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Article |
Endoplasmic reticulum remains continuous and undergoes sheet-to-tubule transformation during cell division in mammalian cells
Correspondence to Eija Jokitalo: eija.jokitalo{at}helsinki.fi
The endoplasmic reticulum (ER) is a multifaceted cellular organelle both structurally and functionally, and its cell cycle–dependent morphological changes are poorly understood. Our quantitative confocal and EM analyses show that the ER undergoes dramatic reorganization during cell division in cultured mammalian cells as mitotic ER profiles become shorter and more branched. 3D modeling by electron tomography reveals that the abundant interphase structures, sheets, are lost and subsequently transform into a branched tubular network that remains continuous. This is confirmed by observing the most prominent ER subdomain, the nuclear envelope (NE). A NE marker protein spreads to the mitotic ER tubules, although it does not show a homogenous distribution within the network. We mimicked the mitotic ER reorganization using puromycin to strip the membrane-bound ribosomes from the interphase ER corresponding to the observed loss of ribosomes normally occurring during mitosis. We propose that the structural changes in mitotic ER are linked to ribosomal action on the ER membranes.
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Introduction |
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ER plays a crucial role in synthesis, modification, transport, and degradation of membrane and secretory proteins, and is the site for biosynthesis, processing, and transport of several lipids, and for regulation of cytosolic calcium level (Sitia and Meldolesi, 1992). Considering the multitude of functions attributed to the ER, it is believed that the complexity of the ER is greater than the tripartite division suggests (Sitia and Meldolesi, 1992; Staehelin, 1997; Papp et al., 2003). Specialized subdomains have been suggested and in some cases shown to exist, but their composition and distribution is still mostly unclear (Geuze et al., 2003; Rizzuto et al., 2004; Robenek et al., 2004; Levine and Rabouille, 2005). Morphological aspects of the ER are very challenging to study because ER is a highly dynamic organelle that constantly rearranges its polygonal structure (Baumann and Walz, 2001). The first proteins involved in tubule generation, maintenance, and branch point generation, the reticulon and DP1 families and p22, have been identified (Andrade et al., 2004; Shibata et al., 2006; Voeltz et al., 2006), but sheet formation is still a mystery.
Little is known about the morphogenesis of the ER during cell division, or the partitioning of the NE or other functional subdomains in relation to, or within, the reticular ER in animal cells (Lowe and Barr, 2007). Although morphologically different in interphase, the distinction between the NE and the rest of the ER is less clear during cell division. Earlier models, which were mainly based on EM data, suggested complete fragmentation of the NE and fragmentation of the rest of the ER to a variable extent, yielding a heterogeneous population of fragments (Zeligs and Wollman, 1979; Warren and Wickner, 1996; Dreier and Rapoport, 2000; Du et al., 2004). On the other hand, FRAP experiments have shown that after the NE breakdown, proteins of the NE disperse into the ER, which remains continuous during cell division (Ellenberg et al., 1997). It is not known whether the NE proteins spread evenly throughout the ER, how subdomains within the reticular ER are partitioned, and how ER network organization itself changes during cell division.
There are few studies where changes in the overall structure of the ER during cell division have been systematically characterized, and it is evident that both structure and distribution of the ER within cells varies among different species and according to developmental stage or differentiation (Terasaki, 2000; Terasaki et al., 2001; Wollert et al., 2002; Bobinnec et al., 2003; McCullough and Lucocq, 2005; Poteryaev et al., 2005). These studies have shown that the ER undergoes dramatic changes during cell cycle reflecting the different needs of each cell type, and suggest that several mechanisms for partitioning have evolved. Study of cultured animal cells that have relatively simple signaling cascades and are unspecialized with respect of secretion or storage may reveal some basic fundamental mechanisms behind the partitioning of the ER and its subdomains.
In the present study, we analyzed the architecture of the ER in mammalian cells using a combination of light microscopy (LM) and EM techniques. We characterized the reticular ER in interphase and in dividing cells and the relation of NE and ER during cell division by quantitative morphometric assays, where tubule or cisternal length and the number of branch points were the two main features measured. We used two different approaches: confocal microscopy for live-cell visualization of the ER, and transmission electron microscopy (TEM) analysis of fixed cells to ensure higher resolution. In our TEM studies, we used thin sections for morphometric analysis, and for appreciation of the 3D structure, serial semi-thick sections were subjected to electron tomography (ET).
Our results show that the ER partitions as a continuous network and the NE partitions as part of the reticular ER. However, mitotic ER has a very different network structure compared to interphase ER. Morphometric assays revealed that it contains more branch points and shorter profiles than the interphase ER. By ET we were able to show that these changes in the network structure are accomplished by the transitioning of sheets into a tubular network. We provide mechanistic insight into the inheritance of the ER by showing that similar changes in the ER structure are induced by stripping of ribosomes with puromycin from the interphase ER. This is consistent with our observations on the decrease in ribosomes on the ER membranes during mitosis.
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Results |
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For confocal microscopy, we used live CHO-K1 cells stably expressing luminal collagen-specific chaperone Hsp47 (Nagata, 1998) coupled to EGFP (Kano et al., 2005; Hsp47-GFP). Interphase ER appeared as a continuous network that was denser around the nucleus than at the periphery of the cells (Fig. 1 A). Tubules close to the plasma membrane, and especially at the leading edge of migrating cells, seemed longer than elsewhere.
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When analyzing thin EM sections, one disadvantage is that cross-sections of sheets and tubules have similar profiles to tubules and vesicles, respectively. Therefore, the best way to appreciate the complex and pleiomorphic structure of the ER is to make 3D reconstructions by ET. For this, we imaged several relatively thick (200 or 250 nm) successive sections from cytochemically stained ssHRP-KDEL/CHO-K1 cells and produced models of ER by drawing the contours in the tomographic slices. Our models clearly demonstrated a continuity of the ER network in interphase cells. They also revealed the abundance of sheet structures that are otherwise difficult to observe by confocal microscopy or thin section EM. To study the impact of localization within the cell on the ER structure, we produced ER models from two different cell depths. A model derived from three bottom sections (illustration in Fig. 2, box A) demonstrates that the ER close to the plasma membrane as well as in the thin lamellipodia was composed of interconnected long tubules, and that sheets started to appear closer to the nucleus (Fig. 2 A, a and b). On the other hand, a model from the middle sections of the cell (illustration in Fig. 2, box B) showed abundant sheets in the central area of the cell and that tubules were mainly found in the periphery close to the plasma membrane and around the Golgi ribbon (Fig. 2 B). ET revealed three types of branch points, those between two sheets, a sheet and a tubule, or two tubules (Fig. 2 C, a–c; an animated model of sheet-to-sheet branch is included as Video 1, available at http://www.jcb.org/cgi/content/full/jcb.200705112/DC1).
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At EM level, the mitotic ER profiles appeared short and fairly evenly distributed, although excluded from the spindle area (Fig. 4, a–d). ER exit sites quickly reduced in number after early prometaphase and emerged again during telophase.
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The average number of branch points in interphase Hsp47-GFP/CHO-K1 cells was 84 per 100 µm2 (±12, n = 11 cells; Fig. 5 B, Table I). In mitotic cells, there were well over 100 branch points per 100 µm2. The increase was especially high in metaphase cells, showing 36% increase (115 ±13, n = 9). During earlier and later phases of mitosis, the increase ranged between 23 to 27% (104–107 branch points per 100 µm2). Similarly to the branch points, the total number of ER profiles was higher in mitotic cells than in interphase cells (Table I). Whereas interphase cells had 115 profiles per 100 µm2 (±12, n = 11) on average, the number increased to 138 and 136 profiles during early and late mitosis, respectively, and reached the average top value of 142 profiles (±14, n = 9) during metaphase. Distribution of profiles in different length groups indicated that the change was mainly due to increased number of short ER profiles in mitotic cells (Fig. 5 C, Table I). In the shortest category (0.2–0.4 µm), there were 26% more profiles in early mitotic, 31% in metaphase, 20% in anaphase, and 14% in telophase cells. In the next length groups of 0.4–0.8 µm and 0.8–1.5 µm, the increase in the number of profiles during all mitotic phases was between 22–30% and 13–25%, respectively. There were no significant changes in the longest profile category (>1.5 µm).
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Because the shape of the CHO-K1 cells changes from flat at interphase to round during cell division, we investigated whether the cell volume available for the ER diminishes, therefore forcing the network into denser configuration during mitosis. Our measurements from confocal section stacks showed that the cell volume was similar in interphase and mitotic cells. Interphase ER was confined to 80% of the cell volume, leaving 20% to the nucleus (±6%, n = 7). This ratio did not change in mitotic cells, being 78% for the ER and 22% for the chromosomes and mitotic spindle (±8%, n = 22). Thus, the ER takes up a similar volume of the cell throughout the cell cycle despite the changes in cell shape.
The morphometric analysis of the EM images confirmed the confocal microscopy results. The extent of ER network formation as measured by the number of branch points per ER area increased 39% from interphase (102 ± 26, n = 22) to metaphase (142 ± 31, n = 15) and dropped back to the interphase level upon mitotic exit (Fig. 5 D, Table II). Dividing cells had considerably larger number of short profiles and smaller number of long profiles than interphase cells. The change was most pronounced during metaphase, when the increase in short profiles outnumbered the decrease in long profiles such that the total ER profile number per ER area was increased approximately 70%. The sum of the lengths of all ER profiles per ER area, however, remained equal, 112 ± 15 µm per 100 µm2, at all phases of the cell cycle, and showed similar standard deviation within individual mitotic phase groups than between them (Table II).
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Denser ER network is accomplished by transformation of sheets to tubules
What kind of morphological changes can explain the increased numbers of branch points and short profiles in the mitotic ER? To elucidate this, corresponding tomographic models from interphase and dividing cells were done from upper middle sections (see illustration in Fig. 6 D) covering perinuclear area to plasma membrane in interphase cells and an area including some of the condensed chromosomes and reaching to the plasma membrane in dividing cells. In the modeled area of an interphase cell (Fig. 6 A), most of the ER appeared as sheets and only few tubular ER structures were found. Similar sheet structures were found in the tomogram of specimen sectioned perpendicular to the coverslip (Fig. 6 B).
Some of the sheets with low curvature extended through the whole tomographic volume (800 nm, an animated model of interphase ER is included as Video 2, available at http://www.jcb.org/cgi/content/full/jcb.200705112/DC1).
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Partitioning of the NE is subordinate to the ER network
In order to follow the fate of the NE during mitosis, we created CHO-K1 cell lines stably expressing the inner nuclear membrane marker lamin ß receptor coupled to EGFP (LBR-GFP; Ellenberg et al., 1997) or HRP. In both cases, we chose a clone that had a low expression level to avoid abnormal structures of the NE and leaking of the marker into the peripheral ER during interphase (Ellenberg et al., 1997; Ma et al., 2007). First, we re-examined the dispersal of the NE marker during cell division by live cell video and confocal microscopy. The video of four dividing LBR-GFP expressing cells (Video 4, available at http://www.jcb.org/cgi/content/full/jcb.200705112/DC1) shows how the marker is tightly kept on the NE during interphase and is redistributed to the reticular ER during cell division (cell 3 shown in Fig. S1, available at http://www.jcb.org/cgi/content/full/jcb.200705112/DC1) in agreement with the videos shown earlier using the same marker protein expressed in Cos7 cells (Ellenberg et al., 1997). During the mitotic NE dispersal, the fluorescent pattern became very faint. We attempted to quantify the structural changes of the LBR-GFP–positive membranes during mitosis using the method applied to Hsp47-GFP–expressing cells, but good skeleton models were not obtained because of the low expression level. Therefore, we continued the analysis using EM and peroxidase cytochemical staining of the LBR-HRP/CHO-K1 cell line. The NE of interphase cells was darkly stained, whereas the rest of the ER remained unstained (Fig. 7 a).
During prophase, the staining still appeared on the NE, although stained ER profiles were occasionally seen outside the NE (Fig. 7 b and close-up of boxed area in Fig. 7 c). As the NE breakdown started (Fig. 7 d) and progressed further (Fig. 7 e), the number of stained profiles outside the NE increased and conversely, the NE staining became weaker. The NE formed long, still thin and often layered membrane stretches that surrounded the chromosomes. The membranes also retained some nuclear pores even after the NE had been perforated at several points (Fig. 7, d and e). After the NE breakdown and until late anaphase, LBR staining seemed to be distributed all over the reticulum (Fig. 7, e–g). However, a closer look revealed that the majority of the cells had both unstained and stained ER profiles (Fig. 7, f and g).
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Next, the structure of NE during cell division was further analyzed by ET. For this, we used the same stable LBR-HRP/CHO-K1 cell line and ET was implemented as previously described to the corresponding areas from upper middle sections of the cell (illustration in Fig. 6). After the NE breakdown during prometaphase, the remnants of the NE could be seen as heavily stained profiles (see arrowheads in Fig. 8 A, inset). In a 3D model, these membranes appeared as small sheet structures that were thinner than the reticular ER and had still nuclear pores. A part of the reticular ER at this phase clearly appeared unstained. After the cell division progressed, the NE could no longer be morphologically distinguished, and the LBR-HRP was dispersed into the reticular ER (Fig. 8 B). During metaphase, the 3D structures of both LBR-HRP– and ssHRP-KDEL–marked ER membranes were indistinguishable from each other (Figs. 8 B and 6 C, respectively). In a telophase cell, the condensed chromosomes seemed to be surrounded by heavily stained NE membranes (Fig. 8 C, colored in red) that could occasionally be discerned already in anaphase cells (unpublished data). The small, thin sheet structures appeared to have nuclear pores (not seen in the picture). In all phases of cell division, the distribution of LBR-HRP was non-homogeneous, and the stained ER tubules were directly continuous with the unstained ones. The modeled ER network totally lacked sheet structures during metaphase (Fig. 8 B) and at later phases during cell division the only sheet structures found were the ones surrounding the condensed chromosomes (Fig. 8 C). Thus, the NE seemed to lose its identity during cell division while becoming a part of the ER network and thereby, the partitioning of the NE is subordinated to the ER network during cell division.
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Discussion |
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Our ET analysis of interphase ER revealed that sheets were predominant over tubules in the central area of the cell, whereas peripheral areas close to plasma membrane had long interconnected tubules. Both confocal and thin section EM quantifications showed that mitotic ER profiles are shorter and more branched in comparison to interphase ER. The structural change explaining these results was revealed by ET of mitotic ER: sheets were completely lost and transformed into branched tubular network.
These changes were correlated with the partitioning of the most pronounced subdomain of the ER, the NE, during mitosis. Studies concerning the partitioning of the NE at LM level have shown the spreading of the NE membrane proteins into and homogenous distribution within the ER network during mitosis (Ellenberg et al., 1997; Yang et al., 1997). Here, we confirm the redistribution of the inner nuclear membrane marker protein LBR-HRP into the ER network at the ultrastructural level, although the distribution of the label within the metaphase ER appeared somewhat more restricted than the almost homogenous distribution of the ssHRP-KDEL label (quantification data not depicted). We found frequent unstained tubules that were connected to stained ones. This may indicate that LBR is redistributed into distinct subdomains within the mitotic ER. The idea that NE building units would be kept as assembly-ready subdomains during mitosis is not completely new (Mattaj, 2004). However, we cannot make firm conclusions on this because the extent of HRP-induced staining depends heavily on the expression level of the protein. Fortunately, we were able to choose a low expressing cell line that does not induce NE invagination or membrane stacks, as reported in previous works (Ellenberg et al., 1997; Ma et al., 2007). Our estimations from Western blotting suggest that the total amount of LBR-HRP protein is approximately one third of the amount of ssHRP-KDEL protein (unpublished data). Whether this amount of protein could disperse and occupy the entire ER remains unanswered.
The structural changes of the ER from interphase sheets to highly branched tubular network after the onset of cell division is surprisingly radical and presumably requires the action of many molecular players. The first proteins involved in forming branch points and maintaining tubules have been found recently (for review see Shibata et al., 2006; Vedrenne and Hauri, 2006). Reticulons, especially the reticulon 4a (Rtn4a), and DP1, were suggested to function as tubule-forming hairpin proteins of the ER (Voeltz et al., 2006). Overexpression of Rtn4a abolished sheet structures from the interphase ER and created long, non-branching tubules instead. Therefore, reticulon overexpression or relative abundance at the onset of cell division might explain the loss of sheets from the mitotic ER.
We were able to convert interphase ER sheets into highly branched tubules effectively mimicking the mitotic ER morphology by stripping the membrane-bound polysomes from the interphase ER with a puromycin treatment of cells. Similar treatment with cycloheximide that inhibits protein synthesis without causing the dissociation of ribosomes did not induce structural changes to the ER. Furthermore, quantification of ribosomes on ER membranes of interphase and mitotic cells revealed a major stripping of ribosomes during mitosis; prometaphase to anaphase cells had 70% decrease in ER-bound ribosomes compared to interphase cells. Quantification was done on each segment of transversely sectioned ER on random images and ribosomes were counted individually. We analyzed altogether 2.8 mm of ER membrane, and values obtained for interphase cells agree well with previous quantifications done on cultured interphase Buffalo Rat liver cells (Seiser and Nicchitta, 2000).
These results suggest that polysomes are involved in stabilizing sheet structures of the ER. The mechanism behind is unclear, but there are some interesting possibilities. One alternative introduced recently (Shibata et al., 2006) is that large polysomes might not fit on tubules with high membrane curvature; therefore polysomes must trigger formation of sheets. However, tubulation of ER was observed when the inhibition of translation by cycloheximide was prolonged to 2 h. While this treatment stripped a significant part of ribosomes from the ER, deprivation of some sheet stabilizing protein during the 2-h cycloheximide incubation, but not yet within the 15-min incubation, cannot be excluded.
Puromycin treatment splits the ribosome in half (Seiser and Nicchitta, 2000). Bound only by the 60S subunit, calcium has been shown to leak through the translocon (Roy and Wonderlin, 2003; Van Coppenolle et al., 2004). Because ER is the source for Ca2+ -signals during mitosis and mitotic progression is blocked if the signals are inhibited (Parry et al., 2005), the structural resemblance of the puromycin treated interphase ER to mitotic ER might be explained by the resemblance of the cytosolic Ca2+ environments. Interestingly, Ca2+ efflux from the ER has been shown to accompany also the ER network formation in Xenopus vesicles in vitro (Voeltz et al., 2006). A likely candidate responding to the Ca2+ efflux is the evolutionarily highly conserved p22 (Andrade et al., 2004) that promotes branching depending on cytoplasmic Ca2+ level.
Our results concerning the structural changes occurring during cell division are not in agreement with some of the earlier published work. In sea urchin embryos and Xenopus oocytes the ER remained continuous during mitosis, but accumulated at the mitotic poles (Terasaki, 2000; Terasaki et al., 2001). No obvious transformation from sheets to tubules was observed. In the early Caenorhabditis elegans embryo and Drosophila melanogaster syncytial embryos the ER forms sheets during mitosis and cycles between dispersed and accumulated states (Bobinnec et al., 2003; Poteryaev et al., 2005). These cell cycle–induced changes may reflect the rapid cell cycle and emphasis on partitioning of the NE. In this study, we have provided evidence that in cultured mammalian cells the abundant interphase ER structures, sheets, are lost and transform into a branched tubular network that remains continuous. The amount of ER-bound ribosomes is significantly reduced during the middle phases of mitosis. We propose that tubulation provides a simple, yet effective, mechanism for partitioning of the ER in cultured mammalian cells, and that the method to achieve these changes is linked to the ribosomal action on ER membranes—ribosomal stripping and inhibition of translation in the long run lead to loss of sheets and appearance of tubules both in interphase cells and during cell division.
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Materials and methods |
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LM imaging
For quantification of the ER-bound S6 ribosomal protein, cells grown on 50 µg/ml fibronectin (Sigma-Aldrich)-coated coverslips were semi- permeabilized with 40 µg/ml digitonin (Seiser and Nicchitta, 2000), and cytosolic ribosomes were washed away with 1 M KCl for 2x 15 min on ice before fixation with PFA and immunostaining with S6-antibody (Jackson ImmunoResearch Laboratories). The cells were imaged with a UPlanApo 100x/1.35 (oil) objective in a wide-field AX70 Provis microscope using an F-View II CCD camera and AnalySIS software (Olympus) for 20 ms. The fluorescence intensity of S6 staining per cell was determined with ImagePro Plus 5.1 software (MediaCybernetics, Inc.).
For live-cell confocal imaging, cells were grown on fibronectin-coated glass-bottom dishes (MatTek). During imaging, the cells were kept in 40 mM Hepes-buffered growth medium at 37°C and 5 µM in vivo DNA stain DRAQ5 (Alexis Biochemicals) was added just before imaging when needed (Smith et al., 2000). For quantification, confocal image stacks were acquired with SP2 AOBS (Leica) using HCX PL APO 63x/1.2 W Corr/0.17 CS objective, red (HeNe 633 nm/10 mW) laser line for DRAQ5 and blue (Ar 488 nm/70 mW) for EGFP. The image stacks were collected through the cells and recorded by Leica confocal software, using parallel scanning, zoom 6, frame averaging 3 and 0.285-µm z-step. All stacks were deconvoluted with AutoQuant AutoDeblur 3D Blind deconvolution (Media Cybernetics, Inc.) using the same settings.
Imaging for co-compartmentalization analyses were done as above, but with HCX PL APO 63x/1.4–0.6 oil objective and lime laser (DPSS 561 nm/10 mW) for S6-staining using sequential scanning, 0.122-µm z-step and zoom 8. Deconvoluted stacks were analyzed with Imaris 5.7.1 Coloc tool using 5% threshold.
Quantification from confocal images
Every 2nd (interphase cells), 5th or 10th (mitotic cells) section through the confocal image stacks was analyzed, yielding an average of 3–4 sections or 4–5 sections per cell, respectively. A skeleton model of the ER network was formed with the branch and endpoint filter of ImagePro using thresholds 4–6 for CHO-K1/Hsp47-GFP and 4–20 for CHO-K1/ssGFP-KDEL depending on the brightness of individual cells. The cytoplasmic area containing ER and excluding areas devoid of ER was demarcated manually. This area, ER profile lengths, and branch points within the area were quantified with ImagePro. The resulting values from one cell were summed and related to the corresponding area, thus yielding an average value of the whole cell. Then, an average of all the cells within a mitotic phase was calculated.
To measure the volume where ER was confined, optical sections were chosen at regular intervals throughout the image stacks starting and ending with the top- and bottom-most sections of the cell. The ER area and ER-free areas, including the nucleus or chromosomes and the mitotic spindle, were traced by hand, digitized, and measured with ImagePro. The total volume with or without ER in the cell was then calculated by applying the Cavalieri principle (Gundersen and Jensen, 1987).
EM
For EM, cells grown on fibronectin-coated glass coverslips were cytochemically stained and flat embedded as described (Jokitalo et al., 2001). 80-nm- thick sections were cut parallel or perpendicular to the coverslip, post-stained with uranyl acetate and lead citrate, and observed with Tecnai 12 (FEI Corp.) operating at 80 kV. Images for quantification were acquired with ES500W CCD camera (Gatan Corp.) at 11,500x providing a pixel size of 6 nm. Cells for analysis were sampled randomly provided that their cell cycle phase could be reliably identified. The entire cellular profiles were imaged, and all stained ER profiles were traced manually by drawing the longest possible line from end to end along the middle of the profile onto transparencies, which were scanned and analyzed by ImagePro similarly to confocal images.
For ribosomal quantification, the cells were processed similarly, except that they were osmicated in non-reducing conditions and stained en bloc with 1% uranyl acetate, 0.3 M sucrose for 1.5 h at 4°C, and for data collection, the film camera of the microscope was used. ER membrane profiles of 60-nm sections at final magnification of 46,000x were traced along the transversely sectioned membranes. Ribosomes on them were counted individually and profile lengths were measured as above.
ET
For 3D ET, semi-thick (200 or 250 nm) sections were imaged with Tecnai FEG 20 (FEI Corp.) operating at 200 kV. Images for Figs. 6, 8, and 10 were collected with a 1k x 1k Multiscan 794 CCD camera (Gatan Corp.) at 3,500 or 5,000x, providing a pixel size of 5.42 nm or 3.68 nm, respectively, and for Fig. 2 with a 4k x 4k Ultrascan 4000 CCD camera (Gatan Corp.) at 5,000x, providing a 2x binned pixel size of 4.39 nm. For tilt series, the specimens were tilted at 1-degree intervals using a high tilt specimen holder (model 2020; E.A. Fischione Instruments) between ±65° or ±70°. Automated acquisition of the tilt series was carried out using either tomography software package provided by FEI Corp. or SerialEM software (Mastronarde, 2005). SerialEM enables automatic acquisition of montaged tilt series allowing tomography of larger areas, and by utilizing this feature, montages of 2 x 1, 3 x 1, or 2 x 2 frames were collected. For dual-axis tomography (Mastronarde, 1997), one tilt series was recorded from the area of interest, then the grid was rotated approximately 90° and a second tilt series from the same area was acquired.
The alignment of the tilt series was done with IMOD software package (Kremer et al., 1996) using 15-nm colloidal gold particles underneath or on top of the sections as fiducial markers. Tomographic reconstructions were analyzed and modeled with Amira (TGS Inc.). Before modeling the z-scale was elongated by the factor of 1.6 to compensate the compression of section by the electron beam.
Statistical analysis and image enhancements
Statistical significances of the differences between groups were calculated using single factor ANOVA in Excel 2003 (Microsoft). Contrast in images was enhanced by using the brightness and contrast tool of Adobe Photoshop 7.0.
Online supplemental material
Video 1 shows an animated model of a sheet-to-sheet branch from a 3D model of interphase ER shown in Fig. 6 A. Video 2 and Video 3 show z-series of the tomograms and 3D models of interphase and metaphase ER, respectively. Video 4 shows four dividing LBR-GFP/CHO-K1 cells and Fig. S1 still images of cell 3 of the video. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200705112/DC1.
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Acknowledgments |
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This work was funded by the Academy of Finland (projects 201198 and 115025) and University of Helsinki Research Funds. M. Puhka is a student of the Viikki Graduate School in Biosciences.
Submitted: 18 May 2007
Accepted: 5 November 2007
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