Mechanisms for Protein Redistribution in Thylakoids of Anabaena During Cell Differentiation

Mechanisms for Protein Redistribution in Thylakoids of Anabaena During Cell Differentiation Abstract Thylakoid membranes are far from being homogeneous in composition. On the contrary, compositional heterogeneity of lipid and protein content is well known to exist in these membranes. The mechanisms for the confinement of proteins at a particular membrane domain have started to be unveiled, but we are far from a thorough understanding, and many issues remain to be elucidated. During the differentiation of heterocysts in filamentous cyanobacteria of the Anabaena and Nostoc genera, thylakoids undergo a complete reorganization, separating into two membrane domains of different appearance and subcellular localization. Evidence also indicates different functionality and protein composition for these two membrane domains. In this work, we have addressed the mechanisms that govern the specific localization of proteins at a particular membrane domain. Two classes of proteins were distinguished according to their distribution in the thylakoids. Our results indicate that the specific accumulation of proteins of the CURVATURE THYLAKOID 1 (CURT1) family and proteins containing the homologous CAAD domain at subpolar honeycomb thylakoids is mediated by multiple mechanisms including a previously unnoticed phenomenon of thylakoid membrane migration. Introduction The hydrophobic nature of some proteins or protein domains makes them suitable for insertion in biological membranes. However, the multiplicity of membrane systems within a cell requires the existence of mechanisms to allocate proteins to the proper lipid bilayer where they can accomplish their function (Teasdale and Jackson 1996, Derby and Gleeson 2007). Once inserted in the proper membrane, some proteins can be confined to a particular membrane domain, which involves mechanisms for the generation and maintenance of a compositional heterogeneity in the membrane. Compositional heterogeneity is conspicuous in chloroplast thylakoids, where protein complexes are confined in distinct membrane domains, i.e. PSII occupies the grana thylakoids while PSI and the ATP synthase are mostly in the stroma lamellae (Andersson and Anderson 1980, Dekker and Boekema 2005, Nevo et al. 2012). In chloroplast thylakoids, this phenomenon has been well characterized and is commonly referred to as lateral heterogeneity. However, how this uneven arrangement of protein complexes is established and maintained is not fully understood. It is also known that the distribution of protein complexes in the thylakoids is not static and may vary according to the conditions. For instance, light conditions promote state transitions or the PSII repair cycle, both involving long-range migration of membrane macrocomplexes (Joshua and Mullineaux 2004, Bellafiore et al. 2005, Bonardi et al. 2005, Nixon et al. 2005). Cyanobacteria are phylogenetically related to plant chloroplasts (Sagan 1967) and also contain thylakoids, which form an independent membrane system in the cytoplasm that is topologically equivalent to the stroma. However, thylakoids do not form grana or lamellae in cyanobacteria. On the contrary, thylakoid ultrastructure appears in general homogeneous within a species and no domains are distinguishable (Liberton et al. 2006, van de Meene et al. 2006, Nevo et al. 2007, Liberton and Pakrasi 2008, Liberton et al. 2011, Gonzalez-Esquer et al. 2016). An exception to this is observed in heterocysts of filamentous cyanobacteria where thylakoids are partitioned in two domains clearly discernible by their different appearance and subcellular distribution (Lang and Fay 1971, Wilcox et al. 1973, Giddings and Staehelin 1979). A domain known as honeycomb thylakoids is formed by highly contorted membranes densely packed at subpolar regions of the cell, while a second domain, referred to as peripheral thylakoids, is formed by membranes with a less convoluted appearance and a loose distribution in the cytoplasm (Lang and Fay 1971, Wilcox et al. 1973, Sherman et al. 2000). These membrane domains are not related in structure or function to the grana and lamella of chloroplast thylakoids but represent a suitable model to investigate the compositional heterogeneity of thylakoid membranes in cyanobacteria, an issue little investigated in these organisms. Heterocysts are cells specialized in nitrogen fixation that can be observed in multicellular filamentous species of cyanobacteria (Wolk 1996). Nitrogen scarcity triggers a regulatory cascade that provokes the differentiation of some vegetative cells of the filament into heterocysts. Differentiation is aimed at the physical separation into distinct cell types of two incompatible processes: oxygenic photosynthesis and nitrogen fixation. Thus, nitrogen-fixing filaments are composed of heterocysts that perform nitrogen fixation, separated by spacers of 10–15 vegetative cells that fix CO2 by photosynthesis. Heterocysts and vegetative cells exchange carbon and nitrogen compounds, thereby ensuring the correct nutrition of all cells in the filament (Flores and Herrero 2010). Differentiation induces extensive changes in the transcriptome, proteome, ultrastructure and metabolism of the cell (Lang and Fay 1971, Wilcox et al. 1973, Flaherty et al. 2011, Mitschke et al. 2011). Many of these changes are aimed at the expression of the oxygen-sensitive nitrogenase complex and the creation of a microoxic environment that preserves it from the oxygen produced by neighboring cells. In differentiating cells, most of the photosynthetic antenna complex is degraded, oxygen production at PSII is abrogated and photosynthetic CO2 fixation is blocked (Wolk 1996, Herrero et al. 2004). The large requirements of nitrogenase for reducing power and ATP are respectively fulfilled by the catabolism of sugars imported from vegetative cells and by the activity of F0F1-ATP synthase sustained by the electrochemical gradient created by cyclic electron transport around PSI under illumination (Magnuson and Cardona 2016). Another metabolic trait of heterocysts is a high oxygen detoxification activity based on terminal respiratory oxidases and flavodiiron proteins that directly reduce oxygen by the Mehler reaction (Pils et al. 2004, Valladares et al. 2007, Ermakova et al. 2014). Ultrastructural changes that occur during differentiation include the enlargement of the cell, the thickening of the cell wall by deposition of extra polysaccharide and glycolipid layers, the formation of polar granules made of a reserve polymer called cyanophycin and a complete reorganization of thylakoid membranes (Lang and Fay 1971, Wilcox et al. 1973, Giddings and Staehelin 1979). In the genera Anabaena and Nostoc, thylakoids are arranged in vegetative cells forming parallel layers at the periphery of the cytoplasm, the central part of the cytosol being mostly free from membranes and occupied by the nucleoid. Differentiation provokes an extensive rearrangement, which may involve synthesis of new membranes, so that in mature heterocysts thylakoids are partitioned into honeycomb and peripheral thylakoids. These two domains are not only different in localization and appearance, but evidence also indicates the existence of compositional heterogeneity, with proteins that specifically reside in only one of these two domains. However, the protein constituents of each domain remain to be fully characterized (Murry et al. 1981, Valladares et al. 2007, Cardona et al. 2009). Despite this information, many aspects about the formation of these two membrane domains and the distribution of proteins in each of them remain obscure. Furthermore, it is still not clear where to draw the line between honeycomb and peripheral thylakoids and whether they are physically interconnected. Photosynthetic electron transport is the major role of thylakoids, and most proteins anchored to or associated with these membranes are related to this function. However, the thylakoids of some cyanobacterial species also harbor proteins involved in gene translation, namely aminoacyl-tRNA synthetases (aaRSs). Membrane anchoring of these enzymes occurs through an extra domain called CAAD that contains two transmembrane segments. CAAD has an inherent capacity for directing proteins to the thylakoid membrane, not requiring any signal peptide (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016). Interestingly, CAAD is homologous to proteins of the CURVATURE THYLAKOID 1 (CURT1) family (Luque and Ochoa de Alda 2014). Proteins of this family have membrane-bending capacity and were shown to confer their characteristic shape to the thylakoids of Arabidopsis chloroplasts and the cyanobacterium Synechocystis (Armbruster et al. 2013, Heinz et al. 2016). In Arabidopsis, CURT1 proteins contribute to the lateral heterogeneity of thylakoids, showing a specific distribution at grana margins, where they induce acute curvature to the membrane (Armbruster et al. 2013). How proteins select their proper localization in the thylakoids of heterocysts is still an open question. We have analyzed the subcellular localization of thylakoidal proteins of Anabaena sp. PCC 7120 (also known as Nostoc sp. PCC 7120, hereafter Anabaena). Two distinct behaviors were observed that allowed their partitioning into two classes. One class is represented by the F0F1-ATP synthase and HetN, which in mature heterocysts occupied both honeycomb and peripheral thylakoids. A second class includes proteins specifically located at honeycomb thylakoids and includes valyl-tRNA synthetase (ValRS), CurT proteins and FraH. We have investigated how proteins of this second class select their specific localization at this particular domain. Results presented here indicate that multiple mechanisms contribute to the accumulation and specific confinement of some class 2 proteins at honeycomb thylakoids during heterocyst differentiation. Results Specific localization of proteins at discrete domains of the thylakoids in heterocysts Fig. 1A illustrates the ultrastructure of vegetative cells and heterocysts, evidencing the different appearance of the thylakoids in each cell type and their segregation in two domains in heterocysts (see also Lang and Fay 1971, Wilcox et al. 1973, Sherman et al. 2000). Honeycomb thylakoids were observed at a subpolar position in the vicinity of the cyanophycin granule, whereas peripheral thylakoids colonized other areas of the cytoplasm (Fig. 1A). Since these two domains are only discernible at late stages of differentiation or in mature heterocysts, the terms honeycomb (h) and peripheral (p) thylakoids will be used throughout this work to refer to membrane domains of mature heterocysts (or late pro-heterocysts). For premature differentiation stages, all membranes in the cytoplasm will be referred to as internal membranes. To investigate the distribution of proteins in h and p thylakoids of Anabaena heterocysts, the fluorescence of green fluorescent protein (GFP) fusions (see the Materials and Methods) was monitored by confocal microscopy. Mature heterocysts can be easily distinguished from surrounding vegetative cells by their enlarged body, their low autofluorescence and conspicuous refringent cyanophycin granules at both poles (Fig. 1B). Two distinct patterns were observed. The F0F1-ATP synthase complex and HetN showed fluorescent signals at subpolar regions, as well as discrete irregular signals elsewhere in the cytoplasm that varied in shape from cell to cell (Fig. 2A, B). The distribution of GFP-tagged ATP synthase (Santamaría-Gómez et al. 2016) in the tridimensional space of the heterocyst cytoplasm was analyzed in detail by z-axis montages (Fig. 2C) and 3-D image reconstruction (Supplementary Movie S1). Z-axis montages revealed that the subpolar signals showed a lenticular form whereas the non-polar irregular signals traversed the cytoplasm in distinct directions apparently forming bridges that connected the subpolar signals (Fig. 2C; Supplementary Movie S1). Tomography images corroborated the subcellular localization of the ATP synthase observed by z-montages (Supplementary Movie S2). These results indicated the localization of ATP-synthase and HetN in h and p thylakoids, which is consistent with previous reports showing decoration of both membrane domains with antibodies against the α- and β-subunits of ATP synthase (Sherman et al. 2000). Fig. 1 View largeDownload slide Cell structure of Anabaena cells. (A) Ultrastructure of vegetative cells and heterocysts in Anabaena. cw, cell wall; c, cyanophycin granule; t, vegetative cell thylakoids; n, neck. Size bar corresponds to 1 µm. (B) Bright field image (top) and confocal fluorescence image of the photosynthetic pigments of Anabaena filament (bottom). H, heterocysts; V, vegetative cell; c, cyanophycin granules. Confocal fluorescence images were obtained using a Leica TCS SP2 confocal microscope (see the Materials and Methods for details). Fig. 1 View largeDownload slide Cell structure of Anabaena cells. (A) Ultrastructure of vegetative cells and heterocysts in Anabaena. cw, cell wall; c, cyanophycin granule; t, vegetative cell thylakoids; n, neck. Size bar corresponds to 1 µm. (B) Bright field image (top) and confocal fluorescence image of the photosynthetic pigments of Anabaena filament (bottom). H, heterocysts; V, vegetative cell; c, cyanophycin granules. Confocal fluorescence images were obtained using a Leica TCS SP2 confocal microscope (see the Materials and Methods for details). Fig. 2 View largeDownload slide Subcellular localization of GFP fusion proteins in heterocysts. (A) Panels show the fluorescence of Anabaena derivative strains. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Other panels show the fluorescence of filaments expressing GFP fusion proteins. The name of the protein fused to GFP is indicated on the left of each panel. For each fusion, several panels are shown. The first panel (from left to right) corresponds to the fluorescence of GFP (λemision = 500–540 nm), the second panel corresponds to the autofluorescence of photosynthetic pigments (λemision = 630–700 nm), the third panel is the superposition of the two previous panels and the fourth panel is the superposition of the GFP fluorescence image and the bright field image. Plots on the right show the quantification of GFP fluorescence. Heterocysts are indicated with an ‘H’. In the plot, the image of the fourth panel is projected on the horizontal plane and the fluorescence of each area is represented on the vertical axis. Units of the x- and z-axis are microns and those of the y-axis are arbitrary units of fluorescence intensity. Size bars corresponds to 1 µm. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions (λemision = 500–540 nm) at these regions was quantified in heterocysts (n = 30) of filaments expressing each fusion protein. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units on the y-axis. Horizontal lines indicate the average value. (C) Z-axis montage of confocal microscopy images of cells expressing AtpA–GFP. Panels corresponding to serial layers of the preparation are presented in an ordered array. H, heterocyst; V, vegetative cell. Panels on the left show the fluorescence of the wild-type strain used as a negative control as in (A). Fig. 2 View largeDownload slide Subcellular localization of GFP fusion proteins in heterocysts. (A) Panels show the fluorescence of Anabaena derivative strains. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Other panels show the fluorescence of filaments expressing GFP fusion proteins. The name of the protein fused to GFP is indicated on the left of each panel. For each fusion, several panels are shown. The first panel (from left to right) corresponds to the fluorescence of GFP (λemision = 500–540 nm), the second panel corresponds to the autofluorescence of photosynthetic pigments (λemision = 630–700 nm), the third panel is the superposition of the two previous panels and the fourth panel is the superposition of the GFP fluorescence image and the bright field image. Plots on the right show the quantification of GFP fluorescence. Heterocysts are indicated with an ‘H’. In the plot, the image of the fourth panel is projected on the horizontal plane and the fluorescence of each area is represented on the vertical axis. Units of the x- and z-axis are microns and those of the y-axis are arbitrary units of fluorescence intensity. Size bars corresponds to 1 µm. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions (λemision = 500–540 nm) at these regions was quantified in heterocysts (n = 30) of filaments expressing each fusion protein. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units on the y-axis. Horizontal lines indicate the average value. (C) Z-axis montage of confocal microscopy images of cells expressing AtpA–GFP. Panels corresponding to serial layers of the preparation are presented in an ordered array. H, heterocyst; V, vegetative cell. Panels on the left show the fluorescence of the wild-type strain used as a negative control as in (A). In contrast, GFP fusions of other membrane proteins including ValRS, the two CurT homologs, Alr0805 and Alr4119, and FraH were detected as conspicuous fluorescent subpolar foci in mature heterocysts, with signals not being observed elsewhere in the cell, which indicated their specific localization at h thylakoids. A GFP fusion of CAAD, the domain that anchors ValRS to the membrane, showed the same localization pattern as full-length ValRS in heterocysts (Fig. 2A, B), which is consistent with previous observations (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016) and indicates that the determinants for the specific localization of ValRS in h thylakoids reside in the CAAD domain. These results allowed the partition of thylakoidal proteins into two classes: class 1, represented by the F0F1-ATP synthase and HetN; and class 2, which includes ValRS, Alr0805, Alr4119 and FraH. Fluorescence quantification clearly revealed two patterns corresponding to classes 1 and 2 (Fig. 2A, B). Furthermore, these observations lend support to the existence of a compositional heterogeneity between h and p thylakoids (Cardona et al. 2009, Magnuson and Cardona 2016), which entails the existence of mechanisms that promote the specific localization of proteins at particular membrane domains. An interesting observation was that the CurT proteins, Alr0805 and Alr4119, showed opposite patterns of expression: Alr0805 was abundant in vegetative cells and scarce in heterocysts, while Alr4119 was abundant in heterocysts and not detected in vegetative cells (Fig. 3). Here we propose to name these proteins as CurT1 (Alr0805) and CurT2 (Alr4119). The confinement of these proteins in h thylakoids suggests that they may be involved in conferring acute curvature to these membranes. Fig. 3 View largeDownload slide Differential expression profile of CurT proteins in Anabaena. Panels corresponding to composite images of cells expressing Alr0805–GFP or Alr4119–GFP from their native promoter (as indicated) are shown. Composite images of GFP fluorescence and the bright field image are shown on the left, and composite images of the red autofluorescence of photosynthetic pigments and GFP fluorescence are shown on the right. Some heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The three bottom panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Fig. 3 View largeDownload slide Differential expression profile of CurT proteins in Anabaena. Panels corresponding to composite images of cells expressing Alr0805–GFP or Alr4119–GFP from their native promoter (as indicated) are shown. Composite images of GFP fluorescence and the bright field image are shown on the left, and composite images of the red autofluorescence of photosynthetic pigments and GFP fluorescence are shown on the right. Some heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The three bottom panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Redistribution of GFP–CAAD during cell differentiation is a sequential process with two distinct phases Expression of the GFP–CAAD fusion described above had a low impact on the physiology of Anabaena and was chosen as a model to analyze how class 2 proteins localize specifically at honeycomb membranes. This fusion protein was considered a good representative of class 2 proteins since it reproduced the localization of ValRS (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016) and was homologous to CurT1 and CurT2 (Supplementary Fig. S1), which suggested that they would share the same mechanisms for their specific localization at h thylakoids. GFP–CAAD was observed uniformly distributed in the thylakoids of vegetative cells, whereas in heterocysts it was confined at h thylakoids (Fig. 2A) (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016). Therefore, this change in the subcellular distribution of GFP–CAAD must be completed by the 24 h that a vegetative cell takes to differentiate into a heterocyst. To gather information on the subcellular localization at intermediate stages of differentiation, GFP–CAAD was expressed in Anabaena from the heterocyst-specific patS promoter, which is induced early in differentiating cells (Yoon and Golden 2001), and GFP fluorescence was monitored at different time points after the initiation of heterocyst differentiation elicited by nitrogen step-down (see the Materials and Methods). At the initiation of the experiment, filaments showed no GFP fluorescence. After 6 h, cells regularly spaced in the filament, separated by approximately 10–20 cells, showed dim GFP fluorescence. These fluorescent cells were cells initiating differentiation, where the signal was observed at the cell periphery, co-localizing with the thylakoid membranes, detectable by the red fluorescent signal of photosynthetic pigments (Fig. 4A, B; Supplementary Fig. S2). At 12 h after the initiation, GFP–CAAD fluorescence remained at the cell periphery, but some accumulation was detected close to the poles of the cell, and 6 h later (18 h) the fluorescent signal was more intense at the poles and dimmer at the periphery (Fig. 4A–C; Supplementary Fig. S2). In contrast, no enrichment of the red signal at the poles was observed at 12 or 18 h (Fig. 4C; Supplementary Fig. S2). At 24 h, filaments contained mature heterocysts, where GFP fluorescence no longer remained at the cell periphery, being confined exclusively at a subpolar position (Fig. 4A, B). Fig. 4 View largeDownload slide Phases in the redistribution of GFP–CAAD during heterocyst differentiation. (A) Cultures of Anabaena expressing GFP–CAAD were subjected to nitrogen withdrawal and images were taken by confocal microscopy at different time points, as indicated. Details of the different panels are as in Fig. 2. Pro-heterocysts and heterocysts are situated at the center of each panel. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions in these regions was quantified in heterocysts (n = 20) of filaments expressing GFP–CAAD. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units in the y-axis. Horizontal lines indicate the average value. (C) Fluorescence intensity profiles of the periphery of differentiating cells. Panels correspond to the images shown on the right in (A). Fluorescence intensity of the green and red channels was quantitated in the zone covered by the yellow ring and plotted. The ring line starts at the white bar, which corresponds to the 0 point on the x-axis, and continues clockwise. (D) Fluorescence microscopy images of Anabaena cells expressing GFP–CAAD taken at 16 and 24 h after nitrogen compound withdrawal. Pro-heterocysts and heterocysts are situated at the center of each panel. Images were subjected to a deconvolution treatment to improve resolution (see the Materials and Methods). White arrowheads indicate the septal area between the heterocyst and adjacent vegetative cells. Plots correspond to the quantification of the fluorescence of GFP (green) and of photosynthetic pigments (red). Units of the x- and z-axis are microns. The y-axis indicates the relative fluorescence intensity in arbitrary units. Fig. 4 View largeDownload slide Phases in the redistribution of GFP–CAAD during heterocyst differentiation. (A) Cultures of Anabaena expressing GFP–CAAD were subjected to nitrogen withdrawal and images were taken by confocal microscopy at different time points, as indicated. Details of the different panels are as in Fig. 2. Pro-heterocysts and heterocysts are situated at the center of each panel. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions in these regions was quantified in heterocysts (n = 20) of filaments expressing GFP–CAAD. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units in the y-axis. Horizontal lines indicate the average value. (C) Fluorescence intensity profiles of the periphery of differentiating cells. Panels correspond to the images shown on the right in (A). Fluorescence intensity of the green and red channels was quantitated in the zone covered by the yellow ring and plotted. The ring line starts at the white bar, which corresponds to the 0 point on the x-axis, and continues clockwise. (D) Fluorescence microscopy images of Anabaena cells expressing GFP–CAAD taken at 16 and 24 h after nitrogen compound withdrawal. Pro-heterocysts and heterocysts are situated at the center of each panel. Images were subjected to a deconvolution treatment to improve resolution (see the Materials and Methods). White arrowheads indicate the septal area between the heterocyst and adjacent vegetative cells. Plots correspond to the quantification of the fluorescence of GFP (green) and of photosynthetic pigments (red). Units of the x- and z-axis are microns. The y-axis indicates the relative fluorescence intensity in arbitrary units. It is worth noting that signals observed at polar positions at 12 and 18 h were different in appearance and location from those observed at 24 h (Fig. 4A). To obtain a deeper insight on this issue, pro-heterocyst (16 h) and mature heterocysts (24 h) were analyzed by fluorescence microscopy followed by a deconvolution treatment of images to improve resolution (Fig. 4D; Supplementary Fig. S3; Supplementary Movies S3, S4). In pro-heterocysts (16 h), signals of GFP–CAAD at the cell poles were concentrated at discrete points very close to the septum with adjacent vegetative cells. Notice that the green fluorescence at the poles appears to overflow the red fluorescence signal (Fig. 4D, left panel; Supplementary Fig. S3; Supplementary Movie S3). In mature heterocysts (24 h), GFP–CAAD fluorescence was subpolar (closer to the center of the cell); the signal occupied a larger area and showed a crescent-like form (Fig. 4D, right panel; Supplementary Fig. S3; Supplementary Movie S4). These observations indicated that the redistribution of GFP–CAAD during differentiation is a sequential process that includes a long-lasting phase (0–18 h) where the GFP–CAAD concentration gradually decreases at the cell periphery and increases at the poles, very close to the septum; and a second phase (18–24 h) where the protein is shifted 0.3–0.5 µm toward the center of the cell (Fig. A–C). It is important to point out that in the first phase the dynamics of GFP–CAAD appear to be independent of the putative dynamics of the internal membranes (compare green and red signals in Fig. 4C and Supplementary Fig. S2). Mechanism for GFP–CAAD redistribution during cell differentiation Two non-mutually exclusive possibilities were considered for the GFP–CAAD redistribution in differentiating cells: (i) the migration of the protein within the cell or (ii) the replacement of the pre-existing protein by de novo synthesized molecules that are inserted specifically at h membranes. To test the first option, an assay was set up to monitor the dynamics of GFP–CAAD during differentiation in conditions where GFP–CAAD molecules are not synthesized de novo. For this, the PrbcL promoter, specific for vegetative cells and inactive in heterocysts (Elhai and Wolk 1990, Madan and Nierzwicki-Bauer 1993, Ramasubramanian et al. 1994), was chosen to direct the expression of reporter proteins. Since PrbcL is strongly repressed in differentiating cells (Elhai and Wolk 1990, Madan and Nierzwicki-Bauer 1993, Ramasubramanian et al. 1994), fluorescence signals in these cells would correspond to the GFP–CAAD molecules synthesized before the onset of differentiation. Monitoring the fluorescent signal in these cells would reveal whether GFP–CAAD molecules remain immobile or migrate within the cell during differentiation. To test the experimental design, a PrbcL–GFP construction was introduced into Anabaena. In nitrogen-rich medium, all cells of the filament showed intense fluorescence, consistent with the strong activity of the PrbcL promoter (Fig. 5A, central panel). When filaments were transferred to medium with no nitrogen compound, cells at semi-regular intervals, identified as differentiating cells by their larger size, showed a gradual decay of GFP fluorescence (Fig. 5A, right panel). The fluorescence decay was attributed to the absence of PrbcL activity and the degradation of GFP protein with time. In line with this, prolonged incubation times (>36 h) yielded a large proportion of heterocysts with no fluorescence signal. However, due to the great stability of GFP (in vitro half-life >26 h) (Corish and Tyler-Smith 1999), faint but detectable signals remained in pro-heterocysts (18–23 h) and mature heterocysts (24 h), validating this approach. In filaments expressing GFP–CAAD from PrbcL, very strong fluorescent signals of punctate appearance, mostly overlapping with the red fluorescence of thylakoids, were distinguished in vegetative cells (Fig. 5B; Supplementary Fig. S4). In contrast, in mature heterocysts (24 h), faint fluorescence signals were observed. Importantly, in virtually all mature heterocysts showing remnant fluorescence, these signals were at subpolar positions (93%, n = 57) (Fig. 5B; Supplementary Fig. S4). These results indicated that molecules synthesized previously (i.e. in the progenitor vegetative cell) had migrated during differentiation from the periphery to a subpolar position of the cell. Fig. 5 View largeDownload slide Diffusion of GFP–CAAD in internal membranes of heterocysts. (A) Cells expressing GFP from the PrbcL promoter were cultured in the presence of nitrate (central panel) or in the absence of nitrogen compounds (right panel). Heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The left panel corresponds to a composite image of wild-type filaments in the bright field and green fluorescence channels shown as a control of the absence of fluorescence in the 500–540 nm band. (B) Composite fluorescent images of Anabaena cells expressing GFP–CAAD from the PrbcL promoter. A heterocyst occupies the center of each panel. The size bar corresponds to 2 µm. The three panels on the left show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (C) FRAP analysis of a pro-heterocyst expressing GFP–CAAD. FRAP experiments were done using a Leica TCS SP5 confocal microscope (see the Materials and Methods for details). The first panel labeled as ‘Pre’ shows an image of the heterocyst before bleaching. Other panels correspond to images taken at the time indicated in seconds after bleaching. A white arrow indicates the bleached area. The plot on the right shows the evolution of the fluorescence signal during the experiment. Bleaching was performed at time = 0. The average and SD are indicated; n is the number of repeats of this experiment. (D) FRAP analysis of mature heterocyst expressing GFP–CAAD. Details are as in (C). (E) FRAP analysis of mature heterocyst expressing AtpA–GFP. Details are as in (C). (F) FRAP analysis of a mature heterocyst expressing GFP–CAAD. The entire region occupied by the heterocyst (indicated by a bracket in the left panel) was bleached and fluorescence recovery was followed for 300 s. Details are as in (C). Fig. 5 View largeDownload slide Diffusion of GFP–CAAD in internal membranes of heterocysts. (A) Cells expressing GFP from the PrbcL promoter were cultured in the presence of nitrate (central panel) or in the absence of nitrogen compounds (right panel). Heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The left panel corresponds to a composite image of wild-type filaments in the bright field and green fluorescence channels shown as a control of the absence of fluorescence in the 500–540 nm band. (B) Composite fluorescent images of Anabaena cells expressing GFP–CAAD from the PrbcL promoter. A heterocyst occupies the center of each panel. The size bar corresponds to 2 µm. The three panels on the left show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (C) FRAP analysis of a pro-heterocyst expressing GFP–CAAD. FRAP experiments were done using a Leica TCS SP5 confocal microscope (see the Materials and Methods for details). The first panel labeled as ‘Pre’ shows an image of the heterocyst before bleaching. Other panels correspond to images taken at the time indicated in seconds after bleaching. A white arrow indicates the bleached area. The plot on the right shows the evolution of the fluorescence signal during the experiment. Bleaching was performed at time = 0. The average and SD are indicated; n is the number of repeats of this experiment. (D) FRAP analysis of mature heterocyst expressing GFP–CAAD. Details are as in (C). (E) FRAP analysis of mature heterocyst expressing AtpA–GFP. Details are as in (C). (F) FRAP analysis of a mature heterocyst expressing GFP–CAAD. The entire region occupied by the heterocyst (indicated by a bracket in the left panel) was bleached and fluorescence recovery was followed for 300 s. Details are as in (C). To test whether such migration of GFP–CAAD occurs by diffusion of this protein in the lipid bilayer, FRAP (fluorescence recovery after photobleaching) experiments were performed. FRAP has been extensively used to analyze the lateral movement of fluorescent proteins in the plane of thylakoid membranes (Mullineaux et al. 1997, Mullineaux and Sarcina 2002, Mullineaux 2004, Kaňa 2013). FRAP was first performed in pro-heterocysts expressing GFP–CAAD (16–20 h) by bleaching regions roughly at the equator of the cell (Fig. 5C, top panels) or at one of the poles (Fig. 5C, bottom panels). Recovery of fluorescence in the bleached area was observed in the order of seconds to minutes. Such recovery cannot be attributed to new protein synthesis since no recovery was observed in entirely bleached cells after 5 min (Fig. 5F). Therefore, FRAP results indicated that in pro-heterocysts GFP–CAAD can diffuse in thylakoids at different sites in the cell. Diffusion was tested in mature heterocysts (24 h), where GFP–CAAD is confined at h thylakoids, by bleaching the signal at one of the poles. No recovery was observed after 300 s (Fig. 5D), and similar results were obtained when half of the area was bleached, indicating that the protein is not mobile at h thylakoids. This lack of mobility is a specific feature of GFP–CAAD that may be shared by other class 2 proteins, but does not appear to apply to class 1 proteins, as recovery of AtpA–GFP fluorescence was observed when one pole of mature heterocysts was bleached (Fig. 5E). This latter observation has important implications as it lends empirical support to the existence of physical continuity between h and p thylakoids (Fig. 5E), an issue that so far has remained controversial. The shift in the position of GFP–CAAD from a polar to a subpolar position observed in the second phase (see above, Fig. 4) was intriguing. To obtain an insight into this issue, cells at late stages of differentiation (16–24 h) were analyzed by electron microscopy with a particular focus on polar and subpolar regions (Fig. 6A–D). In parallel, the subcellular localization of GFP–CAAD was monitored by fluorescence microscopy (Fig. 6E–L). In pro-heterocysts (16 h), the neck of the cell, a narrow enlargement of the cytoplasm at the poles that reduces the section of the septum in contact with the neighboring vegetative cell (Fig. 1A), was observed to be full of internal membranes (Fig. 6A). This was concomitant with the observation of GFP–CAAD as a strong fluorescent signal very close to the neighboring cell (Fig. 6E, I), suggesting that this protein could be colonizing the internal membranes located within the neck. At later time points, cyanophycin was observed to accumulate starting close to the septum and gradually filling the neck till it overflowed it, which results in the characteristic cup-like form of cyanophycin in mature heterocysts (Fig. 6B–D). The growth of the cyanophycin granule paralleled the changes observed for the GFP–CAAD signal at these stages of differentiation, including a displacement of about 0.3–0.5 µm toward the center of the cell and a reshaping of the signal from an intense concentrated dot to a more expanded crescent-like form, reminiscent of the shape of h thylakoids in mature heterocysts. Taken together, these observations suggested that at these stages of differentiation (16–24 h) the accumulation of cyanophycin dislodges internal membranes out of the neck of the heterocyst. It follows that in contrast to the first phase, the displacement of GFP–CAAD in this second phase would be sustained by a distinct phenomenon: the migration of membranes that carry this protein as an integral component. Fig. 6 View largeDownload slide Ultrastructure of cells in the last stages of differentiation, and subcellular localization of GFP–CAAD. Panels show cells in different stages of differentiation (A, E, I, 16 h; B, F, J, 20 h; C, G, K, 22 h; D, H, L, 24 h). (A–D) Electron microscopy of pro-heterocysts and heterocysts in different stages of differentiation as indicated above. The bracket indicates the neck of the cell, ‘c’ denotes cyanophycin. The size bar corresponds to 1 µm. Images (A), (B), (C) and (D) are representative of four, seven, six and nine different heterocysts, respectively. (E–H) Composite images of the bright field and GFP fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. (I–L) Composite images of the green and red fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. Images (E–L) are representative of at least 10 different heterocysts. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Fig. 6 View largeDownload slide Ultrastructure of cells in the last stages of differentiation, and subcellular localization of GFP–CAAD. Panels show cells in different stages of differentiation (A, E, I, 16 h; B, F, J, 20 h; C, G, K, 22 h; D, H, L, 24 h). (A–D) Electron microscopy of pro-heterocysts and heterocysts in different stages of differentiation as indicated above. The bracket indicates the neck of the cell, ‘c’ denotes cyanophycin. The size bar corresponds to 1 µm. Images (A), (B), (C) and (D) are representative of four, seven, six and nine different heterocysts, respectively. (E–H) Composite images of the bright field and GFP fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. (I–L) Composite images of the green and red fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. Images (E–L) are representative of at least 10 different heterocysts. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. The results above illustrate that existing GFP–CAAD molecules can be displaced within the cell during the differentiation process, contributing to its accumulation in thylakoids at polar or subpolar positions. The fate of class 2 proteins synthesized de novo remains to be investigated. This issue was analyzed at late stages of differentiation once the two membrane domains, h and p thylakoids, are fully developed. In order to monitor only de novo synthetized molecules and avoid the interference by pre-existing ones, GFP–CAAD was expressed from a heterocyst-specific late promoter, PhetN. This promoter was reported to be induced about 17 h after nitrogen compound withdrawal (Bauer et al. 1997, Callahan and Buikema 2001, Flaherty et al. 2011). In our experiments, GFP–CAAD fluorescent signals started to be faintly visible 20 h after the initiation of differentiation (Fig. 7). As a control, cells expressing the HetN–GFP fusion protein from the PhetN promoter were also analyzed. Interestingly, mature heterocysts expressing HetN–GFP showed fluorescence at subpolar and peripheral positions, indicating its presence at h and p thylakoids (100%, n = 46). In contrast, virtually all cells expressing GFP–CAAD showed fluorescence exclusively at the subpoles (96%, n = 75). These observations indicated that the GFP–CAAD molecules synthetized de novo at the late stages of differentiation insert specifically at h thylakoids. Fig. 7 View largeDownload slide Subcellular localization of GFP–CAAD and HetN–GFP fusion proteins expressed from the late heterocyst-specific hetN promoter. Images correspond to cells bearing the construction indicated on the left. Bright field images (left) and composite fluorescence images (right) are shown. Heterocysts are indicated by white arrowheads. Size bars correspond to 20 µm. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Fig. 7 View largeDownload slide Subcellular localization of GFP–CAAD and HetN–GFP fusion proteins expressed from the late heterocyst-specific hetN promoter. Images correspond to cells bearing the construction indicated on the left. Bright field images (left) and composite fluorescence images (right) are shown. Heterocysts are indicated by white arrowheads. Size bars correspond to 20 µm. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Before differentiation (i.e. in the progenitor vegetative cell), GFP–CAAD shows a peripheral distribution coincident with the position of thylakoids, whereas in mature heterocysts it is confined at h thylakoids at subpolar positions. Data presented above allow us to conclude that three different phenomena contribute to this redistribution: (i) protein diffusion in the thylakoid membrane; (ii); thylakoid migration; and (iii) specific insertion of de novo synthesized proteins at h thylakoids. Discussion Cyanobacterial thylakoids do not form grana and show a uniform appearance with no discernible domains (Gonzalez-Esquer et al. 2016). However, in heterocysts of filamentous cyanobacteria of the Anabaena and Nostoc genera, the separation of thylakoids into two domains was described decades ago (Lang and Fay 1971, Wilcox et al. 1973, Wolk 1996, Sherman et al. 2000). Despite this, many issues about the generation of h and p thylakoids remain unknown. For instance, it is still not clear whether the separation of these two domains occurs by remodeling of pre-existing thylakoids, if it requires the synthesis of new membranes, or both. Previous evidence suggested distinct functionality and protein composition for h and p domains (Murry et al. 1981, Valladares et al. 2007, Ferimazova et al. 2013). Cardona et al. could separate two fractions in their preparations of heterocyst internal membranes and showed that some proteins were specifically found in only one of them (Cardona et al. 2009). However, it is not clear whether these two fractions corresponded to h and p thylakoids. Therefore, the precise localization of proteins at either or both membrane domains remained mostly an open question. Results in this work add further evidence to the existence of a compositional heterogeneity between h and p thylakoids and allowed the partition of proteins into two classes according to their distribution in these membranes (Fig. 2). Some observations (Fig. 5C, E) indicate that there is continuity between h and p thylakoids in mature heterocysts. However, the uneven distribution of proteins in these two domains entails the existence of mechanisms for the generation and maintenance of compositional heterogenetity. We have investigated how class 2 proteins are specifically confined at h thylakoids. An important issue is how this specificity is established (i.e. how class 2 proteins can distinguish h from p thylakoids). A widespread mechanism for the specific placement of proteins at defined membrane areas is diffusion and capture, consisting of the free lateral diffusion of proteins in the bilayer until they interact with a trapping factor that can be a molecule (e.g. a lipid or protein partner already present at the destination) (Shapiro et al. 2009, Laloux and Jacobs-Wagner 2014) or a geometrical cue (e.g. membrane curvature) (Huang and Ramamurthi 2010). The redistribution of GFP–CAAD during differentiation could fit with this model as ab initio this protein is relatively mobile in the membrane but later on it becomes motionless in the h thylakoids (Fig. 5C). Though this lack of mobility may partially derive from macromolecular crowding or high viscosity of this membrane domain (Kirchhoff et al. 2004, Mullineaux 2008, Kaňa 2013, Kirchhoff 2014, Liu 2016), the observation that other proteins (i.e. AtpA–GFP) can diffuse in these membranes supports the existence of (a) specific trapping molecule(s) for GFP–CAAD. Alternatively, given the highly contorted nature of honeycomb membranes, class 2 proteins may localize there by direct sensing of membrane curvature (Huang and Ramamurthi 2010, Laloux and Jacobs-Wagner 2014, Strahl et al. 2015). Interestingly, CurT1 and CurT2 proteins contain predicted amphipathic helices in their N-termini (Supplementary Fig. S1B) and some reports have described that amphipathic helices have the ability to sense membrane curvature whereas other reports show that these helices induce membrane remodeling (see Drin and Antonny 2010, and references therein). In line with this, proteins of the CURT1 family from Synechocystis and Arabidopsis were found enriched at thylakoid regions of acute curvature and were shown to have the intrinsic ability to introduce curvature to liposomes in in vitro assays (Armbruster et al. 2013, Heinz et al. 2016). An interesting observation was the contrasting expression profile of CurT proteins in Anabaena (Fig. 3). CurT1 is expressed in all cells, although it is less abundant in heterocysts, which points to this protein as the housekeeping CurT in Anabaena. Both CurT1 and CurT2 are expressed in heterocysts, and are specifically found at h thylakoids in mature heterocysts (Fig. 2). This suggests that either the specific presence of CurT2, or the joint action of both proteins (i.e. by hetero-oligomerization) could be responsible for conferring the characteristic intricate shape to h thylakoids. Given the homology of the ValRS CAAD domain with proteins of the CURT1 family, it is tempting to speculate that this domain could also contribute to shaping h thylakoids. However, is worth noting that the CAAD sequence from Anabaena ValRS was not predicted to contain amphipathic helices (Supplementary Fig. S1B) (Luque and Ochoa de Alda 2014). Diffusion and trapping by a lipid or protein partner or by membrane curvature contributes to but it is not entirely responsible for the redistribution of GFP–CAAD during the differentiation of heterocysts. Data herein show that such a redistribution results from the additive effect of multiple phenomena (i.e. protein diffusion, membrane migration and specific insertion at polar membranes). However, these three distinct mechanisms appear to operate at successive stages of differentiation. At early stages (phase 1) the protein would freely diffuse in the membrane until at about 12 h an attracting or trapping element emerges at polar membranes (Figs. 4, 5C, D) and provokes accumulation of GFP–CAAD at membranes that fill the neck of the cell (Figs. 4D, 6A; Supplementary Fig. S3). In this phase, the redistribution of GFP–CAAD is apparently independent of putative membrane dynamics (Fig. 4C; Supplementary Fig. S2). At later stages (phase 2; 18–24 h), membranes containing GFP–CAAD would migrate out of the neck, expelled by the growth of the cyanophycin granule (Fig. 6). Hence, GFP–CAAD is passively transported to its final destination as an integral component of these membranes. Though global reorganization of thylakoids during differentiation was long suspected to involve the migration of membranes, to our knowledge this was not demonstrated. The use of GFP–CAAD as a marker of membrane localization has provided empirical evidence for the occurrence of this phenomenon in the neck of late pro-heterocysts. Our observations find further support in electron micrographs from previous studies that focused on other issues of differentiation (Lang and Fay 1971, Wilcox et al. 1973, Sherman et al. 2000, Zheng et al. 2017). Finally, in the last stages of phase 2, when the h domain is developed (in late pro-heterocysts and mature heterocysts), the de novo synthesized GFP–CAAD protein would insert specifically in such membrane domain, further contributing to the accumulation of GFP–CAAD protein in these membranes. It is not known whether this happens by in situ translation in ribosomes associated with h thylakoids (Matsumoto et al. 2015), or if the protein is synthesized by cytosolic ribosomes and then directed to these membranes by unknown factor(s). In any case, data presented in this work revealed the existence of sorting mechanisms to direct mRNA or proteins to specific domains of the internal membranes in mature heterocysts. The three mechanisms shown here to direct the relocalization of GFP–CAAD most probably operate for class 2 proteins such as ValRS, CurT1 and perhaps CurT2. However, this may not be the case for other class 2 proteins such as FraH (Merino-Puerto et al. 2011). In heterocysts, FraH is only expressed at late stages of differentiation. Thus, it is possible that subpolar accumulation of FraH occurs only by specific insertion of proteins synthesized de novo. Interestingly, fraH mutants lack h thylakoids, so FraH was suggested to have a role in the development or in the maintenance of this membrane domain (Merino-Puerto et al. 2011), which deserves further investigation. Cyanobacteria and plastids are highly ordered entities and their structure determines how they function (Jarvis and Lopez-Juez 2013, Gonzalez-Esquer et al. 2016). Proper arrangement of thylakoid proteins at specific membrane domains is important for normal functioning as well as for acclimation to particular conditions (Pribil et al. 2014). In addition, redistribution of some proteins in the thylakoids was shown to be important for acclimation and developmental phenomena, including the PSII repair cycle during photoinhibition, the placement of the PSII antenna during state transitions or the differentiation of plastids (Joshua and Mullineaux 2004, Bellafiore et al. 2005, Bonardi et al. 2005, Nixon et al. 2005,Jarvis and Lopez-Juez 2013). The differentiation of heterocysts, which can be regarded both as an acclimation response to nitrogen scarcity and a developmental phenomenon within the filament, is accompanied by a redistribution of thylakoid proteins. The consequent compositional heterogeneity reinforces the idea of a division of labor between h and p thylakoids and underlines the importance of the peculiar ultrastructure of heterocysts for their functioning. Results shown herein identify mechanisms responsible for the compositional heterogeneity of heterocyst thylakoids. We propose that similar mechanisms may operate jointly or individually for the establishment of compositional heterogeneity in thylakoids of other cyanobacteria or plant chloroplasts. Materials and Methods Organisms and growth conditions Anabaena sp. PCC 7120 and derivative strains were routinely grown in BG11 medium (Rippka 1988) at 30°C, under continuous illumination (75 photon µmol m–2 s–1), either continuously shaken or bubbled with a mixture of CO2 and air (1% v/v), and in this case supplemented with 10 mM NaHCO3; 1% (w/v). Difco agar was added to BG11 for the preparation of solid medium. When required, antibiotics were used at the following concentrations: neomycin, 10–50 µg ml–1; streptomycin 2–5 µg ml–1; and spectinomycin 2–5 µg ml–1. For the induction of heterocyst differentiation, filaments growing in BG11 medium were filtered through 0.2 µm pore size nylon filters, washed twice with BG110 medium (similar to BG11 but lacking NaNO3), inoculated in BG110 medium and cultured during 24 h at 30°C under continuous illumination. Cyanobacterial strains used in this work are described in Supplementary Table S1. Escherichia coli strain DH5α, used as a recipient for cloning, was grown in Luria–Bertani medium supplemented with antibiotics at a standard concentration when required (Ausubel et al. 2010). Plasmid construction Plasmids used in this work are described in Supplementary Table S2, and oligonucleotides used for PCR and other purposes are detailed in Supplementary Table S3. Plasmids were introduced into E. coli by transformation and into Anabaena by conjugal transfer as described (Elhai and Wolk 1988). Confocal and fluorescence microscopy For confocal microscopy of GFP fluorescence, Anabaena filaments expressing GFP were visualized with a Leica TCS SP2 confocal microscope using a HCX PLAM-APO ×63 1.4 NA oil immersion objective. GFP was excited with 488 nm irradiation from an argon ion laser. Fluorescence emission was monitored across windows of 500–540 nm (for GFP) and 630–700 nm (for cyanobacterial autofluorescence). To ensure that the signal observed in the 500–540 nm band corresponded to GFP and not to cyanobacterial pigments or degradation products, prior to every experiment filaments of the wild-type strain, not expressing GFP or GFP fusion proteins, were observed and microscope settings were selected so that no signal was detected in the 500–540 window. For fluorescence microscopy, Anabaena filaments were analyzed using a Leica DM6000B fluorescence microscope with ×100 and ×63 objectives. GFP fluorescence was monitored using a fluorescein isothiocyanate (FITC) L5 filter with an excitation range of 480/40 nm and an emission range of 572/30 nm. Prior to every experiment, filaments of the wild-type strain, not expressing GFP or GFP fusion proteins, were observed and microscope settings were selected so that no signal was detected at 572 nm. The cyanobacterial autofluorescence was monitored using a Texas red TX2 filter with an excitation window of 560/40 nm and emission window of 645/75 nm. Images were analyzed and convolved with the Leica Application Suite Advanced Fluorescence software and the Image J program (version 1.41). Fluorescence intensity was quantitated with the Image J program (version 1.41); values were processed with the Microsoft Excel Program (version 14.7) and plots were generated with Microsoft Excel or with Kaleidagraph (version 4.1). Fluorescence recovery after photobleaching (FRAP) Anabaena filaments expressing GFP–CAAD or AtpA–GFP were visualized by confocal microscopy as above using a Leica TCS SP5 confocal microscope. Cells were maintained in a chamber at 30°C during the FRAP assay. After an initial image was recorded, the bleaching was carried out by selecting a region of interest (ROI) of a fixed size. Bleaching was performed by exciting the GFP at 488 nm with an increase of the laser intensity by a factor of 8 and scanning the ROI during 0.137 s. After photobleaching, fluorescence recovery was monitored, acquiring images typically every 5 and 10 s or 1 min. Images were analyzed using the ImageJ version 1.41 software. Relative fluorescence intensity was calculated as described by (Kaňa 2013). The photobleaching depth was calculated as B = (Fi – F0)/F0, where Fi is the intensity of the region before bleaching and F0 is the intensity immediately after bleaching. For each time point (t), the relative fluorescence intensity (RFI) was calculated as RFI(t) = (Ft – F0)/(Fi – F0). For normalization, Fi was made equal to 1. Electron microscopy Filaments incubated in BG110 for 24 h were harvested by centrifugation, fixed with 2.5% (w/v) glutaraldehyde for 2 h at 4°C with gentle agitation and washed three times with 0.1 M cacodylate pH 7.4. Samples were stained with 0.5% (w/v) KMnO4 at room temperature for 1 h, and then washed, dehydrated with increasing concentrations of acetone and embedded in Spurr resin. Serial ultra-thin sections (50–100 nm) were prepared and deposited in formvar film-coated grids. All the samples were examined with a ZEISS LIBRA 120 PLUS electron microscope at 120 kV. Supplementary Data Supplementary data are available at PCP online. Funding This work was supported by Ministerio de Economía y Competitividad [grants BFU2013-44686P and BFU2016-77097P] and the European Regional Development Fund (FEDER). Abbreviations Abbreviations CAAD cyanobacterial aminoacyl-tRNA synthetase appended domain CURT1 CURVATURE THYLAKOID 1 protein family FRAP fluorescence recovery after photobleaching GFP green fluorescent protein ValRS valyl-tRNA synthetase Acknowledgements We are grateful to Dr. Antonia Herrero and Dr. Enrique Flores for providing Anabaena strains CSL108 and CSVT26 expressing GFP fusions of HetN and FraH, respectively. We are indebted to Alicia Orea (IBVF), Paloma Domínguez Giménez (Cabimer) and Juan Luis Ribas (CitiUS) for excellent technical assistance. We thank A. Herrero and E. Flores for stimulating discussions, and A. Herrero for a critical reading of the manuscript. Disclosures The authors have no conflicts of interest to declare. 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Google Scholar CrossRef Search ADS © The Author 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oup.com This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Plant and Cell Physiology Oxford University Press

Mechanisms for Protein Redistribution in Thylakoids of Anabaena During Cell Differentiation

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Oxford University Press
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© The Author 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oup.com
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0032-0781
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1471-9053
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Abstract

Abstract Thylakoid membranes are far from being homogeneous in composition. On the contrary, compositional heterogeneity of lipid and protein content is well known to exist in these membranes. The mechanisms for the confinement of proteins at a particular membrane domain have started to be unveiled, but we are far from a thorough understanding, and many issues remain to be elucidated. During the differentiation of heterocysts in filamentous cyanobacteria of the Anabaena and Nostoc genera, thylakoids undergo a complete reorganization, separating into two membrane domains of different appearance and subcellular localization. Evidence also indicates different functionality and protein composition for these two membrane domains. In this work, we have addressed the mechanisms that govern the specific localization of proteins at a particular membrane domain. Two classes of proteins were distinguished according to their distribution in the thylakoids. Our results indicate that the specific accumulation of proteins of the CURVATURE THYLAKOID 1 (CURT1) family and proteins containing the homologous CAAD domain at subpolar honeycomb thylakoids is mediated by multiple mechanisms including a previously unnoticed phenomenon of thylakoid membrane migration. Introduction The hydrophobic nature of some proteins or protein domains makes them suitable for insertion in biological membranes. However, the multiplicity of membrane systems within a cell requires the existence of mechanisms to allocate proteins to the proper lipid bilayer where they can accomplish their function (Teasdale and Jackson 1996, Derby and Gleeson 2007). Once inserted in the proper membrane, some proteins can be confined to a particular membrane domain, which involves mechanisms for the generation and maintenance of a compositional heterogeneity in the membrane. Compositional heterogeneity is conspicuous in chloroplast thylakoids, where protein complexes are confined in distinct membrane domains, i.e. PSII occupies the grana thylakoids while PSI and the ATP synthase are mostly in the stroma lamellae (Andersson and Anderson 1980, Dekker and Boekema 2005, Nevo et al. 2012). In chloroplast thylakoids, this phenomenon has been well characterized and is commonly referred to as lateral heterogeneity. However, how this uneven arrangement of protein complexes is established and maintained is not fully understood. It is also known that the distribution of protein complexes in the thylakoids is not static and may vary according to the conditions. For instance, light conditions promote state transitions or the PSII repair cycle, both involving long-range migration of membrane macrocomplexes (Joshua and Mullineaux 2004, Bellafiore et al. 2005, Bonardi et al. 2005, Nixon et al. 2005). Cyanobacteria are phylogenetically related to plant chloroplasts (Sagan 1967) and also contain thylakoids, which form an independent membrane system in the cytoplasm that is topologically equivalent to the stroma. However, thylakoids do not form grana or lamellae in cyanobacteria. On the contrary, thylakoid ultrastructure appears in general homogeneous within a species and no domains are distinguishable (Liberton et al. 2006, van de Meene et al. 2006, Nevo et al. 2007, Liberton and Pakrasi 2008, Liberton et al. 2011, Gonzalez-Esquer et al. 2016). An exception to this is observed in heterocysts of filamentous cyanobacteria where thylakoids are partitioned in two domains clearly discernible by their different appearance and subcellular distribution (Lang and Fay 1971, Wilcox et al. 1973, Giddings and Staehelin 1979). A domain known as honeycomb thylakoids is formed by highly contorted membranes densely packed at subpolar regions of the cell, while a second domain, referred to as peripheral thylakoids, is formed by membranes with a less convoluted appearance and a loose distribution in the cytoplasm (Lang and Fay 1971, Wilcox et al. 1973, Sherman et al. 2000). These membrane domains are not related in structure or function to the grana and lamella of chloroplast thylakoids but represent a suitable model to investigate the compositional heterogeneity of thylakoid membranes in cyanobacteria, an issue little investigated in these organisms. Heterocysts are cells specialized in nitrogen fixation that can be observed in multicellular filamentous species of cyanobacteria (Wolk 1996). Nitrogen scarcity triggers a regulatory cascade that provokes the differentiation of some vegetative cells of the filament into heterocysts. Differentiation is aimed at the physical separation into distinct cell types of two incompatible processes: oxygenic photosynthesis and nitrogen fixation. Thus, nitrogen-fixing filaments are composed of heterocysts that perform nitrogen fixation, separated by spacers of 10–15 vegetative cells that fix CO2 by photosynthesis. Heterocysts and vegetative cells exchange carbon and nitrogen compounds, thereby ensuring the correct nutrition of all cells in the filament (Flores and Herrero 2010). Differentiation induces extensive changes in the transcriptome, proteome, ultrastructure and metabolism of the cell (Lang and Fay 1971, Wilcox et al. 1973, Flaherty et al. 2011, Mitschke et al. 2011). Many of these changes are aimed at the expression of the oxygen-sensitive nitrogenase complex and the creation of a microoxic environment that preserves it from the oxygen produced by neighboring cells. In differentiating cells, most of the photosynthetic antenna complex is degraded, oxygen production at PSII is abrogated and photosynthetic CO2 fixation is blocked (Wolk 1996, Herrero et al. 2004). The large requirements of nitrogenase for reducing power and ATP are respectively fulfilled by the catabolism of sugars imported from vegetative cells and by the activity of F0F1-ATP synthase sustained by the electrochemical gradient created by cyclic electron transport around PSI under illumination (Magnuson and Cardona 2016). Another metabolic trait of heterocysts is a high oxygen detoxification activity based on terminal respiratory oxidases and flavodiiron proteins that directly reduce oxygen by the Mehler reaction (Pils et al. 2004, Valladares et al. 2007, Ermakova et al. 2014). Ultrastructural changes that occur during differentiation include the enlargement of the cell, the thickening of the cell wall by deposition of extra polysaccharide and glycolipid layers, the formation of polar granules made of a reserve polymer called cyanophycin and a complete reorganization of thylakoid membranes (Lang and Fay 1971, Wilcox et al. 1973, Giddings and Staehelin 1979). In the genera Anabaena and Nostoc, thylakoids are arranged in vegetative cells forming parallel layers at the periphery of the cytoplasm, the central part of the cytosol being mostly free from membranes and occupied by the nucleoid. Differentiation provokes an extensive rearrangement, which may involve synthesis of new membranes, so that in mature heterocysts thylakoids are partitioned into honeycomb and peripheral thylakoids. These two domains are not only different in localization and appearance, but evidence also indicates the existence of compositional heterogeneity, with proteins that specifically reside in only one of these two domains. However, the protein constituents of each domain remain to be fully characterized (Murry et al. 1981, Valladares et al. 2007, Cardona et al. 2009). Despite this information, many aspects about the formation of these two membrane domains and the distribution of proteins in each of them remain obscure. Furthermore, it is still not clear where to draw the line between honeycomb and peripheral thylakoids and whether they are physically interconnected. Photosynthetic electron transport is the major role of thylakoids, and most proteins anchored to or associated with these membranes are related to this function. However, the thylakoids of some cyanobacterial species also harbor proteins involved in gene translation, namely aminoacyl-tRNA synthetases (aaRSs). Membrane anchoring of these enzymes occurs through an extra domain called CAAD that contains two transmembrane segments. CAAD has an inherent capacity for directing proteins to the thylakoid membrane, not requiring any signal peptide (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016). Interestingly, CAAD is homologous to proteins of the CURVATURE THYLAKOID 1 (CURT1) family (Luque and Ochoa de Alda 2014). Proteins of this family have membrane-bending capacity and were shown to confer their characteristic shape to the thylakoids of Arabidopsis chloroplasts and the cyanobacterium Synechocystis (Armbruster et al. 2013, Heinz et al. 2016). In Arabidopsis, CURT1 proteins contribute to the lateral heterogeneity of thylakoids, showing a specific distribution at grana margins, where they induce acute curvature to the membrane (Armbruster et al. 2013). How proteins select their proper localization in the thylakoids of heterocysts is still an open question. We have analyzed the subcellular localization of thylakoidal proteins of Anabaena sp. PCC 7120 (also known as Nostoc sp. PCC 7120, hereafter Anabaena). Two distinct behaviors were observed that allowed their partitioning into two classes. One class is represented by the F0F1-ATP synthase and HetN, which in mature heterocysts occupied both honeycomb and peripheral thylakoids. A second class includes proteins specifically located at honeycomb thylakoids and includes valyl-tRNA synthetase (ValRS), CurT proteins and FraH. We have investigated how proteins of this second class select their specific localization at this particular domain. Results presented here indicate that multiple mechanisms contribute to the accumulation and specific confinement of some class 2 proteins at honeycomb thylakoids during heterocyst differentiation. Results Specific localization of proteins at discrete domains of the thylakoids in heterocysts Fig. 1A illustrates the ultrastructure of vegetative cells and heterocysts, evidencing the different appearance of the thylakoids in each cell type and their segregation in two domains in heterocysts (see also Lang and Fay 1971, Wilcox et al. 1973, Sherman et al. 2000). Honeycomb thylakoids were observed at a subpolar position in the vicinity of the cyanophycin granule, whereas peripheral thylakoids colonized other areas of the cytoplasm (Fig. 1A). Since these two domains are only discernible at late stages of differentiation or in mature heterocysts, the terms honeycomb (h) and peripheral (p) thylakoids will be used throughout this work to refer to membrane domains of mature heterocysts (or late pro-heterocysts). For premature differentiation stages, all membranes in the cytoplasm will be referred to as internal membranes. To investigate the distribution of proteins in h and p thylakoids of Anabaena heterocysts, the fluorescence of green fluorescent protein (GFP) fusions (see the Materials and Methods) was monitored by confocal microscopy. Mature heterocysts can be easily distinguished from surrounding vegetative cells by their enlarged body, their low autofluorescence and conspicuous refringent cyanophycin granules at both poles (Fig. 1B). Two distinct patterns were observed. The F0F1-ATP synthase complex and HetN showed fluorescent signals at subpolar regions, as well as discrete irregular signals elsewhere in the cytoplasm that varied in shape from cell to cell (Fig. 2A, B). The distribution of GFP-tagged ATP synthase (Santamaría-Gómez et al. 2016) in the tridimensional space of the heterocyst cytoplasm was analyzed in detail by z-axis montages (Fig. 2C) and 3-D image reconstruction (Supplementary Movie S1). Z-axis montages revealed that the subpolar signals showed a lenticular form whereas the non-polar irregular signals traversed the cytoplasm in distinct directions apparently forming bridges that connected the subpolar signals (Fig. 2C; Supplementary Movie S1). Tomography images corroborated the subcellular localization of the ATP synthase observed by z-montages (Supplementary Movie S2). These results indicated the localization of ATP-synthase and HetN in h and p thylakoids, which is consistent with previous reports showing decoration of both membrane domains with antibodies against the α- and β-subunits of ATP synthase (Sherman et al. 2000). Fig. 1 View largeDownload slide Cell structure of Anabaena cells. (A) Ultrastructure of vegetative cells and heterocysts in Anabaena. cw, cell wall; c, cyanophycin granule; t, vegetative cell thylakoids; n, neck. Size bar corresponds to 1 µm. (B) Bright field image (top) and confocal fluorescence image of the photosynthetic pigments of Anabaena filament (bottom). H, heterocysts; V, vegetative cell; c, cyanophycin granules. Confocal fluorescence images were obtained using a Leica TCS SP2 confocal microscope (see the Materials and Methods for details). Fig. 1 View largeDownload slide Cell structure of Anabaena cells. (A) Ultrastructure of vegetative cells and heterocysts in Anabaena. cw, cell wall; c, cyanophycin granule; t, vegetative cell thylakoids; n, neck. Size bar corresponds to 1 µm. (B) Bright field image (top) and confocal fluorescence image of the photosynthetic pigments of Anabaena filament (bottom). H, heterocysts; V, vegetative cell; c, cyanophycin granules. Confocal fluorescence images were obtained using a Leica TCS SP2 confocal microscope (see the Materials and Methods for details). Fig. 2 View largeDownload slide Subcellular localization of GFP fusion proteins in heterocysts. (A) Panels show the fluorescence of Anabaena derivative strains. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Other panels show the fluorescence of filaments expressing GFP fusion proteins. The name of the protein fused to GFP is indicated on the left of each panel. For each fusion, several panels are shown. The first panel (from left to right) corresponds to the fluorescence of GFP (λemision = 500–540 nm), the second panel corresponds to the autofluorescence of photosynthetic pigments (λemision = 630–700 nm), the third panel is the superposition of the two previous panels and the fourth panel is the superposition of the GFP fluorescence image and the bright field image. Plots on the right show the quantification of GFP fluorescence. Heterocysts are indicated with an ‘H’. In the plot, the image of the fourth panel is projected on the horizontal plane and the fluorescence of each area is represented on the vertical axis. Units of the x- and z-axis are microns and those of the y-axis are arbitrary units of fluorescence intensity. Size bars corresponds to 1 µm. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions (λemision = 500–540 nm) at these regions was quantified in heterocysts (n = 30) of filaments expressing each fusion protein. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units on the y-axis. Horizontal lines indicate the average value. (C) Z-axis montage of confocal microscopy images of cells expressing AtpA–GFP. Panels corresponding to serial layers of the preparation are presented in an ordered array. H, heterocyst; V, vegetative cell. Panels on the left show the fluorescence of the wild-type strain used as a negative control as in (A). Fig. 2 View largeDownload slide Subcellular localization of GFP fusion proteins in heterocysts. (A) Panels show the fluorescence of Anabaena derivative strains. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Other panels show the fluorescence of filaments expressing GFP fusion proteins. The name of the protein fused to GFP is indicated on the left of each panel. For each fusion, several panels are shown. The first panel (from left to right) corresponds to the fluorescence of GFP (λemision = 500–540 nm), the second panel corresponds to the autofluorescence of photosynthetic pigments (λemision = 630–700 nm), the third panel is the superposition of the two previous panels and the fourth panel is the superposition of the GFP fluorescence image and the bright field image. Plots on the right show the quantification of GFP fluorescence. Heterocysts are indicated with an ‘H’. In the plot, the image of the fourth panel is projected on the horizontal plane and the fluorescence of each area is represented on the vertical axis. Units of the x- and z-axis are microns and those of the y-axis are arbitrary units of fluorescence intensity. Size bars corresponds to 1 µm. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions (λemision = 500–540 nm) at these regions was quantified in heterocysts (n = 30) of filaments expressing each fusion protein. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units on the y-axis. Horizontal lines indicate the average value. (C) Z-axis montage of confocal microscopy images of cells expressing AtpA–GFP. Panels corresponding to serial layers of the preparation are presented in an ordered array. H, heterocyst; V, vegetative cell. Panels on the left show the fluorescence of the wild-type strain used as a negative control as in (A). In contrast, GFP fusions of other membrane proteins including ValRS, the two CurT homologs, Alr0805 and Alr4119, and FraH were detected as conspicuous fluorescent subpolar foci in mature heterocysts, with signals not being observed elsewhere in the cell, which indicated their specific localization at h thylakoids. A GFP fusion of CAAD, the domain that anchors ValRS to the membrane, showed the same localization pattern as full-length ValRS in heterocysts (Fig. 2A, B), which is consistent with previous observations (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016) and indicates that the determinants for the specific localization of ValRS in h thylakoids reside in the CAAD domain. These results allowed the partition of thylakoidal proteins into two classes: class 1, represented by the F0F1-ATP synthase and HetN; and class 2, which includes ValRS, Alr0805, Alr4119 and FraH. Fluorescence quantification clearly revealed two patterns corresponding to classes 1 and 2 (Fig. 2A, B). Furthermore, these observations lend support to the existence of a compositional heterogeneity between h and p thylakoids (Cardona et al. 2009, Magnuson and Cardona 2016), which entails the existence of mechanisms that promote the specific localization of proteins at particular membrane domains. An interesting observation was that the CurT proteins, Alr0805 and Alr4119, showed opposite patterns of expression: Alr0805 was abundant in vegetative cells and scarce in heterocysts, while Alr4119 was abundant in heterocysts and not detected in vegetative cells (Fig. 3). Here we propose to name these proteins as CurT1 (Alr0805) and CurT2 (Alr4119). The confinement of these proteins in h thylakoids suggests that they may be involved in conferring acute curvature to these membranes. Fig. 3 View largeDownload slide Differential expression profile of CurT proteins in Anabaena. Panels corresponding to composite images of cells expressing Alr0805–GFP or Alr4119–GFP from their native promoter (as indicated) are shown. Composite images of GFP fluorescence and the bright field image are shown on the left, and composite images of the red autofluorescence of photosynthetic pigments and GFP fluorescence are shown on the right. Some heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The three bottom panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Fig. 3 View largeDownload slide Differential expression profile of CurT proteins in Anabaena. Panels corresponding to composite images of cells expressing Alr0805–GFP or Alr4119–GFP from their native promoter (as indicated) are shown. Composite images of GFP fluorescence and the bright field image are shown on the left, and composite images of the red autofluorescence of photosynthetic pigments and GFP fluorescence are shown on the right. Some heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The three bottom panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Redistribution of GFP–CAAD during cell differentiation is a sequential process with two distinct phases Expression of the GFP–CAAD fusion described above had a low impact on the physiology of Anabaena and was chosen as a model to analyze how class 2 proteins localize specifically at honeycomb membranes. This fusion protein was considered a good representative of class 2 proteins since it reproduced the localization of ValRS (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016) and was homologous to CurT1 and CurT2 (Supplementary Fig. S1), which suggested that they would share the same mechanisms for their specific localization at h thylakoids. GFP–CAAD was observed uniformly distributed in the thylakoids of vegetative cells, whereas in heterocysts it was confined at h thylakoids (Fig. 2A) (Olmedo-Verd et al. 2011, Santamaría-Gómez et al. 2016). Therefore, this change in the subcellular distribution of GFP–CAAD must be completed by the 24 h that a vegetative cell takes to differentiate into a heterocyst. To gather information on the subcellular localization at intermediate stages of differentiation, GFP–CAAD was expressed in Anabaena from the heterocyst-specific patS promoter, which is induced early in differentiating cells (Yoon and Golden 2001), and GFP fluorescence was monitored at different time points after the initiation of heterocyst differentiation elicited by nitrogen step-down (see the Materials and Methods). At the initiation of the experiment, filaments showed no GFP fluorescence. After 6 h, cells regularly spaced in the filament, separated by approximately 10–20 cells, showed dim GFP fluorescence. These fluorescent cells were cells initiating differentiation, where the signal was observed at the cell periphery, co-localizing with the thylakoid membranes, detectable by the red fluorescent signal of photosynthetic pigments (Fig. 4A, B; Supplementary Fig. S2). At 12 h after the initiation, GFP–CAAD fluorescence remained at the cell periphery, but some accumulation was detected close to the poles of the cell, and 6 h later (18 h) the fluorescent signal was more intense at the poles and dimmer at the periphery (Fig. 4A–C; Supplementary Fig. S2). In contrast, no enrichment of the red signal at the poles was observed at 12 or 18 h (Fig. 4C; Supplementary Fig. S2). At 24 h, filaments contained mature heterocysts, where GFP fluorescence no longer remained at the cell periphery, being confined exclusively at a subpolar position (Fig. 4A, B). Fig. 4 View largeDownload slide Phases in the redistribution of GFP–CAAD during heterocyst differentiation. (A) Cultures of Anabaena expressing GFP–CAAD were subjected to nitrogen withdrawal and images were taken by confocal microscopy at different time points, as indicated. Details of the different panels are as in Fig. 2. Pro-heterocysts and heterocysts are situated at the center of each panel. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions in these regions was quantified in heterocysts (n = 20) of filaments expressing GFP–CAAD. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units in the y-axis. Horizontal lines indicate the average value. (C) Fluorescence intensity profiles of the periphery of differentiating cells. Panels correspond to the images shown on the right in (A). Fluorescence intensity of the green and red channels was quantitated in the zone covered by the yellow ring and plotted. The ring line starts at the white bar, which corresponds to the 0 point on the x-axis, and continues clockwise. (D) Fluorescence microscopy images of Anabaena cells expressing GFP–CAAD taken at 16 and 24 h after nitrogen compound withdrawal. Pro-heterocysts and heterocysts are situated at the center of each panel. Images were subjected to a deconvolution treatment to improve resolution (see the Materials and Methods). White arrowheads indicate the septal area between the heterocyst and adjacent vegetative cells. Plots correspond to the quantification of the fluorescence of GFP (green) and of photosynthetic pigments (red). Units of the x- and z-axis are microns. The y-axis indicates the relative fluorescence intensity in arbitrary units. Fig. 4 View largeDownload slide Phases in the redistribution of GFP–CAAD during heterocyst differentiation. (A) Cultures of Anabaena expressing GFP–CAAD were subjected to nitrogen withdrawal and images were taken by confocal microscopy at different time points, as indicated. Details of the different panels are as in Fig. 2. Pro-heterocysts and heterocysts are situated at the center of each panel. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (B) Plots represent the relative fluorescence of regions A, B, C and D as depicted in the diagram at the top. The fluorescence of GFP fusions in these regions was quantified in heterocysts (n = 20) of filaments expressing GFP–CAAD. Values of fluorescence were normalized with respect to the sum of the fluorescence of all four regions and represented as a dot plot. The relative fluorescence of each region is indicated in arbitrary units in the y-axis. Horizontal lines indicate the average value. (C) Fluorescence intensity profiles of the periphery of differentiating cells. Panels correspond to the images shown on the right in (A). Fluorescence intensity of the green and red channels was quantitated in the zone covered by the yellow ring and plotted. The ring line starts at the white bar, which corresponds to the 0 point on the x-axis, and continues clockwise. (D) Fluorescence microscopy images of Anabaena cells expressing GFP–CAAD taken at 16 and 24 h after nitrogen compound withdrawal. Pro-heterocysts and heterocysts are situated at the center of each panel. Images were subjected to a deconvolution treatment to improve resolution (see the Materials and Methods). White arrowheads indicate the septal area between the heterocyst and adjacent vegetative cells. Plots correspond to the quantification of the fluorescence of GFP (green) and of photosynthetic pigments (red). Units of the x- and z-axis are microns. The y-axis indicates the relative fluorescence intensity in arbitrary units. It is worth noting that signals observed at polar positions at 12 and 18 h were different in appearance and location from those observed at 24 h (Fig. 4A). To obtain a deeper insight on this issue, pro-heterocyst (16 h) and mature heterocysts (24 h) were analyzed by fluorescence microscopy followed by a deconvolution treatment of images to improve resolution (Fig. 4D; Supplementary Fig. S3; Supplementary Movies S3, S4). In pro-heterocysts (16 h), signals of GFP–CAAD at the cell poles were concentrated at discrete points very close to the septum with adjacent vegetative cells. Notice that the green fluorescence at the poles appears to overflow the red fluorescence signal (Fig. 4D, left panel; Supplementary Fig. S3; Supplementary Movie S3). In mature heterocysts (24 h), GFP–CAAD fluorescence was subpolar (closer to the center of the cell); the signal occupied a larger area and showed a crescent-like form (Fig. 4D, right panel; Supplementary Fig. S3; Supplementary Movie S4). These observations indicated that the redistribution of GFP–CAAD during differentiation is a sequential process that includes a long-lasting phase (0–18 h) where the GFP–CAAD concentration gradually decreases at the cell periphery and increases at the poles, very close to the septum; and a second phase (18–24 h) where the protein is shifted 0.3–0.5 µm toward the center of the cell (Fig. A–C). It is important to point out that in the first phase the dynamics of GFP–CAAD appear to be independent of the putative dynamics of the internal membranes (compare green and red signals in Fig. 4C and Supplementary Fig. S2). Mechanism for GFP–CAAD redistribution during cell differentiation Two non-mutually exclusive possibilities were considered for the GFP–CAAD redistribution in differentiating cells: (i) the migration of the protein within the cell or (ii) the replacement of the pre-existing protein by de novo synthesized molecules that are inserted specifically at h membranes. To test the first option, an assay was set up to monitor the dynamics of GFP–CAAD during differentiation in conditions where GFP–CAAD molecules are not synthesized de novo. For this, the PrbcL promoter, specific for vegetative cells and inactive in heterocysts (Elhai and Wolk 1990, Madan and Nierzwicki-Bauer 1993, Ramasubramanian et al. 1994), was chosen to direct the expression of reporter proteins. Since PrbcL is strongly repressed in differentiating cells (Elhai and Wolk 1990, Madan and Nierzwicki-Bauer 1993, Ramasubramanian et al. 1994), fluorescence signals in these cells would correspond to the GFP–CAAD molecules synthesized before the onset of differentiation. Monitoring the fluorescent signal in these cells would reveal whether GFP–CAAD molecules remain immobile or migrate within the cell during differentiation. To test the experimental design, a PrbcL–GFP construction was introduced into Anabaena. In nitrogen-rich medium, all cells of the filament showed intense fluorescence, consistent with the strong activity of the PrbcL promoter (Fig. 5A, central panel). When filaments were transferred to medium with no nitrogen compound, cells at semi-regular intervals, identified as differentiating cells by their larger size, showed a gradual decay of GFP fluorescence (Fig. 5A, right panel). The fluorescence decay was attributed to the absence of PrbcL activity and the degradation of GFP protein with time. In line with this, prolonged incubation times (>36 h) yielded a large proportion of heterocysts with no fluorescence signal. However, due to the great stability of GFP (in vitro half-life >26 h) (Corish and Tyler-Smith 1999), faint but detectable signals remained in pro-heterocysts (18–23 h) and mature heterocysts (24 h), validating this approach. In filaments expressing GFP–CAAD from PrbcL, very strong fluorescent signals of punctate appearance, mostly overlapping with the red fluorescence of thylakoids, were distinguished in vegetative cells (Fig. 5B; Supplementary Fig. S4). In contrast, in mature heterocysts (24 h), faint fluorescence signals were observed. Importantly, in virtually all mature heterocysts showing remnant fluorescence, these signals were at subpolar positions (93%, n = 57) (Fig. 5B; Supplementary Fig. S4). These results indicated that molecules synthesized previously (i.e. in the progenitor vegetative cell) had migrated during differentiation from the periphery to a subpolar position of the cell. Fig. 5 View largeDownload slide Diffusion of GFP–CAAD in internal membranes of heterocysts. (A) Cells expressing GFP from the PrbcL promoter were cultured in the presence of nitrate (central panel) or in the absence of nitrogen compounds (right panel). Heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The left panel corresponds to a composite image of wild-type filaments in the bright field and green fluorescence channels shown as a control of the absence of fluorescence in the 500–540 nm band. (B) Composite fluorescent images of Anabaena cells expressing GFP–CAAD from the PrbcL promoter. A heterocyst occupies the center of each panel. The size bar corresponds to 2 µm. The three panels on the left show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (C) FRAP analysis of a pro-heterocyst expressing GFP–CAAD. FRAP experiments were done using a Leica TCS SP5 confocal microscope (see the Materials and Methods for details). The first panel labeled as ‘Pre’ shows an image of the heterocyst before bleaching. Other panels correspond to images taken at the time indicated in seconds after bleaching. A white arrow indicates the bleached area. The plot on the right shows the evolution of the fluorescence signal during the experiment. Bleaching was performed at time = 0. The average and SD are indicated; n is the number of repeats of this experiment. (D) FRAP analysis of mature heterocyst expressing GFP–CAAD. Details are as in (C). (E) FRAP analysis of mature heterocyst expressing AtpA–GFP. Details are as in (C). (F) FRAP analysis of a mature heterocyst expressing GFP–CAAD. The entire region occupied by the heterocyst (indicated by a bracket in the left panel) was bleached and fluorescence recovery was followed for 300 s. Details are as in (C). Fig. 5 View largeDownload slide Diffusion of GFP–CAAD in internal membranes of heterocysts. (A) Cells expressing GFP from the PrbcL promoter were cultured in the presence of nitrate (central panel) or in the absence of nitrogen compounds (right panel). Heterocysts are indicated with white arrowheads. The size bar corresponds to 20 µm. The left panel corresponds to a composite image of wild-type filaments in the bright field and green fluorescence channels shown as a control of the absence of fluorescence in the 500–540 nm band. (B) Composite fluorescent images of Anabaena cells expressing GFP–CAAD from the PrbcL promoter. A heterocyst occupies the center of each panel. The size bar corresponds to 2 µm. The three panels on the left show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. (C) FRAP analysis of a pro-heterocyst expressing GFP–CAAD. FRAP experiments were done using a Leica TCS SP5 confocal microscope (see the Materials and Methods for details). The first panel labeled as ‘Pre’ shows an image of the heterocyst before bleaching. Other panels correspond to images taken at the time indicated in seconds after bleaching. A white arrow indicates the bleached area. The plot on the right shows the evolution of the fluorescence signal during the experiment. Bleaching was performed at time = 0. The average and SD are indicated; n is the number of repeats of this experiment. (D) FRAP analysis of mature heterocyst expressing GFP–CAAD. Details are as in (C). (E) FRAP analysis of mature heterocyst expressing AtpA–GFP. Details are as in (C). (F) FRAP analysis of a mature heterocyst expressing GFP–CAAD. The entire region occupied by the heterocyst (indicated by a bracket in the left panel) was bleached and fluorescence recovery was followed for 300 s. Details are as in (C). To test whether such migration of GFP–CAAD occurs by diffusion of this protein in the lipid bilayer, FRAP (fluorescence recovery after photobleaching) experiments were performed. FRAP has been extensively used to analyze the lateral movement of fluorescent proteins in the plane of thylakoid membranes (Mullineaux et al. 1997, Mullineaux and Sarcina 2002, Mullineaux 2004, Kaňa 2013). FRAP was first performed in pro-heterocysts expressing GFP–CAAD (16–20 h) by bleaching regions roughly at the equator of the cell (Fig. 5C, top panels) or at one of the poles (Fig. 5C, bottom panels). Recovery of fluorescence in the bleached area was observed in the order of seconds to minutes. Such recovery cannot be attributed to new protein synthesis since no recovery was observed in entirely bleached cells after 5 min (Fig. 5F). Therefore, FRAP results indicated that in pro-heterocysts GFP–CAAD can diffuse in thylakoids at different sites in the cell. Diffusion was tested in mature heterocysts (24 h), where GFP–CAAD is confined at h thylakoids, by bleaching the signal at one of the poles. No recovery was observed after 300 s (Fig. 5D), and similar results were obtained when half of the area was bleached, indicating that the protein is not mobile at h thylakoids. This lack of mobility is a specific feature of GFP–CAAD that may be shared by other class 2 proteins, but does not appear to apply to class 1 proteins, as recovery of AtpA–GFP fluorescence was observed when one pole of mature heterocysts was bleached (Fig. 5E). This latter observation has important implications as it lends empirical support to the existence of physical continuity between h and p thylakoids (Fig. 5E), an issue that so far has remained controversial. The shift in the position of GFP–CAAD from a polar to a subpolar position observed in the second phase (see above, Fig. 4) was intriguing. To obtain an insight into this issue, cells at late stages of differentiation (16–24 h) were analyzed by electron microscopy with a particular focus on polar and subpolar regions (Fig. 6A–D). In parallel, the subcellular localization of GFP–CAAD was monitored by fluorescence microscopy (Fig. 6E–L). In pro-heterocysts (16 h), the neck of the cell, a narrow enlargement of the cytoplasm at the poles that reduces the section of the septum in contact with the neighboring vegetative cell (Fig. 1A), was observed to be full of internal membranes (Fig. 6A). This was concomitant with the observation of GFP–CAAD as a strong fluorescent signal very close to the neighboring cell (Fig. 6E, I), suggesting that this protein could be colonizing the internal membranes located within the neck. At later time points, cyanophycin was observed to accumulate starting close to the septum and gradually filling the neck till it overflowed it, which results in the characteristic cup-like form of cyanophycin in mature heterocysts (Fig. 6B–D). The growth of the cyanophycin granule paralleled the changes observed for the GFP–CAAD signal at these stages of differentiation, including a displacement of about 0.3–0.5 µm toward the center of the cell and a reshaping of the signal from an intense concentrated dot to a more expanded crescent-like form, reminiscent of the shape of h thylakoids in mature heterocysts. Taken together, these observations suggested that at these stages of differentiation (16–24 h) the accumulation of cyanophycin dislodges internal membranes out of the neck of the heterocyst. It follows that in contrast to the first phase, the displacement of GFP–CAAD in this second phase would be sustained by a distinct phenomenon: the migration of membranes that carry this protein as an integral component. Fig. 6 View largeDownload slide Ultrastructure of cells in the last stages of differentiation, and subcellular localization of GFP–CAAD. Panels show cells in different stages of differentiation (A, E, I, 16 h; B, F, J, 20 h; C, G, K, 22 h; D, H, L, 24 h). (A–D) Electron microscopy of pro-heterocysts and heterocysts in different stages of differentiation as indicated above. The bracket indicates the neck of the cell, ‘c’ denotes cyanophycin. The size bar corresponds to 1 µm. Images (A), (B), (C) and (D) are representative of four, seven, six and nine different heterocysts, respectively. (E–H) Composite images of the bright field and GFP fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. (I–L) Composite images of the green and red fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. Images (E–L) are representative of at least 10 different heterocysts. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Fig. 6 View largeDownload slide Ultrastructure of cells in the last stages of differentiation, and subcellular localization of GFP–CAAD. Panels show cells in different stages of differentiation (A, E, I, 16 h; B, F, J, 20 h; C, G, K, 22 h; D, H, L, 24 h). (A–D) Electron microscopy of pro-heterocysts and heterocysts in different stages of differentiation as indicated above. The bracket indicates the neck of the cell, ‘c’ denotes cyanophycin. The size bar corresponds to 1 µm. Images (A), (B), (C) and (D) are representative of four, seven, six and nine different heterocysts, respectively. (E–H) Composite images of the bright field and GFP fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. (I–L) Composite images of the green and red fluorescence of cells expressing GFP–CAAD from the patS promoter. The size bar corresponds to 1 µm. Images (E–L) are representative of at least 10 different heterocysts. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. The results above illustrate that existing GFP–CAAD molecules can be displaced within the cell during the differentiation process, contributing to its accumulation in thylakoids at polar or subpolar positions. The fate of class 2 proteins synthesized de novo remains to be investigated. This issue was analyzed at late stages of differentiation once the two membrane domains, h and p thylakoids, are fully developed. In order to monitor only de novo synthetized molecules and avoid the interference by pre-existing ones, GFP–CAAD was expressed from a heterocyst-specific late promoter, PhetN. This promoter was reported to be induced about 17 h after nitrogen compound withdrawal (Bauer et al. 1997, Callahan and Buikema 2001, Flaherty et al. 2011). In our experiments, GFP–CAAD fluorescent signals started to be faintly visible 20 h after the initiation of differentiation (Fig. 7). As a control, cells expressing the HetN–GFP fusion protein from the PhetN promoter were also analyzed. Interestingly, mature heterocysts expressing HetN–GFP showed fluorescence at subpolar and peripheral positions, indicating its presence at h and p thylakoids (100%, n = 46). In contrast, virtually all cells expressing GFP–CAAD showed fluorescence exclusively at the subpoles (96%, n = 75). These observations indicated that the GFP–CAAD molecules synthetized de novo at the late stages of differentiation insert specifically at h thylakoids. Fig. 7 View largeDownload slide Subcellular localization of GFP–CAAD and HetN–GFP fusion proteins expressed from the late heterocyst-specific hetN promoter. Images correspond to cells bearing the construction indicated on the left. Bright field images (left) and composite fluorescence images (right) are shown. Heterocysts are indicated by white arrowheads. Size bars correspond to 20 µm. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Fig. 7 View largeDownload slide Subcellular localization of GFP–CAAD and HetN–GFP fusion proteins expressed from the late heterocyst-specific hetN promoter. Images correspond to cells bearing the construction indicated on the left. Bright field images (left) and composite fluorescence images (right) are shown. Heterocysts are indicated by white arrowheads. Size bars correspond to 20 µm. The three top panels show the fluorescence of the wild-type strain, used as a negative control for fluorescence in the 500–540 nm band. Before differentiation (i.e. in the progenitor vegetative cell), GFP–CAAD shows a peripheral distribution coincident with the position of thylakoids, whereas in mature heterocysts it is confined at h thylakoids at subpolar positions. Data presented above allow us to conclude that three different phenomena contribute to this redistribution: (i) protein diffusion in the thylakoid membrane; (ii); thylakoid migration; and (iii) specific insertion of de novo synthesized proteins at h thylakoids. Discussion Cyanobacterial thylakoids do not form grana and show a uniform appearance with no discernible domains (Gonzalez-Esquer et al. 2016). However, in heterocysts of filamentous cyanobacteria of the Anabaena and Nostoc genera, the separation of thylakoids into two domains was described decades ago (Lang and Fay 1971, Wilcox et al. 1973, Wolk 1996, Sherman et al. 2000). Despite this, many issues about the generation of h and p thylakoids remain unknown. For instance, it is still not clear whether the separation of these two domains occurs by remodeling of pre-existing thylakoids, if it requires the synthesis of new membranes, or both. Previous evidence suggested distinct functionality and protein composition for h and p domains (Murry et al. 1981, Valladares et al. 2007, Ferimazova et al. 2013). Cardona et al. could separate two fractions in their preparations of heterocyst internal membranes and showed that some proteins were specifically found in only one of them (Cardona et al. 2009). However, it is not clear whether these two fractions corresponded to h and p thylakoids. Therefore, the precise localization of proteins at either or both membrane domains remained mostly an open question. Results in this work add further evidence to the existence of a compositional heterogeneity between h and p thylakoids and allowed the partition of proteins into two classes according to their distribution in these membranes (Fig. 2). Some observations (Fig. 5C, E) indicate that there is continuity between h and p thylakoids in mature heterocysts. However, the uneven distribution of proteins in these two domains entails the existence of mechanisms for the generation and maintenance of compositional heterogenetity. We have investigated how class 2 proteins are specifically confined at h thylakoids. An important issue is how this specificity is established (i.e. how class 2 proteins can distinguish h from p thylakoids). A widespread mechanism for the specific placement of proteins at defined membrane areas is diffusion and capture, consisting of the free lateral diffusion of proteins in the bilayer until they interact with a trapping factor that can be a molecule (e.g. a lipid or protein partner already present at the destination) (Shapiro et al. 2009, Laloux and Jacobs-Wagner 2014) or a geometrical cue (e.g. membrane curvature) (Huang and Ramamurthi 2010). The redistribution of GFP–CAAD during differentiation could fit with this model as ab initio this protein is relatively mobile in the membrane but later on it becomes motionless in the h thylakoids (Fig. 5C). Though this lack of mobility may partially derive from macromolecular crowding or high viscosity of this membrane domain (Kirchhoff et al. 2004, Mullineaux 2008, Kaňa 2013, Kirchhoff 2014, Liu 2016), the observation that other proteins (i.e. AtpA–GFP) can diffuse in these membranes supports the existence of (a) specific trapping molecule(s) for GFP–CAAD. Alternatively, given the highly contorted nature of honeycomb membranes, class 2 proteins may localize there by direct sensing of membrane curvature (Huang and Ramamurthi 2010, Laloux and Jacobs-Wagner 2014, Strahl et al. 2015). Interestingly, CurT1 and CurT2 proteins contain predicted amphipathic helices in their N-termini (Supplementary Fig. S1B) and some reports have described that amphipathic helices have the ability to sense membrane curvature whereas other reports show that these helices induce membrane remodeling (see Drin and Antonny 2010, and references therein). In line with this, proteins of the CURT1 family from Synechocystis and Arabidopsis were found enriched at thylakoid regions of acute curvature and were shown to have the intrinsic ability to introduce curvature to liposomes in in vitro assays (Armbruster et al. 2013, Heinz et al. 2016). An interesting observation was the contrasting expression profile of CurT proteins in Anabaena (Fig. 3). CurT1 is expressed in all cells, although it is less abundant in heterocysts, which points to this protein as the housekeeping CurT in Anabaena. Both CurT1 and CurT2 are expressed in heterocysts, and are specifically found at h thylakoids in mature heterocysts (Fig. 2). This suggests that either the specific presence of CurT2, or the joint action of both proteins (i.e. by hetero-oligomerization) could be responsible for conferring the characteristic intricate shape to h thylakoids. Given the homology of the ValRS CAAD domain with proteins of the CURT1 family, it is tempting to speculate that this domain could also contribute to shaping h thylakoids. However, is worth noting that the CAAD sequence from Anabaena ValRS was not predicted to contain amphipathic helices (Supplementary Fig. S1B) (Luque and Ochoa de Alda 2014). Diffusion and trapping by a lipid or protein partner or by membrane curvature contributes to but it is not entirely responsible for the redistribution of GFP–CAAD during the differentiation of heterocysts. Data herein show that such a redistribution results from the additive effect of multiple phenomena (i.e. protein diffusion, membrane migration and specific insertion at polar membranes). However, these three distinct mechanisms appear to operate at successive stages of differentiation. At early stages (phase 1) the protein would freely diffuse in the membrane until at about 12 h an attracting or trapping element emerges at polar membranes (Figs. 4, 5C, D) and provokes accumulation of GFP–CAAD at membranes that fill the neck of the cell (Figs. 4D, 6A; Supplementary Fig. S3). In this phase, the redistribution of GFP–CAAD is apparently independent of putative membrane dynamics (Fig. 4C; Supplementary Fig. S2). At later stages (phase 2; 18–24 h), membranes containing GFP–CAAD would migrate out of the neck, expelled by the growth of the cyanophycin granule (Fig. 6). Hence, GFP–CAAD is passively transported to its final destination as an integral component of these membranes. Though global reorganization of thylakoids during differentiation was long suspected to involve the migration of membranes, to our knowledge this was not demonstrated. The use of GFP–CAAD as a marker of membrane localization has provided empirical evidence for the occurrence of this phenomenon in the neck of late pro-heterocysts. Our observations find further support in electron micrographs from previous studies that focused on other issues of differentiation (Lang and Fay 1971, Wilcox et al. 1973, Sherman et al. 2000, Zheng et al. 2017). Finally, in the last stages of phase 2, when the h domain is developed (in late pro-heterocysts and mature heterocysts), the de novo synthesized GFP–CAAD protein would insert specifically in such membrane domain, further contributing to the accumulation of GFP–CAAD protein in these membranes. It is not known whether this happens by in situ translation in ribosomes associated with h thylakoids (Matsumoto et al. 2015), or if the protein is synthesized by cytosolic ribosomes and then directed to these membranes by unknown factor(s). In any case, data presented in this work revealed the existence of sorting mechanisms to direct mRNA or proteins to specific domains of the internal membranes in mature heterocysts. The three mechanisms shown here to direct the relocalization of GFP–CAAD most probably operate for class 2 proteins such as ValRS, CurT1 and perhaps CurT2. However, this may not be the case for other class 2 proteins such as FraH (Merino-Puerto et al. 2011). In heterocysts, FraH is only expressed at late stages of differentiation. Thus, it is possible that subpolar accumulation of FraH occurs only by specific insertion of proteins synthesized de novo. Interestingly, fraH mutants lack h thylakoids, so FraH was suggested to have a role in the development or in the maintenance of this membrane domain (Merino-Puerto et al. 2011), which deserves further investigation. Cyanobacteria and plastids are highly ordered entities and their structure determines how they function (Jarvis and Lopez-Juez 2013, Gonzalez-Esquer et al. 2016). Proper arrangement of thylakoid proteins at specific membrane domains is important for normal functioning as well as for acclimation to particular conditions (Pribil et al. 2014). In addition, redistribution of some proteins in the thylakoids was shown to be important for acclimation and developmental phenomena, including the PSII repair cycle during photoinhibition, the placement of the PSII antenna during state transitions or the differentiation of plastids (Joshua and Mullineaux 2004, Bellafiore et al. 2005, Bonardi et al. 2005, Nixon et al. 2005,Jarvis and Lopez-Juez 2013). The differentiation of heterocysts, which can be regarded both as an acclimation response to nitrogen scarcity and a developmental phenomenon within the filament, is accompanied by a redistribution of thylakoid proteins. The consequent compositional heterogeneity reinforces the idea of a division of labor between h and p thylakoids and underlines the importance of the peculiar ultrastructure of heterocysts for their functioning. Results shown herein identify mechanisms responsible for the compositional heterogeneity of heterocyst thylakoids. We propose that similar mechanisms may operate jointly or individually for the establishment of compositional heterogeneity in thylakoids of other cyanobacteria or plant chloroplasts. Materials and Methods Organisms and growth conditions Anabaena sp. PCC 7120 and derivative strains were routinely grown in BG11 medium (Rippka 1988) at 30°C, under continuous illumination (75 photon µmol m–2 s–1), either continuously shaken or bubbled with a mixture of CO2 and air (1% v/v), and in this case supplemented with 10 mM NaHCO3; 1% (w/v). Difco agar was added to BG11 for the preparation of solid medium. When required, antibiotics were used at the following concentrations: neomycin, 10–50 µg ml–1; streptomycin 2–5 µg ml–1; and spectinomycin 2–5 µg ml–1. For the induction of heterocyst differentiation, filaments growing in BG11 medium were filtered through 0.2 µm pore size nylon filters, washed twice with BG110 medium (similar to BG11 but lacking NaNO3), inoculated in BG110 medium and cultured during 24 h at 30°C under continuous illumination. Cyanobacterial strains used in this work are described in Supplementary Table S1. Escherichia coli strain DH5α, used as a recipient for cloning, was grown in Luria–Bertani medium supplemented with antibiotics at a standard concentration when required (Ausubel et al. 2010). Plasmid construction Plasmids used in this work are described in Supplementary Table S2, and oligonucleotides used for PCR and other purposes are detailed in Supplementary Table S3. Plasmids were introduced into E. coli by transformation and into Anabaena by conjugal transfer as described (Elhai and Wolk 1988). Confocal and fluorescence microscopy For confocal microscopy of GFP fluorescence, Anabaena filaments expressing GFP were visualized with a Leica TCS SP2 confocal microscope using a HCX PLAM-APO ×63 1.4 NA oil immersion objective. GFP was excited with 488 nm irradiation from an argon ion laser. Fluorescence emission was monitored across windows of 500–540 nm (for GFP) and 630–700 nm (for cyanobacterial autofluorescence). To ensure that the signal observed in the 500–540 nm band corresponded to GFP and not to cyanobacterial pigments or degradation products, prior to every experiment filaments of the wild-type strain, not expressing GFP or GFP fusion proteins, were observed and microscope settings were selected so that no signal was detected in the 500–540 window. For fluorescence microscopy, Anabaena filaments were analyzed using a Leica DM6000B fluorescence microscope with ×100 and ×63 objectives. GFP fluorescence was monitored using a fluorescein isothiocyanate (FITC) L5 filter with an excitation range of 480/40 nm and an emission range of 572/30 nm. Prior to every experiment, filaments of the wild-type strain, not expressing GFP or GFP fusion proteins, were observed and microscope settings were selected so that no signal was detected at 572 nm. The cyanobacterial autofluorescence was monitored using a Texas red TX2 filter with an excitation window of 560/40 nm and emission window of 645/75 nm. Images were analyzed and convolved with the Leica Application Suite Advanced Fluorescence software and the Image J program (version 1.41). Fluorescence intensity was quantitated with the Image J program (version 1.41); values were processed with the Microsoft Excel Program (version 14.7) and plots were generated with Microsoft Excel or with Kaleidagraph (version 4.1). Fluorescence recovery after photobleaching (FRAP) Anabaena filaments expressing GFP–CAAD or AtpA–GFP were visualized by confocal microscopy as above using a Leica TCS SP5 confocal microscope. Cells were maintained in a chamber at 30°C during the FRAP assay. After an initial image was recorded, the bleaching was carried out by selecting a region of interest (ROI) of a fixed size. Bleaching was performed by exciting the GFP at 488 nm with an increase of the laser intensity by a factor of 8 and scanning the ROI during 0.137 s. After photobleaching, fluorescence recovery was monitored, acquiring images typically every 5 and 10 s or 1 min. Images were analyzed using the ImageJ version 1.41 software. Relative fluorescence intensity was calculated as described by (Kaňa 2013). The photobleaching depth was calculated as B = (Fi – F0)/F0, where Fi is the intensity of the region before bleaching and F0 is the intensity immediately after bleaching. For each time point (t), the relative fluorescence intensity (RFI) was calculated as RFI(t) = (Ft – F0)/(Fi – F0). For normalization, Fi was made equal to 1. Electron microscopy Filaments incubated in BG110 for 24 h were harvested by centrifugation, fixed with 2.5% (w/v) glutaraldehyde for 2 h at 4°C with gentle agitation and washed three times with 0.1 M cacodylate pH 7.4. Samples were stained with 0.5% (w/v) KMnO4 at room temperature for 1 h, and then washed, dehydrated with increasing concentrations of acetone and embedded in Spurr resin. Serial ultra-thin sections (50–100 nm) were prepared and deposited in formvar film-coated grids. All the samples were examined with a ZEISS LIBRA 120 PLUS electron microscope at 120 kV. Supplementary Data Supplementary data are available at PCP online. Funding This work was supported by Ministerio de Economía y Competitividad [grants BFU2013-44686P and BFU2016-77097P] and the European Regional Development Fund (FEDER). Abbreviations Abbreviations CAAD cyanobacterial aminoacyl-tRNA synthetase appended domain CURT1 CURVATURE THYLAKOID 1 protein family FRAP fluorescence recovery after photobleaching GFP green fluorescent protein ValRS valyl-tRNA synthetase Acknowledgements We are grateful to Dr. Antonia Herrero and Dr. Enrique Flores for providing Anabaena strains CSL108 and CSVT26 expressing GFP fusions of HetN and FraH, respectively. We are indebted to Alicia Orea (IBVF), Paloma Domínguez Giménez (Cabimer) and Juan Luis Ribas (CitiUS) for excellent technical assistance. We thank A. Herrero and E. Flores for stimulating discussions, and A. Herrero for a critical reading of the manuscript. Disclosures The authors have no conflicts of interest to declare. 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Plant and Cell PhysiologyOxford University Press

Published: Jun 6, 2018

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