/TBST overnight at 4°C. The membrane was washed in TBST and incubated in 1:5,000 HRP-conjugated donkey anti-goat IgG (Santa Cruz) diluted in 0.1% donkey serum/TBST for 1 hr at RT. Protein bands were detected with the ECL kit (Amersham Biosciences).
In Situ Hybridization
E8.25 implantation sites from Mrj+/− intercrosses were removed from the uterus, fixed overnight in 4% paraformaldehyde (PFA)/1× phosphate buffered saline (PBS) at 4°C, processed through a sucrose gradient and embedded in OCT compound (Sakura Finetek USA Inc.) for preparation of frozen sections. Eight μm frozen sections were adhered to Super Frost Plus slides (VWR International, Inc.) and stored at −80°C until used. The in situ hybridization protocol was carried out as previously described (Simmons et al.,2007).
Implantation sites from Mrj+/− intercrosses were dissected at E8.25, fixed, stained as whole mount for β-galactosidase expression and then embedded for sectioning as previously described (Hunter et al.,1999).
Equipment and Software
A Leica DMR light microscope (for fluorescent and light images) and Leica DMIL inverted light microscope (for cell culture images) with Photometrics Coolsnap cf camera and Openlab 2.2.2 imaging software program were used to obtain micrographs. Filters used were from Chroma: DAPI (31000) and TRITC (31002). Minimal image processing was performed using Adobe Photoshop 7.0 and figures were constructed using Canvas X.
We thank X. Zhao for help with electron microscopy images and Dr. J. Rossant for the Rosa26 TS cell line. This work was funded by grants to J.C. from the Canadian Institutes of Health Research (CIHR) and the Alberta Heritage Foundation for Medical Research (AHFMR). E.W. was supported by a studentship from the CIHR Strategic Training Program in Genetics, Child Health and Development and fellowships from the Centre for Trophoblast Research and the CIHR. D.S. was supported by fellowships from the Lalor Foundation and the AHFMR. D.N. was supported by fellowships from the CIHR and AHFMR. J.C. is a Scientist of the AHFMR.