TY - JOUR
AU - Coelho, Susana M.
AB - Abstract Brown algae are one of the most developmentally complex groups within the eukaryotes. As in many land plants and animals, their main body axis is established early in development, when the initial cell gives rise to two daughter cells that have apical and basal identities, equivalent to shoot and root identities in land plants, respectively. We show here that mutations in the Ectocarpus DISTAG (DIS) gene lead to loss of basal structures during both the gametophyte and the sporophyte generations. Several abnormalities were observed in the germinating initial cell in dis mutants, including increased cell size, disorganization of the Golgi apparatus, disruption of the microtubule network, and aberrant positioning of the nucleus. DIS encodes a TBCCd1 protein, which has a role in internal cell organization in animals, Chlamydomonas reinhardtii, and trypanosomes. Our study highlights the key role of subcellular events within the germinating initial cell in the determination of apical/basal cell identities in a brown alga and emphasizes the remarkable functional conservation of TBCCd1 in regulating internal cell organization across extremely distant eukaryotic groups. Events in the zygote leading up to and including the first cell division play a key role in the developmental patterning of multicellular organisms. In many plants and animals, asymmetric division of the zygote establishes the principal body axis of the early embryo, representing the first major patterning event during embryogenesis (Gönczy and Rose, 2005; Ueda and Laux, 2012). In land plants, the first cell division usually coincides with the establishment of apical and basal cell identities (Ueda and Laux, 2012) (but note that some fern gametophytes have tripolar germination patterns; Schneider, 2013). In Arabidopsis thaliana, the asymmetrical division of the zygote is associated with a number of cellular events, including movement of the nucleus to the apex of the cell, repositioning of other major organelles, formation of a large vacuole in the basal part of the cell, and reorganization of the microtubules into transverse cortical arrays associated with cell outgrowth in the apical direction (Jeong et al., 2011a; Kimata et al., 2016; Lau et al., 2012). The establishment of the apical-basal axis in Arabidopsis has been shown to be controlled by a complex genetic network involving both auxin-dependent and auxin-independent pathways (Lau et al., 2012). The auxin-dependent network involves the transcription factor MONOPTEROS/AUXIN RESPONSE FACTOR5, the auxin response inhibitor BODENLOS/INDOLE-3-ACETIC-ACID12, and the auxin efflux carrier PIN-FORMED7, whereas the auxin-independent pathway involves EMBRYO SURROUNDING FACTOR1, the receptor-like cytosolic kinase SHORT SUSPENSOR, the MAP kinase kinase YODA, the MAP kinases MPK3 and MPK6, and several transcription factors, including GROUNDED/RKD4 and WOX2 (Jeong et al., 2011b; Waki et al., 2011; Costa et al., 2014; Lau et al., 2012; Rademacher et al., 2012). However, it is not yet clear how these genetic networks implement the cellular events that underlie the formation of the apical-basal axis and the establishment of apical and basal cell identities. The brown algae have evolved complex multicellularity independently of land plants, but the two lineages share several key developmental characteristics, such as the importance of the establishment of the apical/basal axis during development, extensive postembryonic developmental patterning, and developmental programs constrained by the absence of cell migration. The apical and basal structures of brown algae and land plants can also be considered to be analogous, with the basal systems of brown algae being functionally equivalent to rooting structures in land plants. In both lineages, the basal systems are composed of tip-growing filamentous rhizoid cells that extend into the substrate or air/water surrounding the plant and are used for anchoring (Jones and Dolan, 2012). In both brown algae and flowering plants, apical/basal polarity is usually determined very early, before the first cell division (Fritsch, 1935). However, this process may be less complex in brown algae because the development of initial cells (e.g., the zygote) does not occur within maternal tissues. Consequently, axis formation is expected to occur in a cell autonomous manner (but see Whitaker, 1931) and not to be influenced by signals emitted by nearby parental cells (Costa et al., 2014). The brown alga Fucus has been used for many years as a model system to study apical-basal axis formation during embryogenesis (Coelho et al., 2002; Brownlee and Bouget, 1998; Kropf et al., 1988). As in land plants, the zygote cell divides asymmetrically to establish the apical-basal axis (Bouget et al., 1998; Goodner and Quatrano, 1993). The two products of this division go on to produce the apical and basal systems of the alga, the thallus and the rhizoid, respectively (Brownlee and Bouget, 1998). The establishment of the apical-basal axis in Fucus has been studied in detail at the cellular level and has been shown to involve position-dependent information from the cell wall (Berger et al., 1994). Apoplastic diffusible gradients appear also to be involved in pattern formation in the multicellular Fucus embryo (Bouget et al., 1998). While the large zygote and the external fertilization process (fusion of male and female gametes released from the parent thallus) of Fucus makes it well adapted for the study of the cell biology of axis formation, this genus is not amenable to genetic approaches and, consequently, the genetic networks that control apical-basal axis formation have not been characterized in the brown alga lineage. Recently, the filamentous alga Ectocarpus sp has emerged as a genetic model for the brown algae (Coelho et al., 2012c; Cock et al., 2014). A high-quality genome sequence is available for this species (Cock et al., 2010; Cormier et al., 2017), together with extensive transcriptomic data (Lipinska et al., 2015; Luthringer et al., 2015) and genetic tools including a dense genetic map (Heesch et al., 2010; Avia et al., 2017). Genetic transformation is not yet possible, but gene knockdown using RNA interference has been demonstrated (Macaisne et al., 2017). These various tools were employed in a recent study that was the first to identify a brown algal developmental gene using a forward genetic approach (Macaisne et al., 2017). Ectocarpus has a complex life cycle involving alternation between multicellular sporophyte and gametophyte generations (Coelho et al., 2012c; Peters et al., 2008) (Supplemental Figure 1). Unlike land plants, both the sporophyte and the gametophyte are derived from single cells, which are released into the seawater medium (gametes and meiospores, respectively) greatly facilitating the analysis of early events during development. This characteristic also means that any parental influence on developmental processes is essentially limited to information provided to the propagules before their release (but see Arun et al., 2013). Both the gametophyte and the sporophyte possess an apical-basal axis and clearly defined apical and basal filamentous systems (Coelho et al., 2011; Peters et al., 2008). In the gametophyte, the basal system consists of rhizoids and the apical system is composed of upright filaments that bear the gametangia. In the sporophyte, the basal system is more extensive, consisting of a network of firmly attached basal filaments, whereas the apical system resembles that of the gametophyte, consisting of branched upright filaments that bear the sexual structures. The general similarities between the gametophyte and sporophyte generations, in terms of their size and overall morphologies, make Ectocarpus an ideal system to investigate how two distinct developmental programs can be deployed from the same genome. In this study, we report the identification of the DISTAG (DIS) locus, which is required for the formation of basal structures during both the sporophyte and gametophyte generations of the Ectocarpus life cycle. DIS encodes a Tubulin Binding Cofactor C (TBCC) domain protein of the TBCCd1 class. Mutations in the Ectocarpus DIS gene are associated with several modifications at the initial cell stage: disorganization of the Golgi apparatus, increased cell size, disruption of the pattern of the microtubule network, and aberrant positioning of the nucleus. dis mutants therefore link subcellular events within the initial cell with the acquisition of apical/basal cell identities. The phenotypes of dis mutants also confirm that the basal filament system of the sporophyte is developmentally equivalent to the rhizoid of the gametophyte generation, providing insight into the evolutionary events that led to the emergence of the sporophyte and gametophyte developmental programs in this species. RESULTS dis Mutants Lack a Basal System During the Ectocarpus gametophyte generation, the apical/basal axis is established in the initial cell, prior to the first cell division. The two cells derived from the division of the initial cell grow, in the form of two germ tubes, to establish a rhizoid (basal, root-like organ) and a filamentous thallus (apical, shoot-like organ) (Figure 1). A UV mutagenesis screen identified two mutant strains (Ec722 and Ec799; Supplemental Table 1 and Supplemental Figure 2) that failed to develop any of the basal structures normally observed in the wild-type gametophyte generation, i.e., the initial cells of the mutants immediately developed as apical upright filaments. Figure 1. Open in new tabDownload slide Developmental Patterning Is Affected in dis Mutants. The apical systems of both the sporophyte and gametophyte generations are composed of apical upright cells (dark yellow) and reproductive structures: sporangia in the sporophyte (green) and gametangia in the gametophyte (purple). The basal system (red) of the wild-type gametophyte generation is composed of a rhizoid, whereas that of the sporophyte generation is composed of basal prostrate cells. The establishment of the basal systems during both the gametophyte and sporophyte generations is affected in dis mutants. Basal structures are indicated in red, apical structures in yellow, green, and purple, and the initial cell and nondifferentiated cells in gray. Bars = 15 µm. (A) Wild-type gametophyte; rhizoid marked with an arrowhead. (B) Apical upright cells of a wild-type gametophyte. (C) Rhizoids forming on apical cells of a wild-type gametophyte. (D) Plurilocular gametangium on a wild-type gametophyte. (E) Young dis-1 gametophyte; the asterisk marks the initial cell. (F) Apical upright cells of a dis-1 gametophyte. (G) Plurilocular gametangium of a dis-1 gametophyte. (H) Prostrate cells of a wild-type sporophyte basal system. (I) Upright cells of the apical system of a wild-type sporophyte. (J) Wild-type unilocular sporangium. (K) Wild-type plurilocular sporangium. (L) Secondary rhizoid developing from the apical system of the sporophyte generation. (M) Young dis-1 sporophyte; the initial cell is marked with an asterisk. (N) Apical upright cells of a dis-1 sporophyte. (O) Unilocular sporangium on a dis-1 upright filament. (P) Plurilocular sporangium on a dis-1 sporophyte. Figure 1. Open in new tabDownload slide Developmental Patterning Is Affected in dis Mutants. The apical systems of both the sporophyte and gametophyte generations are composed of apical upright cells (dark yellow) and reproductive structures: sporangia in the sporophyte (green) and gametangia in the gametophyte (purple). The basal system (red) of the wild-type gametophyte generation is composed of a rhizoid, whereas that of the sporophyte generation is composed of basal prostrate cells. The establishment of the basal systems during both the gametophyte and sporophyte generations is affected in dis mutants. Basal structures are indicated in red, apical structures in yellow, green, and purple, and the initial cell and nondifferentiated cells in gray. Bars = 15 µm. (A) Wild-type gametophyte; rhizoid marked with an arrowhead. (B) Apical upright cells of a wild-type gametophyte. (C) Rhizoids forming on apical cells of a wild-type gametophyte. (D) Plurilocular gametangium on a wild-type gametophyte. (E) Young dis-1 gametophyte; the asterisk marks the initial cell. (F) Apical upright cells of a dis-1 gametophyte. (G) Plurilocular gametangium of a dis-1 gametophyte. (H) Prostrate cells of a wild-type sporophyte basal system. (I) Upright cells of the apical system of a wild-type sporophyte. (J) Wild-type unilocular sporangium. (K) Wild-type plurilocular sporangium. (L) Secondary rhizoid developing from the apical system of the sporophyte generation. (M) Young dis-1 sporophyte; the initial cell is marked with an asterisk. (N) Apical upright cells of a dis-1 sporophyte. (O) Unilocular sporangium on a dis-1 upright filament. (P) Plurilocular sporangium on a dis-1 sporophyte. During the early development of the wild-type sporophyte generation, establishment of apical structures is delayed and an extensive system of prostrate filaments consisting of round cells is formed before the apical thallus filaments develop (Peters et al., 2008; Figure 1). Several observations indicate that this basal network of prostrate filaments is equivalent to the basal system of the gametophyte generation, represented by the rhizoid: (1) both structures develop at the base of the alga (in contact with the substratum), (2) both structures serve an anchoring function, and (3) in sporophytes carrying the immediate upright (imm) mutation the system of prostrate filaments is homeotically replaced by a rhizoid similar to that of the gametophyte (Peters et al., 2008; Macaisne et al., 2017). The sporophyte generations of the Ec722 and Ec799 mutants failed to produce a network of prostrate filaments and therefore also lacked basal structures (Figure 1). The establishment of reproductive structures on apical systems was unaffected in both generations, and the mutants were fully fertile as both gametophytes and sporophytes (Figure 1). The absence of a basal system significantly affected the capacity of the mutants to adhere to the substratum (Figure 2); consequently, Ec722 and Ec799 were named distag-1 (dis-1) and dis-2, respectively (“distag” means “detached” in the Breton language). The loss of basal systems during both the gametophyte and sporophyte generation in dis mutants is consistent with these two systems (rhizoids and prostrate filaments, respectively) being developmentally equivalent between the two generations. Developmental equivalence between the basal systems of the two generations was further supported by the observation that when dis-1 imm double mutants were constructed, they lacked the rhizoid that replaces the prostrate filaments in the imm mutant (Figure 2). Analysis of a segregating family generated from a cross between dis IMM and DIS imm parents indicated that DIS is epistatic to IMM (Supplemental Table 2). Figure 2. Open in new tabDownload slide Loss of Basal Cell-Type Specification in dis Mutants. (A) and (B) Representative image of a dis-1 imm double mutant (A) and a dis-1 individual (B) for comparison. Note that the two mutants have the same phenotype, indicating that DIS is epistatic to IMM. (C) Wild-type gametophyte at the two-cell stage; (D) Eight-day-old wild-type gametophyte; arrows indicate the bipolar germination pattern. (E) Wild-type sporophyte at the two-cell stage. (F) Wild-type sporophyte at the five-cell stage; arrows indicate the bipolar germination pattern. (G) dis-2 gametophyte at the two-cell stage. (H) Seven-day-old dis-2 gametophyte; arrow indicates the unipolar germination pattern. (I) dis-2 sporophyte at the two-cell stage. (J) Eight-day-old dis-2 sporophyte. Arrow indicates the unipolar germination pattern. In (B) to (J), arrowheads indicate the position of the first cell division plane and asterisks indicate the initial cells. (K) Abnormal basal system cells that replace the second germ tube in the dis-1 mutant. (L) Absence of rhizoids in the gametophyte generation of dis mutants causes premature detachment from the substratum. At 7 d after release, only 15% of the wild-type individuals had detached from the substratum (free-floating), compared with 90% of dis-1 and 94% of dis-2 individuals (n = 1225). Different letters above the bars indicate significant differences (Wilcoxon test, P value = 0.0079 for the wild-type versus dis-1 comparison and P value = 0.011 for the wild type versus dis-2 comparison). (M) Cell regeneration in dis versus wild-type filaments. Regeneration of rhizoids from wild-type sporophyte apical cells 4 d after cutting the filament by microdissection. The dashed line indicates the position of the cut, and the arrowheads indicate the regenerating cell. (N) dis-1 individuals do not regenerate rhizoids in response to wounding: dis-1 representative filament 72 h after microdissection. Cylindrical apical cells regenerate instead of rhizoids. The same results were obtained for dis-1 and for dis-2: 100% of the microdissected cells regenerated as apical cells, in contrast to the wild type, where 100% of the cells regenerated as rhizoids. The images shown are representative of the 82 individuals that were analyzed in three independent experiments. Bars = 5 µm in (C), (E), (G), and (I), 20 µm in (B), (D), (F), (H), (J), and (K), and 10 µm in (M) and (N). Figure 2. Open in new tabDownload slide Loss of Basal Cell-Type Specification in dis Mutants. (A) and (B) Representative image of a dis-1 imm double mutant (A) and a dis-1 individual (B) for comparison. Note that the two mutants have the same phenotype, indicating that DIS is epistatic to IMM. (C) Wild-type gametophyte at the two-cell stage; (D) Eight-day-old wild-type gametophyte; arrows indicate the bipolar germination pattern. (E) Wild-type sporophyte at the two-cell stage. (F) Wild-type sporophyte at the five-cell stage; arrows indicate the bipolar germination pattern. (G) dis-2 gametophyte at the two-cell stage. (H) Seven-day-old dis-2 gametophyte; arrow indicates the unipolar germination pattern. (I) dis-2 sporophyte at the two-cell stage. (J) Eight-day-old dis-2 sporophyte. Arrow indicates the unipolar germination pattern. In (B) to (J), arrowheads indicate the position of the first cell division plane and asterisks indicate the initial cells. (K) Abnormal basal system cells that replace the second germ tube in the dis-1 mutant. (L) Absence of rhizoids in the gametophyte generation of dis mutants causes premature detachment from the substratum. At 7 d after release, only 15% of the wild-type individuals had detached from the substratum (free-floating), compared with 90% of dis-1 and 94% of dis-2 individuals (n = 1225). Different letters above the bars indicate significant differences (Wilcoxon test, P value = 0.0079 for the wild-type versus dis-1 comparison and P value = 0.011 for the wild type versus dis-2 comparison). (M) Cell regeneration in dis versus wild-type filaments. Regeneration of rhizoids from wild-type sporophyte apical cells 4 d after cutting the filament by microdissection. The dashed line indicates the position of the cut, and the arrowheads indicate the regenerating cell. (N) dis-1 individuals do not regenerate rhizoids in response to wounding: dis-1 representative filament 72 h after microdissection. Cylindrical apical cells regenerate instead of rhizoids. The same results were obtained for dis-1 and for dis-2: 100% of the microdissected cells regenerated as apical cells, in contrast to the wild type, where 100% of the cells regenerated as rhizoids. The images shown are representative of the 82 individuals that were analyzed in three independent experiments. Bars = 5 µm in (C), (E), (G), and (I), 20 µm in (B), (D), (F), (H), (J), and (K), and 10 µm in (M) and (N). In wild-type Ectocarpus, secondary rhizoids are produced from the apical system cells at a late stage of development (Peters et al., 2008) (Figure 1). These rhizoids can be considered to be analogous to the adventitious roots produced from the stems of some land plants (Atkinson et al., 2014). The dis-1 and dis-2 mutants failed to produce secondary rhizoids during both the sporophyte and the gametophyte generations. Hence, production of all basal structures, both primary and secondary, was blocked in these mutants. In fucoid brown algae, the apical system (the thallus) responds to damage by producing rhizoids (Bouget et al., 1998), and we observed a similar phenomenon in wild-type Ectocarpus. In contrast to the wild type, wounded apical filaments of the dis mutants failed to regenerate rhizoids and instead produced new cylindrical apical filament cells (Figure 2; Supplemental Figure 3). Taken together, our results indicate that basal system formation fails to occur, at all developmental stages, in the absence of a functional DIS gene product. One consequence of the loss of basal structures in the dis mutants is that the sporophyte and gametophyte generations are more similar than in the wild type because in both cases the vegetative thalli consist uniquely of upright filaments made up of cylindrical cells. Despite this resemblance, the sporophyte and gametophyte generations of the dis mutants retained generation-specific features. First, morphometric measurements showed that the length and width of the filament cylindrical cells and the angle at which secondary upright filaments branch off from primary filaments statistically distinguished sporophyte from gametophyte thalli in both wild-type and dis mutant strains. In contrast, no significant differences were detected when these parameters were compared between the same generation of wild type and mutant strains (Supplemental Table 3). Second, at maturity, dis gametophytes produced plurilocular gametangia, which contained gametes that were able to fuse with gametes of the opposite sex to produce zygotes. The sporophyte generation of the dis mutants, on the other hand, produced plurilocular sporangia containing spores (incapable of fusion) and unilocular sporangia with haploid, meiosis-derived meiospores. Finally, the expression patterns of generation-specific marker genes (Peters et al., 2008) were consistent with assigned life cycle generations in both wild-type and dis individuals (Supplemental Figure 4). Analysis of the dis Transcriptome To further characterize the Dis− phenotype, an RNA-seq approach was employed to study gene expression in the dis-1 mutant compared with wild-type basal and apical tissues (Supplemental Data Set 1). Consistent with the fact that dis has lost the ability to produce a basal system and is composed exclusively of apical system cells, analysis of dis transcriptional profiles indicated that they were more similar to apical than to basal transcriptomes of the wild-type strain (Figures 3A and 3B; Supplemental Figure 5). Figure 3. Open in new tabDownload slide Gene Expression Patterns in dis-1 Compared with Wild-Type Basal and Apical Systems. (A) Heat map representing log2(FC) of gene expression in pairwise analyses of (1) dis-1 and wild-type apical system, (2) dis-1 and wild-type basal system, and (3) wild-type apical and wild-type basal system transcriptomes. The analysis included only genes that were significantly differentially expressed in at least one of the comparisons (transcripts per million [TPM] > 1, FC > 2, or FC < 0.5, Padj < 0.1). Black color indicates no significant difference in expression between samples, red color indicates overexpression (TPM > 1, FC > 2, Padj < 0.1) and green color indicates under expression (TPM > 1, FC < 0.5, Padj < 0.1). Two clusters of genes are marked on the graph, cluster 1 (containing genes expressed at the same level in the dis-1 mutant and apical system but upregulated in both samples compared with basal system) and cluster 2 (containing genes expressed at the same level in dis-1 and apical system but downregulated in both samples in relation to the basal system). See also Supplemental Data Set 2. (B) Heat map representing log2(TPM) of gene expression in dis-1, wild-type apical system, and wild-type basal system. For clarity, the heat map presents a subset of 163 genes that were significantly differentially expressed in at least one of the pairwise comparisons between dis-1, apical system, and basal system (TPM > 1, FC > 2, Padj < 0.1). These 163 genes belong to two clusters of genes that were either expressed at the same level in the dis-1 mutant and apical system but upregulated in both samples compared with basal system or genes expressed at the same level in dis-1 and apical system but downregulated in both samples in relation to the basal system. (C) Word cloud, where the font size is proportional to the relative word frequency, for the different functional categories. Upper, middle, and lower panels represent gene functional categories from genes belonging to the clusters in Figure 3A (Supplemental Data Set 2), the top 200 most differentially expressed genes (Supplemental Data Set 3), and the set of differentially expressed genes whose proteins have signal peptides (Supplemental Data Set 4), respectively. Genes with unknown functions were excluded from this analysis. A statistical analysis of the enrichment in each of the functional categories is presented in Supplemental Table 4 (χ2 test, P < 0.05). Figure 3. Open in new tabDownload slide Gene Expression Patterns in dis-1 Compared with Wild-Type Basal and Apical Systems. (A) Heat map representing log2(FC) of gene expression in pairwise analyses of (1) dis-1 and wild-type apical system, (2) dis-1 and wild-type basal system, and (3) wild-type apical and wild-type basal system transcriptomes. The analysis included only genes that were significantly differentially expressed in at least one of the comparisons (transcripts per million [TPM] > 1, FC > 2, or FC < 0.5, Padj < 0.1). Black color indicates no significant difference in expression between samples, red color indicates overexpression (TPM > 1, FC > 2, Padj < 0.1) and green color indicates under expression (TPM > 1, FC < 0.5, Padj < 0.1). Two clusters of genes are marked on the graph, cluster 1 (containing genes expressed at the same level in the dis-1 mutant and apical system but upregulated in both samples compared with basal system) and cluster 2 (containing genes expressed at the same level in dis-1 and apical system but downregulated in both samples in relation to the basal system). See also Supplemental Data Set 2. (B) Heat map representing log2(TPM) of gene expression in dis-1, wild-type apical system, and wild-type basal system. For clarity, the heat map presents a subset of 163 genes that were significantly differentially expressed in at least one of the pairwise comparisons between dis-1, apical system, and basal system (TPM > 1, FC > 2, Padj < 0.1). These 163 genes belong to two clusters of genes that were either expressed at the same level in the dis-1 mutant and apical system but upregulated in both samples compared with basal system or genes expressed at the same level in dis-1 and apical system but downregulated in both samples in relation to the basal system. (C) Word cloud, where the font size is proportional to the relative word frequency, for the different functional categories. Upper, middle, and lower panels represent gene functional categories from genes belonging to the clusters in Figure 3A (Supplemental Data Set 2), the top 200 most differentially expressed genes (Supplemental Data Set 3), and the set of differentially expressed genes whose proteins have signal peptides (Supplemental Data Set 4), respectively. Genes with unknown functions were excluded from this analysis. A statistical analysis of the enrichment in each of the functional categories is presented in Supplemental Table 4 (χ2 test, P < 0.05). Several analyses were performed to characterize the set of genes that were differentially expressed in the three tissues analyzed. These included an analysis of the putative functions of clusters of genes with shared patterns of regulation (Supplemental Data Set 2), an analysis of the 200 most differentially expressed genes (100 most up- and 100 most downregulated) in these comparisons (Supplemental Data Set 3), an analysis of the differentially regulated genes that coded for proteins with putative signal peptides (representing putative secreted proteins; Supplemental Data Set 4) and a KEGG pathway analysis (Supplemental Data Set 5). Global patterns in the changes in gene expression were analyzed using a set of manually assigned functional categories (based on the functional categories used in Tarver et al., 2015). Gene Ontology (GO) term enrichment analysis (Blast2Go; Conesa and Götz, 2008) and KEGG analysis were then used to provide more detailed information about the processes and pathways affected. The relative frequency of the manually assigned functional categories was visualized using a word cloud (Figure 3C). Note that about half of the genes in the transcriptome data sets were of unknown function, and these were excluded from our analysis. Putative Basal and Apical System Effector Genes The gene cluster analysis focused on genes that were either up- or downregulated in both the dis-1 mutant and wild-type apical tissues compared with wild-type basal tissues (Figures 3A and 3B; Supplemental Data Set 2). The aim of this approach was to identify potential basal and apical system effector genes. The set of genes that was upregulated in dis and the wild-type apical system compared with the wild-type basal system (potential apical system effectors) were enriched in the GO categories “kinase activity,” “protein modification,” “cell differentiation,” and “cell communication” (Fisher's exact test, P < 0.05), while the genes that were downregulated in both dis-1 and the apical system compared with the basal system (potential basal system effectors) were enriched in categories related to “transcriptional activity,” “signal transduction,” and “extracellular” or “membrane-located processes” (Supplemental Data Set 2). Accordingly, the assignment of basal and apical effector genes to manually curated functional groups highlighted an important proportion of genes coding for proteins putatively involved in membrane functions and transport, cell wall biosynthesis and modification, vesicle trafficking, adhesion, cell regulation, and signaling (Figure 3C, upper panel; Supplemental Table 4). Top 200 Most Differentially Regulated Genes GO term enrichment analysis of the 200 most differentially regulated genes in the dis-1 mutant compared with wild-type basal and apical systems highlighted a range of GO term categories (Supplemental Data Set 3), but there was a particular abundance of genes with functions related to “membrane,” “oxidation-reduction process,” and “carbohydrate metabolic processes.” The cellular components highlighted by the GO term analyses were “membrane” and “extracellular region.” Manual assignment to functional categories revealed that the common functional groups represented within this set of genes included cell wall biosynthesis and organization, membrane function and transport, cell regulation and signaling, adhesion, vesicle transport, and cytoskeleton (Figure 3C, middle panel; Supplemental Table 4). Secreted Proteins The set of differentially expressed genes that were predicted to encode secreted proteins were analyzed in detail because the phenotypes of the dis mutants are associated with disruption of the Golgi apparatus. Interestingly, a disproportionate number of genes coding for proteins with a putative signal peptide were present in the pool of genes that were differentially regulated in dis versus wild-type basal system (Fisher's test, P value = 0.00002877 and P value = 0.01992 for up- and downregulated genes, respectively; Supplemental Data Set 1). Analysis of the GO terms associated with these genes indicated enrichment for a range of GO categories including carbohydrate transport, transmembrane transport, extracellular components, integral components of the membrane, transmembrane signaling, and Golgi-associated vesicles (Supplemental Data Set 4). The most commonly represented manually annotated functional categories represented within this data set included, as above, cell wall, adhesion, extracellular processes, membrane function and transporters, cytoskeleton, and cellular regulation and signaling (Figure 3C, lower panel). KEGG Analysis Analysis of the assignment of the differentially expressed genes to KEGG pathways (Xie et al., 2011) revealed a number of processes that were differentially regulated in dis versus wild-type samples. Pathways involved in interactions with the extracellular matrix, protein processing in the endoplasmic reticulum, and protein export were upregulated in the dis-1 mutant compared with the wild type basal system, whereas signaling pathways and protein processing in the endoplasmic reticulum were downregulated (Supplemental Figure 7 and Supplemental Data Set 5). Taken together, comparison of the dis-1 transcriptome with wild-type basal and apical tissues suggested that carbohydrate/cell wall-related processes, membrane transport, cellular signaling, and secretory activity may be affected in dis mutants. Note that transcript abundance was also measured using an independent method (RT-qPCR) for 12 of the differentially regulated genes to verify that the differential expression patterns observed were robust (Supplemental Figure 6). Globally, the results of the RT-qPCR analysis confirmed those obtained based on the RNA-seq data (Spearman's correlation R 2 = 0.619, P value = 0.0012). Loss of Bipolar Germination from the Initial Cell in dis Mutants The vast majority of initial cells of both the sporophyte and gametophyte generations of the dis mutants produced a single germ tube rather than the two germ tubes normally observed with wild-type strains (Peters et al., 2008). This phenotype was observed more consistently with the dis-2 mutant than with the dis-1 mutant (85% and 95% for dis-1 and dis-2 sporophytes respectively, n = 591; 82% and 95% for dis-1 and dis-2 gametophytes, respectively, n = 54) (Figure 2). We also noted that, while the dis-2 mutant completely failed to produce a second germ tube, a proportion of dis-1 individuals (65%, n = 235) produced one or more enlarged and abnormally shaped cells at the end where the second germ tube would normally emerge, possibly corresponding to an aborted basal system (Figure 2). Genetic Analysis and Identification of the DIS Gene Sporophytes can be propagated asexually through the production of mitospores (Figure 1). The Dis− mutant phenotype was stable through 30 rounds of asexual generations via mitospores. A male dis-1 gametophyte (Ec722) was crossed with an outcrossing wild-type female gametophyte of the strain Ec568 (Coelho et al., 2011; Peters et al., 2008) (Supplemental Table 1 and Supplemental Figure 2). The resulting sporophyte (Ec653) exhibited a wild-type pattern of development, indicating that the dis-1 mutation was recessive. A segregating population of 200 individuals derived from this cross consisted of 92 and 108 phenotypically wild-type and mutant individuals, respectively, consistent with a 1:1 segregation ratio and Mendelian inheritance of a single-locus recessive mutation (χ2 test = 1.28, df = 1, P value = 0.257). A female individual from this progeny that carried the dis-1 mutation was crossed with a male gametophyte of the dis-2 strain (Ec799). The resulting sporophytes (Ec808 and Ec809) exhibited a Dis− phenotype, indicating that the two mutations dis-1 and dis-2 were allelic. A cloning-by-sequencing approach (Schneeberger et al., 2009) identified a candidate locus on chromosome 05 for the location of the mutation in the dis-1 mutant (Figure 4A) and sequencing of this genomic region identified a single nucleotide mutation at position 3,330,523 that was present in the dis-1 mutant (Ec722) but absent from the Ec32 and Ec568 wild-type strains (Figure 4B). This mutation was located in intron 15 of the gene Ec-05_001860. A cleaved amplified polymorphic sequence (CAPS) marker, developed based on the candidate causal mutation, showed absolute cosegregation of this mutation with the Dis− phenotype in 265 individuals of the segregating family derived from sporophyte Ec653. Resequencing of the Ec-05_001860 gene in the dis-2 mutant identified a point mutation in exon 13 that results in the introduction of a stop codon into the coding region of the gene (Figure 4B). Taken together, these analyses provide strong evidence that Ec-05_001860 corresponded to the DIS gene. Figure 4. Open in new tabDownload slide Identification of the dis-1 and dis-2 Mutations and Phylogenetic Analysis of the DIS Protein. (A) Cloning-by-sequencing identification of the genetic lesion responsible for the Dis− phenotype in the dis-1 mutant. Genetic marker density along the genome (marker/kb) and genetic marker frequency (mean per scaffold). (B) Diagram showing the positions of the dis-1 (intron 15) and dis-2 (exon 13) mutations. The point mutation in exon 13 in dis-2 mutant results in the introduction of a stop codon into the coding region of the gene (indicated by an asterisk). Dark-blue boxes indicate untranslated regions, and light-blue indicates protein-coding exons and the corresponding regions of the protein. (C) Introduction of siRNAs targeting the DIS gene mimics the phenotype of the dis mutations. Compared with the control treatment (siRNA designed against another Ectocarpus sp gene, Ec_13-001890), significantly more cells presented a Dis− phenotype in samples treated with siRNA directed against DIS (Kruskal-Wallis χ2 = 9.1988, df = 3, P value = 0.02676). Asterisks indicate significant differences in relation to the control treatment. The graph is representative of three independent experiments. (D) Unrooted maximum likelihood tree of TBCC domain protein sequences. Ec-05_001860 is the DISTAG protein. Please see Supplemental Table 6 for the taxa and protein sequences used. Figure 4. Open in new tabDownload slide Identification of the dis-1 and dis-2 Mutations and Phylogenetic Analysis of the DIS Protein. (A) Cloning-by-sequencing identification of the genetic lesion responsible for the Dis− phenotype in the dis-1 mutant. Genetic marker density along the genome (marker/kb) and genetic marker frequency (mean per scaffold). (B) Diagram showing the positions of the dis-1 (intron 15) and dis-2 (exon 13) mutations. The point mutation in exon 13 in dis-2 mutant results in the introduction of a stop codon into the coding region of the gene (indicated by an asterisk). Dark-blue boxes indicate untranslated regions, and light-blue indicates protein-coding exons and the corresponding regions of the protein. (C) Introduction of siRNAs targeting the DIS gene mimics the phenotype of the dis mutations. Compared with the control treatment (siRNA designed against another Ectocarpus sp gene, Ec_13-001890), significantly more cells presented a Dis− phenotype in samples treated with siRNA directed against DIS (Kruskal-Wallis χ2 = 9.1988, df = 3, P value = 0.02676). Asterisks indicate significant differences in relation to the control treatment. The graph is representative of three independent experiments. (D) Unrooted maximum likelihood tree of TBCC domain protein sequences. Ec-05_001860 is the DISTAG protein. Please see Supplemental Table 6 for the taxa and protein sequences used. A protocol for genetic transformation does not exist for Ectocarpus (or for any other brown alga) but injection of double stranded RNA into zygotes of the brown alga Fucus has been shown to induce an RNA interference (RNAi) response, leading to knockdown of target gene expression (Farnham et al., 2013). We recently adapted this Fucus RNAi protocol for Ectocarpus, using synthetic siRNA molecules instead of long double-stranded RNA molecules and a lipofectant specifically adapted for dsRNA delivery to introduce the siRNAs into the cell rather than the microinjection procedure used for Fucus (Macaisne et al., 2017). Following simultaneous introduction of three siRNA molecules targeting the DIS gene, a small proportion (∼2.5%) of the parthenogenetic gametes exhibited a pattern of development that closely resembled the phenotypes of the dis mutants (Figure 4C); the gametes emitted a germ tube in a unipolar manner. Germination gave rise to an upright filament consisting of the cylindrical cells typical of the apical system. The gametes that showed unipolar germination occasionally produced an aborted second germ tube with an abnormally shaped cell similar to the enlarged and abnormally shaped cells that are sometimes produced by dis-1 mutants. This phenotype was not observed in the control treatments, where initial cells were incubated with siRNA molecules directed against another Ectocarpus sp gene (Ec-13_001890). These observations indicated that RNAi-induced knockdown of DIS gene expression had the same developmental consequences as the dis mutation in a proportion of the treated individuals, further confirming that Ec-05_001860 corresponds to the DIS gene. Expression Pattern of the DIS Gene during the Life Cycle RT-qPCR analysis indicated that, as expected, DIS transcript abundance was significantly reduced in both dis-1 and dis-2 mutants (Supplemental Figure 8). We used RNA-seq to assay the abundance of the DIS transcript throughout the life cycle of Ectocarpus. Specifically, we compared the expression of DIS at several stages of the ontogeny of the gametophyte and in several tissues of the sporophyte generation (Supplemental Figure 1B). This analysis indicated that transcription of DIS was upregulated in the basal system compared with the apical upright system during the sporophyte generation (pairwise analysis between apical system and basal system, fold change [FC] = 1.9; adjusted P value [Padj] = 7.66E−06). The DIS transcript was detected at low abundance during the gametophyte generation (which consists predominantly of apical filament cells), similar to the level detected in the apical system of the sporophyte (Supplemental Figure 8). Taken together, the expression pattern of DIS during the life cycle of Ectocarpus is consistent with a role in the regulation of the basal system, both during the gametophyte and the sporophyte generations. DIS Encodes the Conserved Protein TBCCd1 The DIS gene is predicted to encode a 774-amino acid protein with a TBCC domain (accession number PF07986 in Pfam database; Supplemental Table 5). Three classes of TBCC proteins have been described in other eukaryotic lineages: canonical TBCC, retinitis pigmentosa protein 2 (RP2), and TBCCd1. Phylogenetic analysis indicated that DIS encodes a TBCCd1 protein (Figure 4D; Supplemental Figure 9 and Supplemental Table 6). DIS is the only TBCCd1-encoding gene in the Ectocarpus genome. Note that the stop codon in dis-2 is located within the region encoding the TBCC domain (Figure 4B). Canonical TBCC functions together with several other tubulin cofactors/chaperones in αβ-tubulin assembly and therefore plays a key role in microtubule dynamics (Tian et al., 1996; Nithianantham et al., 2015). RP2 is also thought to be involved in microtubule assembly but the role TBCCd1 is less clear. Phenotypic analyses have shown that TBCCd1 plays an important role in positioning organelles within the cell in diverse organisms (Feldman and Marshall, 2009; André et al., 2013), but the molecular mechanism through which this protein acts is unclear and may not involve direct effects on microtubule dynamics (Gonçalves et al., 2010). These previous studies prompted us to compare intracellular features of germinating dis initial cells with those of the wild type. No Defects in Flagella Structure and Function Were Detected in dis Mutants Chlamydomonas reinhardtii mutants affected in the ASQ2 gene (which encodes a TBCCd1 protein) have a variable number of flagella (Feldman and Marshall, 2009). In contrast, we observed no variation in flagella number in dis mutant individuals and both the posterior and the anterior flagella of dis gametes were morphologically similar to the wild-type equivalents and were positioned normally, being inserted asymmetrically on the “ventral” side, close to the eyespot (Maier, 1997; Supplemental Table 7). dis gametes exhibited no abnormal swimming behavior that might have been indicative of loss of flagella functionality; both wild-type and dis gametes were positively phototactic (Supplemental Table 7). Moreover, male gametes of dis and wild-type strains showed no difference in fertilization success, evidence that the former can swim toward female gametes in response to pheromone release as efficiently as wild-type gametes (Supplemental Table 7). In summary, we did not find any evidence that DIS is required for flagella functionality. Initial Cells of dis Mutants Exhibit Increased Cell Size, Modified Golgi Architecture, Abnormal Positioning of the Nucleus, and Modification of the Microtubule Network The germinating initial cells of dis mutants were significantly larger than wild-type initial cells (mean areas of 53.9 µm2, 54.1 µm2, and 32.0 µm2 for dis-1, dis-2, and the wild type, respectively; Supplemental Table 7; Figure 5D), and analysis of transmission electron microscopy (TEM) images indicated an unusually abundant trans Golgi network in the initial cells of dis mutants, suggesting a perturbation of secretory activity (Figures 5A to 5F). Moreover, Golgi cisternae of dis-1 and dis-2 cells were significantly shorter than those of the wild type (Wilcoxon test, P value = 1.22e−13; Figure 5H). A similar phenotype has been observed in TBCCd1-depleted human cells, which are enlarged and exhibit a disorganized Golgi (Gonçalves et al., 2010). The cellular phenotype of the dis mutants is therefore consistent with TBCCd1 having a conserved role in maintaining the structural integrity of the Golgi during early development across extremely distant eukaryotic groups. Note that depletion of the tubulin binding cofactor TBCE also causes disorganization and fragmentation of the Golgi (Haase and Rabouille, 2015). Figure 5. Open in new tabDownload slide Intracellular Organization of dis Cells Compared with the Wild Type. (A) and (B) TEM images of representative dis-1 and dis-2 cells, respectively, showing that there was no detachment of the centriole (C) in relation to the nucleus (N). (C) TEM image of a representative wild-type cell. (D) Cell area in germinating initial cells of the wild-type strain and the dis-1 and dis-2 mutants. Measurements of cell area (using ImageJ) were performed 36 h after spore release on n = 56 germinating initial cells. Different letters above the bars indicate significant differences (Wilcoxon test, P value = 7.425e-06 for the comparison wild type versus dis-1 and P value = 0.0017 for the comparison wild type versus dis-2). (E) to (G) The initial cells of the dis mutants present an abundant Golgi apparatus (G), with shorter cisternae, compared with wild-type initial cells. (E) and (F) TEM images of representative dis-1 and dis-2 ([E] and [F], respectively) initial cells (sporophyte generation). Note that ultrastructure is normal except that there is an abundant trans-Golgi apparatus and Golgi cisternae appear fragmented. (G) TEM image of a wild-type initial cell (sporophyte generation). Inset panels show details of Golgi structure and correspond to the boxed regions in each image. (H) Golgi cisternae are significantly shorter in dis mutants compared with wild-type cells (Wilcoxon test, P value = 1.22e-13). Different letters above the bars indicate significant differences. Chl, chloroplast; N, nuclei; v, vacuole; C, centrioles; arrowheads, microtubules; G, Golgi. TGN, trans-Golgi network. Figure 5. Open in new tabDownload slide Intracellular Organization of dis Cells Compared with the Wild Type. (A) and (B) TEM images of representative dis-1 and dis-2 cells, respectively, showing that there was no detachment of the centriole (C) in relation to the nucleus (N). (C) TEM image of a representative wild-type cell. (D) Cell area in germinating initial cells of the wild-type strain and the dis-1 and dis-2 mutants. Measurements of cell area (using ImageJ) were performed 36 h after spore release on n = 56 germinating initial cells. Different letters above the bars indicate significant differences (Wilcoxon test, P value = 7.425e-06 for the comparison wild type versus dis-1 and P value = 0.0017 for the comparison wild type versus dis-2). (E) to (G) The initial cells of the dis mutants present an abundant Golgi apparatus (G), with shorter cisternae, compared with wild-type initial cells. (E) and (F) TEM images of representative dis-1 and dis-2 ([E] and [F], respectively) initial cells (sporophyte generation). Note that ultrastructure is normal except that there is an abundant trans-Golgi apparatus and Golgi cisternae appear fragmented. (G) TEM image of a wild-type initial cell (sporophyte generation). Inset panels show details of Golgi structure and correspond to the boxed regions in each image. (H) Golgi cisternae are significantly shorter in dis mutants compared with wild-type cells (Wilcoxon test, P value = 1.22e-13). Different letters above the bars indicate significant differences. Chl, chloroplast; N, nuclei; v, vacuole; C, centrioles; arrowheads, microtubules; G, Golgi. TGN, trans-Golgi network. The microtubule network plays an important role in the positioning of cellular organelles and has been implicated in the organization and positioning of the Golgi apparatus close to the nuclei (reviewed in Rios and Bornens, 2003). We therefore investigated whether the microtubule network was altered in dis mutants. In wild-type initial cells, a well organized interphase microtubule network was visible during initial cell germination. Microtubules nucleated from the pole opposite the first germination tube and microtubule bundles were gently curved and oriented parallel to the germination axis toward the germination pole (Figures 6A and 6B; Supplemental Figure 10). In dis mutant cells, microtubule bundles were more abundant compared with the wild type (Figure 6; Wilcoxon test, P = 0.000156 and P = 0.0069 for the wild type versus dis-1 and the wild type versus dis-2 comparisons, respectively) and the architecture of the microtubule network was disturbed. dis mutant microtubules were organized in wavy bundles with a crisscross pattern (Figures 6A and 6B; Supplemental Figure 10). The perturbations of the microtubule architecture were only observed in the germinating initial cell, before the first cell division. No abnormalities of the microtubule network were detected in dis mutant filament cells at or after the two-cell stage, compared with wild-type cells (Figure 6C). Cells of wild type and dis 15 day old germlings exhibited similar microtubule networks with bundles of a variety of thicknesses. Figure 6. Open in new tabDownload slide The Organization of the Microtubule Cytoskeleton Is Affected in dis Mutant Germinating Cells. (A) to (C) Confocal maximum z-projections showing representative cells of wild-type, dis-1, and dis-2 mutants at several stages of early development 24 h (A) 48 h (B), and 15 d (C) after release of spores from plurilocular sporangia. Microtubules (MT) were immunostained with an antitubulin antibody (green). Nuclear DNA was counterstained with DAPI (mauve). Microtubule bundles were wavy and more abundant in both dis-1 and dis-2 mutant cells compared with the wild type during the germination of the initial cell ([A] and [B]). The patterns of microtubule networks in dis mutant and wild-type cells at later stages did not exhibit any marked differences (C). Note that adult filament cells exhibit more autofluorescence, and DAPI staining is therefore less clear than in germinating cells. Nuclei are highlighted with an asterisk. Cell contours are indicated with a dotted line. In (A) and (B), cells are germinating toward the top of the figure. Images in (A) and (B) are representative of 113 wild-type, 89 dis-1, and 43 dis-2 cells. Images in (C) are representative of 18 wild-type, 21 dis-1, and 10 dis-2 cells in >10-cell filaments. Bars = 2 µm. (D) The initial, germinating cells of dis mutants exhibit, on average, more microtubule bundles than wild-type initial cells. The diagram below the graph indicates the stage of growth measured and the positions of the transepts used to count the number of microtubule bundles. The number of cells measured for each strain is indicated under each sample in parentheses. (E) Confocal maximum z-projections showing representative germinating cells (used for scoring in [F]) of wild-type (n = 42) and dis-2 mutants (n = 31) immunostained with antitubulin and anticentrin (cen) antibodies and counterstained with DAPI. Centrin colocalized with the microtubule nucleation sites in wild-type and dis-2 mutant cells (representative of the pattern for both the dis-1 and dis-2 mutants). Germination direction is indicated with an arrow, and microtubule nucleation sites are highlighted with an arrowhead. Bars = 5 µm. (F) Relative positions of the microtubule nucleation sites and the nucleus in relation to the germination axis in wild-type and dis mutant initial cells. The number of cells scored is indicated in parentheses. (G) Proportion of cells containing nuclei in anterior, central, or posterior position in relation to the germination axis before the first cell division in wild-type, dis-1, and dis-2 mutant initial cells. Nuclei were mispositioned in a significant proportion of dis initial cells before the first cell division. The number of cells scored in each sample is indicated in parentheses. (H) Positions of nuclei in wild-type and dis mutants during germination, at the two- to three-cell stage and in adult filaments (>10 cells). The diagrams below the graph indicate the stage of growth measured and the two distances measured for each cell to calculate the ratio of distances. A ratio of 1 means that nuclei were centrally positioned in the cell along the elongation axis. The number of cells counted for each sample is indicated in parentheses. Figure 6. Open in new tabDownload slide The Organization of the Microtubule Cytoskeleton Is Affected in dis Mutant Germinating Cells. (A) to (C) Confocal maximum z-projections showing representative cells of wild-type, dis-1, and dis-2 mutants at several stages of early development 24 h (A) 48 h (B), and 15 d (C) after release of spores from plurilocular sporangia. Microtubules (MT) were immunostained with an antitubulin antibody (green). Nuclear DNA was counterstained with DAPI (mauve). Microtubule bundles were wavy and more abundant in both dis-1 and dis-2 mutant cells compared with the wild type during the germination of the initial cell ([A] and [B]). The patterns of microtubule networks in dis mutant and wild-type cells at later stages did not exhibit any marked differences (C). Note that adult filament cells exhibit more autofluorescence, and DAPI staining is therefore less clear than in germinating cells. Nuclei are highlighted with an asterisk. Cell contours are indicated with a dotted line. In (A) and (B), cells are germinating toward the top of the figure. Images in (A) and (B) are representative of 113 wild-type, 89 dis-1, and 43 dis-2 cells. Images in (C) are representative of 18 wild-type, 21 dis-1, and 10 dis-2 cells in >10-cell filaments. Bars = 2 µm. (D) The initial, germinating cells of dis mutants exhibit, on average, more microtubule bundles than wild-type initial cells. The diagram below the graph indicates the stage of growth measured and the positions of the transepts used to count the number of microtubule bundles. The number of cells measured for each strain is indicated under each sample in parentheses. (E) Confocal maximum z-projections showing representative germinating cells (used for scoring in [F]) of wild-type (n = 42) and dis-2 mutants (n = 31) immunostained with antitubulin and anticentrin (cen) antibodies and counterstained with DAPI. Centrin colocalized with the microtubule nucleation sites in wild-type and dis-2 mutant cells (representative of the pattern for both the dis-1 and dis-2 mutants). Germination direction is indicated with an arrow, and microtubule nucleation sites are highlighted with an arrowhead. Bars = 5 µm. (F) Relative positions of the microtubule nucleation sites and the nucleus in relation to the germination axis in wild-type and dis mutant initial cells. The number of cells scored is indicated in parentheses. (G) Proportion of cells containing nuclei in anterior, central, or posterior position in relation to the germination axis before the first cell division in wild-type, dis-1, and dis-2 mutant initial cells. Nuclei were mispositioned in a significant proportion of dis initial cells before the first cell division. The number of cells scored in each sample is indicated in parentheses. (H) Positions of nuclei in wild-type and dis mutants during germination, at the two- to three-cell stage and in adult filaments (>10 cells). The diagrams below the graph indicate the stage of growth measured and the two distances measured for each cell to calculate the ratio of distances. A ratio of 1 means that nuclei were centrally positioned in the cell along the elongation axis. The number of cells counted for each sample is indicated in parentheses. In addition to the above-mentioned flagellar phenotype, the Chlamydomonas asq2 mutant exhibits defects in centriole positioning and number (Feldman and Marshall, 2009). We examined two aspects of centriole positioning in the Ectocarpus dis mutants: the degree with which centrioles were associated with the nucleus and the positioning of the associated centriole and nucleus with respect to the germination axis. Regarding the first aspect, centrioles were positioned close to the nuclei in both wild-type and dis mutant cells, and there was no evidence of detachment of centrioles from the nuclear envelope in dis mutants (Figures 5A to 5C). In contrast, we detected marked differences between wild-type and dis mutant cells with respect to the position of the centrioles in relation to the germination pole. In wild-type cells, centrioles (which colocalize with microtubule nucleating sites; Figure 6E) were located close to the nucleus on the side opposite the germination pole (distal), whereas in dis mutant cells, we observed microtubule nucleation both laterally and on the proximate side of the nucleus (Figure 6F). More than 200 dis mutant germlings were inspected at the two-cell stage to determine whether the aberrant positioning of microtubule nucleation sites in germinating initial cells leads to defects in the plane of cell division when these cells divides. This analysis failed to detect any individuals showing signs of aberrant patterns of cell division, i.e., there were no individuals in which the new septum was laid down obliquely, in a division plane that was not perpendicular to the growth axis. We also analyzed the position of the nucleus within the germinating cell. In the majority of wild-type germinating initial cells, the nucleus was positioned near the pole opposite the first germination tube (Figure 6E). In contrast, the majority of the nuclei in dis-1 and dis-2 initial cells were positioned centrally and in up to 30% of the cells the nucleus was located at the germinating end, a configuration that was only observed very rarely (∼2%) in wild-type germinating initial cells (Figures 6E and 6F). Analysis of individuals from the two-cell stage onwards did not detect any aberrations in the positioning of the nuclei in dis mutants compared with the wild type (Wilcoxon test, P value = 0.837 and P = 0.207 for the wild type versus dis-1 and the wild type versus dis-2, respectively), indicating that the phenotype observed in initial cells is limited to that stage of development (Figure 6E). Taken together, these analyses indicate that loss of the DIS protein leads to several defects in subcellular architecture of the initial cell and that these defects are associated with alterations in the microtubule network. To further examine the role of microtubules during early development of Ectocarpus initial cells, we analyzed the effect of drugs that disrupt both microtubule polymerization and depolymerization (although note that the increased number of microtubule bundles in dis mutants is more consistent with a depolymerization defect). Oryzalin, which induces microtubule depolymerization, significantly inhibited elongation of wild-type initial cells and delayed the first cell division but did not result in perturbations of the microtubule network resembling those observed in the dis mutants (Supplemental Figure 11 and Supplemental Table 8). Furthermore, oryzalin treatment did not lead to aberrant positioning of the nuclei, compared with wild-type untreated control cells (Wilcoxon test, W = 101.5, P value = 0.3818; Supplemental Figure 11). Similarly, treatment of wild-type initial cells with taxol, a microtubule stabilizing drug, inhibited germination and cell division but did not produce a cellular phenotype similar to that of the dis mutants. Taxol-treated cells had thick microtubule bundles that were mostly oriented parallel to the cell membrane (Supplemental Figures 10 and 11). This pattern differed from those of both untreated wild-type cells and dis cells. Taxol treatment did not have any significant effect on the position of the nucleus in germinating cells (Wilcoxon test, W = 95, P value = 0.1978; Supplemental Figure 11D). The lack of any detectable effect of pharmacological perturbation of the microtubule network on the positioning of the nucleus within the initial cell suggests that alternative cytoskeletal components, such as actin filaments, may be involved in positioning the nucleus, as observed in plant root hair cells (Ketelaar et al., 2002). Brefeldin A (BFA) has been shown to disrupt the Golgi and to inhibit polarization of the germinating Fucus embryo (Bisgrove and Kropf, 2001; Hable and Kropf, 1998). Treatment of Ectocarpus sp initial cells for 24 h with BFA weakened the cell wall (leading to the release of cytosolic material) and inhibited cell elongation, but did not reproduce the Dis− phenotype (Supplemental Table 8 and Supplemental Figure 11). Taken together, the results of these pharmacological experiments suggest that the phenotypes associated with the dis mutations are not simply due to defects in microtubule polymerization or depolymerization nor to an overall deficiency in secretion. DISCUSSION DIS Is Required for the Establishment of Basal Structures in Ectocarpus Analysis of the phenotypes of the two dis mutants described in this study indicated that the DIS gene is required for the development of basal structures throughout the Ectocarpus life cycle. In wild-type Ectocarpus gametophytes, the initial cell divides asymmetrically to produce two cells with apical and basal identities. Further divisions in opposite directions of each of these cell types generate an apical thallus and a basal rhizoid, respectively. In the gametophytes of dis mutants, the basal germ tube that gives rise to the rhizoid is not produced. The consequent unipolar germination gives rise to individuals with an apical system but no basal structures. We suggest that the basal system is not produced because the dis mutants fail to deploy a putative basal determinant factor (“B”) in the initial cell; as a result, the first division produces an apical cell and a quiescent undifferentiated cell (Figure 7). Figure 7. Open in new tabDownload slide Proposed Model for the Action of the DIS Protein during the Establishment of the Apical/Basal Axis in Ectocarpus sp. In the wild-type gametophyte, axis formation involves the asymmetric distribution of a putative apical determinant “A” (green dots) and of a putative basal determinant “B” (red dots), conferring apical (green shading) and basal (red shading) identities on the two daughter cells, respectively. In the wild-type sporophyte, the (dominant) B factor is expressed at a higher level resulting in two basal cells, which then divide to produce the basal structure. In dis mutants (both gametophyte and sporophyte generations), the B factor is not deployed and the first cell division gives rise to an apical cell (green) and a quiescent, undifferentiated cell (gray). Only the apical cell is capable of growth, resulting in unidirectional growth and a single germ tube. Note that the proposed A factor is necessary to explain the unipolar growth of the dis mutants in the absence of the proposed B factor. Figure 7. Open in new tabDownload slide Proposed Model for the Action of the DIS Protein during the Establishment of the Apical/Basal Axis in Ectocarpus sp. In the wild-type gametophyte, axis formation involves the asymmetric distribution of a putative apical determinant “A” (green dots) and of a putative basal determinant “B” (red dots), conferring apical (green shading) and basal (red shading) identities on the two daughter cells, respectively. In the wild-type sporophyte, the (dominant) B factor is expressed at a higher level resulting in two basal cells, which then divide to produce the basal structure. In dis mutants (both gametophyte and sporophyte generations), the B factor is not deployed and the first cell division gives rise to an apical cell (green) and a quiescent, undifferentiated cell (gray). Only the apical cell is capable of growth, resulting in unidirectional growth and a single germ tube. Note that the proposed A factor is necessary to explain the unipolar growth of the dis mutants in the absence of the proposed B factor. In the wild-type sporophyte, the first division of the initial cell is symmetrical and the two germ tubes form the two ends of a basal filament, which is strongly attached to the substratum. In the absence of the DIS protein, the initial cell produces just one germ tube, which gives rise to the apical system. We interpret the failure to initiate a second germ tube as being a consequence of the failure to establish basal cell identity in the sporophyte of this mutant. We suggest that, in the absence of the basal determinant factor B, the initial cell divides to produce an apical cell and a quiescent undifferentiated cell in the same manner as observed in the gametophyte generation. The dis-2 mutation creates a premature stop codon within the region that encodes the TBCC domain and is therefore likely to be a null mutation. The dis-1 mutation, on the other hand, is located in an intron (Figure 3). This mutation causes a reduction in transcript abundance but does not modify the open reading frame of the transcript. Analysis of the RNA-seq data did not detect any evidence that the mutation affects splicing of the transcript. Therefore, dis-1 may not represent a null mutation. If dis-1 is not a null allele, then under the model shown in Figure 7, the abortive structures produced instead of the second germ tube in dis-1 mutant sporophytes can be interpreted as arising due to incomplete penetrance of the dis-1 mutation, allowing a small amount of the putative basal determinant B to accumulate and the production of an abortive basal system. The model proposes that the observed phenotypes are due to variations in concentration, rather than the subcellular distribution, of the putative B factor. One possible scenario would be that the abnormal Golgi apparatus of the initial cell is unable to produce sufficient amounts either of the putative B factor itself or of a protein that generates the B factor enzymatically. Interestingly, comparison of the dis-1 transcriptome with those of wild-type apical and basal tissues identified an enrichment in genes coding for secreted proteins and highlighted a putative role for DIS in, among other processes, cell wall modification processes, secretion, transport, and vesicle trafficking. Secretion and diffusion of cell fate determinants, particularly from the cell wall, have been shown to play a crucial role in pattern formation in brown algae (Bouget et al., 1998; Arun et al., 2013). Interestingly, mutant analysis indicates that cell wall synthesis, cell wall integrity sensing, and vesicle trafficking are also important processes during the development of Marchantia polymorpha rhizoids and Arabidopsis root hairs (Honkanen et al., 2016). In Ectocarpus, the apical and basal developmental programs appear to be independent, in the sense that removal of the basal system in the dis mutants had no visible effect on patterning of the apical system. In contrast, Arabidopsis mutations that result in the loss of basal structures, such as monopteros and bodenlos for example, also tend to cause more or less subtle modifications to the apical structures (Hardtke and Berleth, 1998; Hamann et al., 1999). This difference may reflect the simpler developmental program of Ectocarpus compared with Arabidopsis but may also be related to the fact that the primary function of brown algal basal systems is to act as a holdfast, whereas the aerial tissues of land plants are dependent on the root system not only for anchorage but also for water and nutrient supply. In land plants, therefore, there may be more of a tendency for the developmental programs of the apical and basal systems to be interdependent. DIS Encodes a TBCC Domain Protein with a Conserved Role in Intracellular Organization The DIS gene encodes a TBCCd1 protein. TBCCd1 is a member of the TBC group of proteins, which also includes the RP2 protein and TBCC, a component of the chaperone complex (TBCA to TBCE) that catalyzes the formation of α,β-tubulin heterodimers (Tian and Cowan, 2013; Steinborn et al., 2002). All three TBCC proteins have been strongly conserved across the eukaryotic tree (Figure 4D; Supplemental Table 5), suggesting that each has a specific function in the cell. However, while the role of TBCC has been described in detail, the exact cellular and molecular function of TBCCd1 remains unclear. The role of TBCC in the α,β-tubulin heterodimerization pathway is to trigger GTP hydrolysis catalyzing the release of newly formed αβ-tubulin heterodimers. Importantly, this pathway is reversible so that TBCC plays a role in both microtubule assembly and disassembly (Nithianantham et al., 2015). Both TBCC and RP2 have GAP activities, mediated by their TBCC domains, and RP2 can complement a TBCC mutant (Schwahn et al., 1998; Schwarz et al., 2012). In contrast, the TBCC domain of TBCCd1 lacks several conserved residues including a critical arginine residue and TBCCd1 is thought not to be a GAP protein (Bartolini et al., 2002; Scheffzek et al., 1998), although it has been suggested that a nearby conserved arginine residue may substitute for the lost arginine residue (Feldman and Marshall, 2009) (Supplemental Figure 9). Therefore, although TBCCd1 probably interacts with the cytoskeleton in some way to mediate its cellular functions, it is currently unclear whether this specifically involves a direct role in microtubule polymerization/depolymerization. TBCCd1 has been localized to the centrosome and the Golgi in organisms as diverse as humans, Chlamydomonas, and trypanosomes, and loss-of-function experiments have indicated a conserved and important role for TBCCd1 in various aspects of the organization of cellular architecture in these diverse eukaryote groups (André et al., 2013; Feldman and Marshall, 2009; Gonçalves et al., 2010). Loss or knockdown of TBCCd1 has severe consequences for cell morphology, for example, leading to alterations in the shape and motility of human cells (Gonçalves et al., 2010), mitotic defects in trypanosomes (André et al., 2013), and defects both in flagella number and functioning and during cell division due to misorientation of the mitotic spindle in Chlamydomonas (Feldman and Marshall, 2009). Although we did not detect any defects in flagella structure or centriole positioning in Ectocarpus dis mutants, the initial cells of these mutants did show defects in the positioning of the nucleus and an atypical microtubule network during germination. Moreover, dis initial cells were markedly larger than the wild-type equivalents and their Golgi network was more abundant and fragmented (shorter cisternae). Increased cell size and disorganization of the Golgi apparatus have been observed in TBCCd1-depleted human and trypanosome cells (Gonçalves et al., 2010; André et al., 2013). The role of TBCCd1 in regulating organelle positioning and microtubule and Golgi architecture, highlighted by our study, can therefore be traced back to the crown divergence of the major eukaryotic groups, more than a billion years ago (Eme et al., 2014). The Golgi defects in the dis mutants are particularly interesting with regard to the model described in Figure 7, which proposes that the Dis− phenotype is due to failure to deploy a hypothetical basal-cell-fate-determining factor B during the first cell division. In animals, the Golgi is critical for cellular differentiation and morphogenesis because it spatially constrains developmental pathways, and it is required for the establishment of cell polarity and normal subcellular organization (Copeland et al., 2016; Vinogradova et al., 2009; Zhong, 2011). Moreover, in the embryo of the brown alga Fucus, targeted secretion from the Golgi to the cell wall has been implicated in axis fixation and establishment of the basal versus the apical system (Shaw and Quatrano, 1996). In Arabidopsis, asymmetrical division of the zygote is preceded by reorganization of the cytoskeleton and repositioning of the nucleus from a central position to the apical region of the cell (Kimata et al., 2016; Ueda et al., 2011). Mutation of the zinc finger transcription factor gene WRKY2 not only disrupts the asymmetrical division of the zygote cell but also leads to partial loss of basal identity in the basal cell lineage (Ueda et al., 2011). Interestingly, the Ectocarpus dis mutants not only fail to produce a basal system but also exhibit disrupted organization of the microtubule cytoskeleton and defects in nucleus positioning in the initial cell. This suggests that, as in flowering plants, the establishment of the basal cell lineage in brown algae depends critically on events leading up to and including the first division of the initial cell. It is not clear how loss of the TBCCd1 protein leads to disorganization of the microtubule cytoskeleton in the dis mutants (whether this is a direct or an indirect effect for example), but this phenotype does not appear to be simply due to defects in microtubule polymerization or depolymerization because treatment with oryzalin or taxol did not result in a phenotype that resembled that of the dis mutants. The observed modifications to the microtubule architecture in dis mutants may be due to more subtle changes in nucleating activity, rates of assembly and disassembly of bundles, modifications of microtubule dynamic instability and/or tubulin, or microtubule posttranslational modifications (Mitchison and Kirschner, 1984; Song and Brady, 2015). Based on the phenotypes associated with TBCCd1 loss in other eukaryotes and on the phenotypes of the dis mutants, we propose that acquisition of basal cell identity in Ectocarpus depends critically on features of the initial cell architecture that are disrupted in the absence of a functional DIS/TBCCd1 protein. Further work is required to understand the exact cellular role of TBCCd1 in Ectocarpus but the identification of this component will allow new insights into the cellular mechanisms involved in establishing apical/basal identity. DIS and the Evolution of Sporophyte and Gametophyte Developmental Programs The pattern of early development of the Ectocarpus sporophyte, which involves deployment of an extensive basal system before the establishment of the apical/basal axis, is unusual because most brown algal sporophytes exhibit early establishment of the apical/basal axis leading directly to the production of a thallus cell and a rhizoid, respectively (Fritsch, 1935). In the Ectocarpus imm mutant the extensive basal system of the sporophyte fails to form and this structure is replaced by a rhizoid (Peters et al., 2008). Based on the phenotype of the imm mutant, we have suggested that the pattern of early sporophyte development in extant Ectocarpus evolved from a simpler, ancestral developmental program that more closely represented that of the gametophyte, i.e., involving an asymmetrical initial cell division that gave rise directly to apical and basal organs (Macaisne et al., 2017). The dis mutants provide further support for this hypothesis because they exhibit complete loss both of the rhizoid during the gametophyte generation and of the extensive network of prostrate filaments during the sporophyte generation, suggesting that the two structures are indeed equivalent. Moreover, in a dis-1 imm double mutant sporophyte, the rhizoid that normally replaces the prostrate filament system fails to develop, providing further evidence that the rhizoid that forms in the imm mutant sporophyte is developmentally equivalent to the basal prostrate filament system of the wild type sporophyte. In land plants, which also have haploid-diploid life cycles, there has been considerable interest in the evolutionary origins of the sporophyte and gametophyte developmental programs, specifically whether each generation has independently evolved its own developmental pathways or, alternatively, whether there has been recruitment of developmental programs from one generation to the other during evolution (Dolan, 2009; Pires and Dolan, 2012; Shaw et al., 2011). Current evidence indicates that the developmental networks that implement land plant sporophyte programs were mainly recruited from the gametophyte generation, which was initially the dominant generation in the land plant lineage (Dolan, 2009; Niklas and Kutschera, 2010), but there is also evidence that there have been sporophyte-specific innovations (Szövényi et al., 2011; Sano et al., 2005). Analyses of the imm (Macaisne et al., 2017) and dis (this study) mutants suggest that the evolution of developmental systems in the brown algae also involved both co-opting of programs from one generation to the other and generation-specific developmental innovations. The phenotypes of the dis mutants suggest that, despite their clear morphological differences, the basal systems of the sporophyte and gametophyte generations share underlying mechanistic features because the TBCCd1 protein is necessary for the deployment of both types of basal structure. In this respect, therefore, the sporophyte and gametophyte basal structures appear to be homologous. On the other hand, it is only in the sporophyte that the rhizoid has been modified to produce an extensive system of prostrate filaments (note that gametophytes carrying the imm mutation are indistinguishable from wild type gametophytes; Peters et al., 2008). The developmental program that depends on the IMM protein therefore appears to have been a sporophyte-specific innovation, which was presumably built onto a more ancient program shared with the gametophyte generation that requires the action of the DIS gene. METHODS UV Mutagenesis and Isolation of Mutant Strains Strain cultivation, genetic crosses, raising of sporophytes from zygotes, and isolation of meiotic families were performed as described previously (Coelho et al., 2012a, 2012d). Gametes of Ectocarpus are able to develop parthenogenically to produce haploid partheno-sporophytes, which are identical morphologically to the sporophytes that develop from diploid zygotes (Peters et al., 2008; Coelho et al., 2011). This phenomenon was exploited to screen directly, in a haploid population, for mutants affected in early sporophyte development. UV mutagenesis of gametes was performed as described previously (Godfroy et al., 2015) and mutant partheno-sporophytes lacking basal structures were identified by visual screening under a light microscope. Genetic Mapping of the DIS Locus To obtain an approximate map position for the DIS gene, the dis-1 mutant (Ec722) was crossed with the outcrossing line Ec568 to generate a mapping population of 265 individuals. Each of the 265 individuals was derived from a different unilocular sporangium (each unilocular sporangium contains 50–100 meiospores, derived from a single meiosis followed by at least five mitotic divisions). The meiospores germinated to produce gametophytes, which were isolated and allowed to germinate parthenogenically. The resulting partheno-sporophytes were then observed under a light microscope to determine whether they exhibited the Dis− phenotype. The dis-1 mutation was then approximately mapped using 21 Dis− individuals from this segregating population and 74 microsatellites markers from the Ectocarpus microsatellite-based genetic map (Heesch et al., 2010). Using this approach, we localized the dis-1 mutation to chromosome 05 at 9.5 cM from the M_114 marker. A cloning-by-sequencing strategy (using the SHOREmap approach; Schneeberger et al., 2009) was then used to precisely localize the dis-1 mutation. Genomic DNA was extracted from 87 wild-type and 96 dis individuals from the mapping population using NucleoSpin 96 Plant II (Macherey-Nagel) and pooled into wild-type and Dis− pools. An Illumina HiSeq 2500 platform (Fasteris) was the used to generate 44.9 Gb of sequence data, corresponding to 117 and 104 million 2 × 100-bp paired-end reads for the wild-type and Dis− pools, respectively. The same DNA extraction protocol and sequencing platform were employed to sequence the dis-2 mutant strain Ec799, producing 49 million 2 × 125 paired-end reads (SRA accession number SRR3714421). Reads were cleaned to remove nucleotides with quality scores of less than 20 from both ends using Prinseq (Schmieder and Edwards, 2011). Reads were then only retained if they were longer than 50 nucleotides, had a mean quality of at least 25, and no non-determined nucleotides. About 3.6% and 4.3% of the raw wild-type and dis reads were removed, respectively. Bowtie2 (Langmead and Salzberg, 2012) was used to map 71.9% and 74.7% of the wild-type and dis cleaned reads, respectively, onto the Ec32 reference genome. The read mapping was then improved by read realignment and base quality recalibration using the GATK software suite. Variant calling was performed using the “consensus” tool of the SHORE pipeline in order to analyze allele frequency. Since we had only one reference genome sequence for Ectocarpus, corresponding to the wild-type strain that was mutagenized to generate the dis mutants, we were not able to apply the strategy described in the SHOREmap publication to generate the list of markers (association of SHOREmap extract and create). We therefore selected genetic markers by comparing singe nucleotide polymorphisms (SNPs) called in wild-type and Dis− pools and extracted SNPs that segregated as expected, i.e., common SNPs for which the sum of the wild-type and dis-1 allele frequencies was between 0.8 and 1.2. In order to prevent loss of markers close to the mutation, we also included SNPs that were specific to one of each of the pools with a frequency of more than 0.9 and less than 0.1. Using this marker list, and crossing SHOREmap annotation results with manual selection of mutations from SHORE consensus, SHORE qVar, and samtools/VarScan (Koboldt et al., 2013; Schneeberger et al., 2009) identified a unique SNP specific to the dis-1 pool at position 50,914 of scaffold sctg_184 (position 3,330,523 on chromosome 5). This A-to-G transition mutation was located in intron 14 of the DIS gene 15 bp after the end of exon 14. A CAPS marker (Supplemental Table 2), developed based on the single nucleotide polymorphism at position 3,330,523 of chromosome 05, was used to genotype the 265 individuals of the mapping population (which included 90 wild-type and 175 dis-1 individuals). This analysis confirmed that the presence of the A-to-G transition was 100% correlated with the Dis− phenotype in this large family. In addition, PCR amplification and Sanger sequencing of the genomic region containing the mutation confirmed its presence in the dis-1 mutant (Ec722) and its absence in the Ec32 and Ec568 strains (the original strain used for the mutagenesis and the outcross female line used to generate the segregating population, respectively). Resequencing of the DIS gene in the dis-2 mutant identified a point mutation (a C-to-T transition) in exon 13 that results in the introduction of a stop codon into the coding region of the gene (chr05, position 3,332,908). Genetic Interaction between DIS and IMM An imm mutant female (Ec602) was crossed with a dis-1 male (Ec722), and the resulting sporophyte (Ec649) gave rise to a 40-gametophyte progeny, all derived from independent meiotic events. The phenotypes of the progeny were scored under an inverted microscope, and genotypes were assessed using CAPS markers for IMM and DIS loci (Supplemental Table 2). RNAi Knockdown of DIS Expression Small interfering RNAs (siRNAs) directed against the DIS gene transcript were designed using version 3.2 of E-RNAi (Horn and Boutros, 2010). The specificity of the designed siRNAs was determined by comparing the sequence (BLASTn) with complete genome and transcriptome sequences. Candidates that matched, even partially, genomic regions or transcripts in addition to DIS were rejected. Three siRNAs with predicted high specificity corresponding to different positions along the DIS transcript were selected (Supplemental Table 9). The control siRNA was directed against the Ectocarpus sp gene Ec-13_001890, which is located within the sex-determining region of the male sex chromosome and therefore not an essential gene for sporophyte function. siRNAs were introduced into Ectocarpus sp strain Ec32 gametes using the transfection reagent HiPerFect (Qiagen). One microliter each of 0.5 μg/μL solutions of siRNAs in 1× Universal siMAX siRNA Buffer (MWG Eurofins) was mixed with 12 μL of HiPerFect transfection reagent in a final volume of 100 μL of natural seawater, vortexed to mix, and incubated for 10 min at room temperature before being added dropwise to 100 μL of freshly released gametes in natural seawater in a Petri dish. After rotating gently to mix, the Petri dish was incubated overnight at 13°C. The following day, 10 mL of culture medium was added (Coelho et al., 2012c) and incubation continued at 13°C. RNAi-induced phenotypes were observed under a light microscope and the number of individuals that resembled the dis mutant were scored in at least 400 individuals for three experimental replicates. Control treatments were performed in the same manner using siRNAs directed against the endogenous gene Ec-13-001890. Unfortunately, it is currently not possible to determine whether the siRNA treatment reduces DIS transcript abundance because, based on the observed phenotypes, gene expression is affected in only a small percentage of the treated cells. However, the use of siRNAs directed against the endogenous gene Ec-13-001890 showed that the effect was sequence specific. It would therefore seem highly unlikely that the observed phenotypes were due to processes that did not involve interference with the expression of the DIS gene. Comparative Transcriptome Analyses RNA-seq analysis was performed to compare the abundances of gene transcripts in the dis-1 mutant with those in basal and upright filaments of the wild-type sporophyte generation. Duplicate synchronous cultures (with more than 100 individuals each) were prepared for each sample under standard conditions (Coelho et al., 2012c). Upright filaments of adult (4 weeks old) wild-type partheno-sporophytes were dissected from prostrate, basal filaments using the sharp end of a Pasteur pipette under a binocular microscope. Visual inspection under the microscope ensured the absence of contaminating basal filaments. Wild-type basal filaments were obtained by harvesting immature individuals before the emergence of the upright filaments, 2 weeks after initial cell release from plurilocular gametangia. The dis-1 mutant was grown under the same conditions as the wild-type tissue and was similarly used at 2 weeks after release of initial cells from plurilocular gametangia. Tissue samples were rapidly frozen in liquid nitrogen and processed for RNA extraction. Total RNA was extracted from each sample using the Qiagen Mini kit as previously described (Lipinska et al., 2015). For each replicate, cDNA was produced by oligo(dT) priming, fragmented, and prepared for 2× 100-bp paired-end sequencing on an Illumina HiSeq 2000 platform by Fasteris. TopHat (v2.0.8) was used to map the RNA-seq reads to the reference genome. Supplemental Table 10 provides details of the sequencing, mapping statistics and accession numbers. Pairwise differential expression analysis between dis-1, basal, and upright filaments was performed with the DESeq2 package (Bioconductor) using an adjusted P value cutoff of 0.1 and a minimal FC of 2. Analysis of Predicted Gene Functions InterProScan (Zdobnov and Apweiler, 2001) and Blast2GO (Conesa and Götz, 2008) were used to recover functional annotations for Ectocarpus proteins. For Blast2GO, a Fisher exact test with a P value cutoff of 0.05 was used to detect enrichment of specific GO terms in the various groups of genes. KEGG pathway enrichment was analyzed using the KOBAS 2.0 platform (Xie et al., 2011). Signal peptides were predicted using Hectar (Gschloessl et al., 2008) implemented through the Galaxy platform (http://webtools.sb-roscoff.fr/). A gene-by-gene manual curation was used to associate each gene in the differential transcriptomic data sets to a functional category. Enrichment in specific functional categories in relation to the Ectocarpus sp genome was performed by statistical analysis using the χ2 test (P < 0.05). Phylogenetic Analysis of TBCC Proteins A multiple alignment of the TBCC domains of diverse TBCC, RP2, and TBCCd1 proteins was generated with Muscle (http://www.ebi.ac.uk/Tools/msa/muscle/). Based on this alignment, an unrooted maximum likelihood phylogenetic tree was built with the LG+G model using MEGA7 (http://www.megasoftware.net/mega.php) and 1000 bootstrap replicates. Model testing was performed in MEGA7. The coordinates of the TBCC domains within the protein sequences were determined using the NCBI Conserved domains tool (http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi). Accession numbers and domain coordinates are provided in Supplemental Table 6. Microdissection and Regeneration of dis Mutants and Wild-Type Filaments Microdissection was performed with the sharp end of a Pasteur pipette under an inverted microscope. After dissection, the material was kept at 13°C under standard culture conditions (Coelho et al., 2012e). Wild-type and dis sporophyte apical filament cells were isolated by microdissection 7 d after the emergence of the first upright filaments (20 d after gamete release); gametophyte apical filament cells were isolated by microdissection 15 d after meiospore release. Morphometric Analysis of Germinating Partheno-Sporophytes Germinating initial cells of dis-1, dis-2, and wild type partheno-sporophytes 36 h after release from gametangia were photographed under an upright microscope (BX41; Olympus). Cell areas were measured for a total of 57 cells using ImageJ (http://imagej.nih.gov/ij). Morphological measurements and analysis of images of adult filaments were also performed under an inverted microscope (CKX41; Olympus) using ImageJ (http://imagej.nih.gov/ij/, 1997-2016). Between 25 and 100 adult filament cells were measured in each of eight different wild-type or dis-1 and dis-2 individuals. The statistical analysis was performed in R. Electron Microscopy Analysis of Cellular Ultrastructure Medium containing mature Ectocarpus dis partheno-sporophytes was pipetted onto a plastic film (gel support films; ATTO). The film was cut into <1-cm side triangles, and these were attached to Petri dishes by adhesive tape. Two days after the release of mitospores from plurilocular sporangia, the resulting germlings, which were attached to the triangles, rapidly immersed in liquid propane cooled to −180°C by liquid nitrogen, and immediately transferred into liquid nitrogen. The samples were submerged in substitution solution containing 2% osmium tetroxide with acetone at −80°C for 2 d, at −40°C for 2 h, and at 4°C for 2 h. Finally, the temperature of the samples was gradually allowed to rise to room temperature, and they were then washed with acetone several times. The gel support films were infiltrated and embedded in Spurr's low-viscosity resin (Polysciences) on aluminum foil dishes. The films with the samples were turned inside out on the upper surface of the resin. Serial sections were cut with a diamond knife on an Ultracut ultramicrotome (Reichert-Jung) and mounted on Formvar-coated slot grids. Sections were stained with TI blue (Nisshin EM) and lead citrate and observed using an electron microscope (JEM-1011; JEOL). The images shown in Figure 5 are representative of 14 initial cells. Golgi fragmentation was quantified using a method adapted from (Tang et al., 2011). TEM photographs were used to measure the lengths of cisternae. Measurements were performed on 14 different individuals, and a total of 113 cisternae from dis-1, dis-2, and wild-type individuals were analyzed. Data from dis-1 and dis-2 were pooled. Cisternae were defined as long membranous structures with a length greater than twice their width, the latter not exceeding 60 nm. Normal cisternae ranged from 20 to 30 nm in width and were longer than 400 nm. Statistical significance was assessed by a Wilcoxon test, implemented in R. RT-qPCR Analysis The abundance of gene transcripts during the Ectocarpus life cycle was assessed using RT-qPCR. RT-qPCR was performed as previously described (Coelho et al., 2011; Lipinska et al., 2013). Primer pairs were designed to amplify regions of the 3′ untranslated region or the most 3′ exon of the gene to be analyzed (Supplemental Table 19). In silico virtual PCR amplifications were performed using the e-PCR program (Rotmistrovsky et al., 2004) and the male Ec32 genome sequence to check the specificity of oligonucleotide pairs. The Plant RNeasy extraction kit (Qiagen) was used to extract total RNA from at least three biological replicates for each of the stages of the life cycle: gametes, young sporophytes, immature gametophytes, mature gametophytes, partheno-sporophytes, and diploid heterozygous sporophytes. The RNA was treated with RNase-free DNase-I according to the manufacturer's instructions (Qiagen) to remove any contaminating DNA and stored at −80°C. The concentration and integrity of the RNA was checked using a NanoDrop 2000 spectrophotometer (ThermoFisher Scientific) and by agarose gel electrophoresis. A control PCR without reverse transcriptase was performed to ensure absence of contaminating DNA. For each sample, up to 1 µg of RNA was reverse-transcribed to cDNA using oligo(dT) and the Superscript II RT kit (Life Technologies) according to the manufacturer's instructions and the cDNA was diluted with water to 1.2 ng equivalent RNA⋅µL−1. cDNAs were amplified using the IQ SYBR Green supermix (Bio-Rad Laboratories) on a Chromo4 System thermocycler (Bio-Rad Laboratories). The amplification efficiency was tested using a genomic dilution series and was always at least 80%. The specificity of amplification was checked with a dissociation curve. The EF1α gene (Ec-21_002980) or the Arp2/3 gene (Ec-09_003710) was chosen as a constitutively expressed controls based on Le Bail et al. (2008), and their constitutive expression was validated by the RNA-seq data sets. The normalized data were expressed as the mean ± sd calculated from three independent biological experiments. Inhibitor Treatments Stock solutions of inhibitors were prepared in DMSO (oryzalin at 100 mM, taxol [paclitaxel] at 5 mg/mL, and BFA at 25 mg/mL) and working solutions were diluted in half strength Provasoli-enriched seawater (Coelho et al., 2012d). Freshly released Ectocarpus sp gametes were settled in Petri dishes and treated for 24 h with 0.01 to 1 µM oryzalin, 5 to 20 µM paclitaxel, or 2.5 to 25 µg/mL BFA. Controls were treated with 0.002% DMSO. Cells were allowed to develop after removal of the inhibitor and observations were performed 4 and 7 d later, using an inverted Olympus CKX41 microscope. For each experiment, between 19 and 310 individuals were counted in triplicate dishes (529, 1071, and 2142 cells scored in total in each of the oryzalin, paclitaxel, and BFA treatments, respectively; Supplemental Table 16). Immunostaining Ectocarpus samples were processed as described by Coelho et al. (2012b) using an adapted protocol from Bisgrove and Kropf (1998). Briefly, Ectocarpus cells were settled on cover slips and at appropriate times after settlement were rapidly frozen in liquid nitrogen and fixed in 2.5% glutaraldehyde and 3.2% paraformaldehyde for 1 h, then washed in PBS and treated with 5% Triton overnight. Samples were then rinsed in PBS and 100 mM NaBH4 was added for 4 h. Cell walls were degraded with cellulase (1% w/v) and hemicellulase (4% w/v) for 1 h, and the preparation was then rinsed with PBS and blocked overnight in 2.5% nonfat dry milk in PBS. Samples were treated with antitubulin antibody (1/200th, DM1A; Sigma-Aldrich) at 20°C overnight and then treated with the secondary antibody (AlexaFluor 488-conjugated goat anti-mouse IgG; Sigma-Aldrich; 1:1000 in PBS) at 20°C overnight. The preparation was rinsed with PBS and blocked overnight in 2.5% nonfat dry milk in PBS and then treated with anticentrin antibody (1/1000th anticentrin 1 ab11257; Abcam) at 20°C overnight, followed by the secondary antibody (1/1000th AlexaFluor 555-conjugated goat anti-rabbit IgG; Sigma-Aldrich) for 8 h. Samples were stained with 4′,6-diamidino-2-phenylindole (DAPI; 0.5 µg/mL in PBS) for 10 min at room temperature and mounted in ProLong Gold (Invitrogen). Confocal Microscopy Confocal microscopy was conducted using an inverted SP8 laser scanning confocal microscope (Leica Microsystems) equipped with a compact supply unit which integrates a LIAchroic scan head, several laser lines (405 and 488 nm), and standard photomultiplier tube detectors. We used the oil immersion lens HC PL APO 63×/1.40 OIL CS2. The scanning speed was set at 400 Hz unidirectional. The pinhole was adjusted to one Airy unit for all channels. The spatial sampling rate was optimized according to Niquist criteria, generating a 0.058 × 0.058 × 0.299-µm voxel size (xyz). The Z-stack height fitted the specimen thickness. A two-step sequential acquisition was designed to collect the signal from three or four channels. The first step recorded the antitubulin fluorescence signal (excitation, 488 nm; emission, 530 nm) and the transmitted light. The second step acquired the DAPI fluorescence signal (excitation, 405 nm; emission, 415–480 nm) and the anticentrin signal (excitation, 552 nm; emission, 560–590 nm). Signal intensity was averaged three times. The Fiji software was used to optimize the raw images, including maximum intensity projection and denoising (3*3 median filter). For any given data, both wild-type and mutant images were analyzed simultaneously with similar settings. The Fiji software was also used to estimate the number of microtubule bundles in each cell, before the first cell division. Tracking of bundles was performed on maximum intensity projections of z-planes covering the whole thickness of the cells. We drew three lines transversely, perpendicular to the growth axis of the cell: one in the center of the cell and other two half way between the center and the cell extremities (see diagram in Figure 6D). Peaks corresponding to the microtubule bundles were then identified in plots of intensity profiles at each of the three positions in the cell and counted. The three values were averaged for each cell to derive an estimation of the number of microtubule bundles in each cell. Note that in the dis mutants, due to the disorganized nature of the microtubule network, average bundle numbers may be somewhat underestimated. This is because this method is well adapted for tracking microtubule bundles oriented with their long axis parallel to the image plane, but we may have missed bundles that were perpendicular to the plane of the transection. Measurement of the Positions of Nuclei and Microtubule Nucleation Sites Nuclei were stained in vivo with Hoechst 33342 (ThermoFisher), imaged with an Olympus BX microscope with a ×40 objective, and distances were measured using Fiji software. In germinating initial cells, the position of the nucleus was calculated by measuring the distance from the nucleus to the germinating pole (i.e., the end corresponding to the emerging germ tube) and the distance from the nucleus to the opposite pole of the cell and dividing the latter by the former. Nuclei were scored as being anterior (located near the emerging germ tube), central, or posterior (located nearer the pole opposite the germination pole). At the two-cell stage, the positions of nuclei were similarly measured in a directional manner by measuring the distance from the nucleus to the cell boundary at the germinating end of the filament and the distance from the nucleus to the opposite pole of the cell and dividing the latter by the former. For cells in multicellular filaments at a later stage of development, it was not possible to assign a direction of growth. Filaments were therefore orientated arbitrarily and the position of each nucleus was calculated by dividing the distance from one end of the cell (x) by the distance from the other end of the cell (y) as illustrated in Figure 6H. Microtubule nucleation site positions were measured by dividing each cell into four diagonal quadrants and scoring the position of the nucleating sites with respect to these quadrants: proximal (located on the same pole as the emerging germ tube), lateral, or distal (located nearer the pole opposite the germination pole) as illustrated in Figure 6F. Accession Numbers Accession numbers are provided in Supplemental Tables 6 and 10. DNA-seq data are uploaded at the NCBI Sequence Read Archive (SRA) under the following accession numbers: SRR3710253 (wild-type pool), SRR3710254 (Dis− pool), and SRR3714421 (dis-2 mutant). Supplemental Data Supplemental Table 1. List of strains used in this study. Supplemental Table 2. Genetic analysis of the progeny from a cross between an imm and a dis mutant. Supplemental Table 3. Morphometric analysis of filament cells of adult wild-type and dis individuals. Supplemental Table 4. Functional categories associated with the genes from the several transcriptomic data sets. Supplemental Table 5. TBC group proteins and components of the prefoldin and chaperonin-containing TCP1 complex in Ectocarpus sp. Supplemental Table 6. TBCC domain protein sequence coordinates used to generate the phylogenetic tree. Supplemental Table 7. Cellular features of early development in wild-type and mutant strains. Supplemental Table 8. Effect of oryzalin, taxol, and brefeldin A treatments on wild-type and dis initial cells. Supplemental Table 9. siRNA molecules used for the RNA interference experiments. Supplemental Table 10. RNA-seq data statistics and accession numbers. Supplemental Table 11. Primer pairs designed to assay transcript abundance for generation-specific genes by RT-qPCR. Supplemental Data Set 1. DESeq2 differentially expressed genes, FC > 2, Padj < 0.05, genes differentially expressed in at least one comparison (dis-1 versus basal system versus apical system). Supplemental Data Set 2. Gene Ontology (GOSlim) analysis of genes in differentially regulated gene clusters 1 and 2 (see also Figure 3). Supplemental Data Set 3. The 100 most differentially expressed genes in pairwise comparisons between the dis-1 mutant and wild-type apical or basal tissues. Supplemental Data Set 4. List of differentially expressed genes containing a signal peptide (predicted using Hectar). Supplemental Data Set 5. KEGG pathway enrichment (KOBAS, BLAST e<10e-6) analysis for the set of genes differentially expressed in dis versus apical system, dis versus basal system, and apical versus basal systems. Supplemental File 1. Text file of the alignment corresponding to the phylogenetic analysis in Figure 4D. Acknowledgments We thank Philippe Potin, Cécile Hervé and Elizabeth Ficko-Blean for valuable discussions and the ABiMS Bioinformatics Core Service for providing access to Galaxy. This work was supported by the CNRS, the Agence Nationale de la Recherche (ANR-10-BLAN-1727 and ANR-10-BTBR-04-01), the European Commission Interreg program France (Channel)-England (project Marinexus), the UPMC, and the European Commission (grant agreement 638240). T.U. was supported by a fellowship from the Uehara Memorial Foundation. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. AUTHOR CONTRIBUTIONS O.G., T.U., D.S., S.C., C.N., T.M., L.M., S.M.C., and A.F.P. performed the experiments. A.P.L., K.A., J.M.C., and S.M.C. analyzed data. S.M.C designed the experiments, supervised the work, and wrote the article with input from J.M.C. References 1. André , J. , Harrison , S. , Towers , K. , Qi , X. , Vaughan , S. , McKean , P.G. , Ginger , M.L. ( 2013 ). The tubulin cofactor C family member TBCCD1 orchestrates cytoskeletal filament formation . J. Cell Sci. 126 : 5350 – 5356 . 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TI - DISTAG/TBCCd1 Is Required for Basal Cell Fate Determination in Ectocarpus  
JF - The Plant Cell
DO - 10.1105/tpc.17.00440
DA - 2018-01-05
UR - https://www.deepdyve.com/lp/oxford-university-press/distag-tbccd1-is-required-for-basal-cell-fate-determination-in-s06cewj30N
SP - 3102
EP - 3122
VL - 29
IS - 12
DP - DeepDyve
ER -