Polarization and directional migration of X. laevis PGCs in vitro

4.1.1 In vitro PGC migration in the presence of different substrates

Active migration of X. laevis PGCs starts after developmental stage 24 and at tailbud stage embryos (stages 24-44) takes place within the endoderm (Nishiumi et al., 2005;

Terayama et al., 2013). Due to a high content of the yolk in endodermal cells, it is hard to study cellular and molecular mechanisms underlying active PGC migration. This issue can be solved by the establishment of an in vitro migration assay. In our previous studies, active migration of PGCs isolated from tailbud stage embryos was observed on the fibronectin-coated surface (Tarbashevich et al., 2011; Fig. 5). In the same study, it has also been shown that PGCs can be polarized towards crude extract prepared by homogenization of the dorsal part of tailbud stage embryos. However, the number of cells able to migrate in these conditions was very low. To improve migration efficiency, in the context of this study PGCs

were cultivated in the presence of different cellular and artificial substrates (see section 3.1).

These experiments demonstrated that PGCs migrate most efficiently in the so called under-agarose migration assay on top of the culture dish pre-coated with bovine serum albumin (BSA) (Fig. 12; section 3.1.2). In this case, PGCs were placed in between surface of the culture dish and polymerized agarose gel. Pre-coating of the culture dish with bovine serum albumin (BSA) was performed to saturate unspecific binding sites. In the under-agarose migration assay, PGCs migrate via bleb-associated mechanism (section 3.1.3). Migrating cells have elongated polarized morphology and can be characterized by waves of cell body contraction perpendicular to the leading edge–cell rear axis. Since no enrichment of actin filaments was observed at the leading edge of migrating cells (Fig. 14), traction force required for cell motility is most likely generated by contractions of the cell body that pushes the cell forward due to compressive forces applied on the agarose gel and surface of the culture dish. Similar morphology, but inability of PGCs to migrate in the assays with less rigid environment (extracellular matrix, embedding in the agarose gel and cultivation in the endodermal cells) favours this assumption (Fig. 8, 10, 11).

Studies in Drosophila, zebrafish and mouse demonstrated that surrounding somatic cells contribute to the establishment of directionality and survival of PGCs during active migration (see section 1.3). In Drosophila and zebrafish, somatic cells are involved in the shaping of the chemoattractant gradient required for directional migration (Boldajipour et al., 2008; Mahabaleshwar et al., 2008; Richardson and Lehmann, 2010). In addition, E-cadherin-mediated interaction of PGCs with somatic cells in zebrafish is required for PGC motility (Kardash et al., 2010). In mouse, survival and motility of PGCs depends on the Steel factor, expressed by surrounding somatic cells. Steel factor-expressing cells create a ‘motility niche’ and accompany PGCs all the way through their migration to the gonads (Gu et al., 2009). Role of somatic cells was also demonstrated for PGC migration through dorsal mesentery in X. laevis embryos. It was suggested, that distribution of fibronectin filaments produced by mesentery cells can guide PGC migration at the early tadpole stage of X. laevis development (Heasman et al., 1981; Wylie and Heasman, 1982; Brustis et al., 1984). In addition, PGCs, seeded on the layer of isolated fibronectin-producing mesentery cells, could form filopodia-like protrusions and translocate the cell body (Wylie and Roos, 1976;

Heasman et al., 1977). To test possible requirement of somatic cells for active migration of PGCs during tailbud stage of X. laevis development, isolated PGCs were cultured in the surrounding of pre-cultivated somatic endodermal cells (Fig. 8; section 3.1.1). During the first several hours of cultivation, PGCs revealed increased cellular dynamics, but could not translocate the cell body. Although cultivation of PGCs in these conditions was performed up to 24 hours, decrease of cellular dynamics after the first few hours, also observed during cultivation of PGCs in the presence of artificial substrates, indicates that somatic endodermal cells do not produce factors required for PGC motility. This can also be confirmed by our previous experiments with isolated PGCs cultivated in the presence of dorsal and ventral homogenized extracts. PGCs cultivated on top of fibronectin-coated Petri dish can be polarized towards crude homogenized extract prepared from the dorsal part of tailbud stage

embryos, but did not respond to the extract prepared from the ventral part, which consists mostly from endodermal cells (Tarbashevich et al., 2011; Fig. 5)

Active migration of PGCs in zebrafish and mouse depends on the gradient of chemoattractant SDF-1 (Doitsidou et al., 2002; Ara et al., 2003; Molyneaux et al., 2003). In these species, polarization of PGCs towards somatic cells expressing SDF-1 is followed by directional migration within the embryo (Kunwar et al., 2006). Since PGC migration takes place during active tissue- and organogenesis, migrating cells must quickly respond to the changes in chemoattractant gradient. In the under-agarose migration assay (described in sections 3.1.3 and 3.2) migrating X. laevis PGCs alternated between migratory ‘run‘ phase, characterized by an elongated polarized cell morphology and active migration, and

‘tumbling’ phase, characterized by a loss of cell polarity and formation of large bleb-like protrusions in random directions. An alternation between ‘run’ and ‘tumbling’ phases was also described for zebrafish PGCs (Reichman-Fried et al., 2004), and was recently reported for X. laevis PGC migration on fibronectin-coated surface (Terayama et al., 2013). It was suggested that ‘tumbling’ phase is required to re-establish cell polarity in the environment with dynamically changing gradient of chemoattractant. As it was shown previously, X. laevis PGCs can be polarized in vitro by a homogenized extract, prepared from the dorsal part of the tailbud stage embryos (Tarbashevich et al., 2011; Fig. 5). Interestingly, in the under-agarose migration assay, PGCs could initiate active migration even in the absence of any gradient (Fig. 12). This suggests that expression of putative chemoattractant(s) in the dorsal part of X. laevis embryos is important for PGC polarization, but is not required for PGCs migration. On the other hand, PGCs cultivated in different environmental conditions lost polarization and strongly decreased cellular dynamics several hours after dissociation from the endodermal explant. In the same time, ectopic addition of dorsal extract to the PGCs cultivated in the presence of endodermal cells resulted in a more intensive protrusion formation (Fig. 8; section 3.1.1). These observations can be explained by induction of PGC migration by some factors from the dorsal part of X. laevis embryos, and by a remaining potential of PGCs to migrate within few hours after removal of these factors. In the under-agarose migration assay, no increase in the efficiency of PGC migration upon application of dorsal extract can be explained by a poor diffusion of potential factors in the agarose gel.

4.1.2 Role of PIP3 in PGC polarization

Studies in Drosophila, zebrafish and mouse revealed that directionality of PGC migration, similar to chemotaxis in many other cell types, is mediated by G-protein coupled receptors (GPCR) (see section 1.3). In Drosophila, polarization and initiation of PGC migration depends on the expression of GPCR, known as Trapped in endoderm 1 (TRE1) (Richardson and Lehmann, 2010). In zebrafish and mouse, SDF-1 gradient is recognized by chemokine C-X-C motif receptor 4 (CXCR4) expressed in PGCs (Knaut et al., 2003; Molyneaux et al., 2003).

Interaction of GPCRs with their ligands activates dissociation of heterotrimeric G-proteins.

This, in turn, triggers activation of several downstream signaling cascades, including calcium flux, activation of phospholipase C (PLC) and phosphotidylinositide 3-kinase (PI3K) (Dutt et

al., 1998; Wang et al., 2000; Blaser et al., 2006). In Drosophila PGCs, activation of TRE1 was suggested to activate the small GTPase Rho1 and cause redistribution of adherent junctions and Rho1 to the cell rear (Kunwar et al., 2006). In zebrafish, activation of CXCR4b in PGCs leads to activation of the other small GTPase, Rac1, required for formation of the actin brushes at the leading edge of migrating cells (Xu et al., 2012). In addition, it also mediates enrichment of Ca²⁺ at the leading edge that is required for PGC polarization in zebrafish development express CXCR4 that is required for the directionality of PGC migration within the endoderm (Nishiumi et al., 2005; Takeuchi et al., 2010). In the previous studies from our lab, it was shown that formation of bleb-like protrusions and directional migration of X.

laevis PGCs depends on intracellular distribution of phosphatidylinositol (3,4,5)-trisphosphate (PIP3) (Tarbashevich et al., 2011). Aberrations in intracellular PIP3 levels, mediated by the modulation of endogenous activity of PI3K or PTEN, led to the defects in directionality of migration and loss of PGCs. In isolated PGCs, PIP3 is localized at the plasma membrane and is enriched in the bleb-like protrusions formed by these cells (Fig. 32A). This localization depends on the function of kinesin, xKIF13B, is encoded by the germ plasm-specific mRNA. PGC-plasm-specific overexpression of xKIF13B results in the enrichment of PIP3 throughout the plasma membrane and increased formation of bleb-like protrusions, while knock-down leads to the loss of PIP3 localization at the membrane and decreased protrusion formation. Both phenotypes cause defects in directionality of migration and loss of PGCs, similar to the aberrations in intracellular PIP3 levels (Tarbashevich et al., 2011).

In the present study, PGC-specific expression of pleckstrin homology (PH) domain of GRPI protein that served as a PIP3 sensor, was used to monitor endogenous PIP3 distribution in isolated X. laevis PGCs during active migration in the under-agarose assay. During the migratory ‘run’ phase, no enrichment of PIP3 at the leading edge of PGCs was observed.

However, enrichment of PIP3 was restored in the bleb-like protrusions formed by PGCs at the ‘tumbling’ phase (Fig. 15). This suggests PIP3 to be involved in polarization of the cell prior to active migration; however, maintenance of polarized PIP3 distribution during active phase of migration is not required. Interestingly, loss of polarized PIP3 distribution in PGCs, caused by knock-down and overexpression of xKIF13B, or by modulating endogenous activity of PI3K and PTEN, resulted in migration of these cells to ectopic locations in tailbud stage X.

laevis embryos (Tarbashevich et al., 2011). Similar PGC distribution was reported in X. laevis embryos with upregulated expression of SDf-1 (Bonnard et al., 2012). This phenotype is different in comparison to the clustering of PGCs in the endoderm observed in Dnd and Xdazl knock-down (Houston and King, 2000b; Horvay et al., 2006). Ectopic localization of PGCs was also observed in zebrafish and mouse embryos upon knock-down of SDF-1 and its

receptor CXCR4 (Ara et al., 2003; Molyneaux et al., 2003; Reichman-Fried et al., 2004).

Altogether, these observations suggest that polarized PIP3 distribution in X. laevis PGCs is required for cell polarization and directionality of migration at the tailbud stage, but not for cellular motility.