Endoplasmic reticulum remains continuous and undergoes sheet-to-tubule transformation during cell division in mammalian cells

ER undergoes morphological changes during cell cycle

We next analyzed the ER structure during different phases of the cell cycle. In all our experiments, we imaged naturally occurring mitotic cells in asynchronous cell populations. In confocal optical sections of mitotic cells (Fig. 3, b–e), ER resided at areas not occupied by the chromosomes and mitotic spindle and clearly retained the reticular form throughout mitosis. However, in contrast to interphase (Fig. 3 a), the network appeared denser and more evenly distributed.

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Figure 3.

Confocal microscopy analysis of the ER throughout the cell cycle. Confocal optical sections of CHO-K1 cells expressing Hsp47-GFP (a–e) or ssGFP-KDEL (f) were obtained throughout the cell cycle. DNA was stained in vivo (in lilac). The ER of an interphase cell (a) shows a typical polygonal structure with denser network in the perinuclear area than in the periphery of the cells. Arrowheads indicate long tubules found in the cell periphery. In contrast to the interphase ER, the mitotic ER was denser and more evenly distributed. The spindle region excluding ER is visible in metaphase (c) and late anaphase (d) cells. The other phases shown are prometaphase (b), early telophase (e) and metaphase (f). Bars, 5 μm; bar in image f applies to images b–f.

These results were confirmed with a cell line stably expressing EGFP coupled to the ER-targeting signal sequence and ER retention signal KDEL (ssGFP-KDEL). This ER-marker gave similar results as Hsp47-GFP (a metaphase cell shown in Fig. 3 f).

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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Figure 4.

Labeling of the ER network with peroxidase cytochemistry in mitotic ssHRP-KDEL/CHO-K1 cells. The ER network lacks long ER profiles in prometaphase (a), metaphase (b), late anaphase (c), and telophase (d) cells, and has many branch points especially during metaphase. The small insets show the cell cycle phase and the overall distribution of the ER profiles. ER branch points (arrows), mitotic Golgi clusters (G), chromosomes (Chr) and NE are indicated. Bars are 0.5 μm in high and 5 μm in low magnification images.

In order to accurately characterize the global changes in the ER network during cell division, we quantified two parameters from live cells: ER profile lengths including tubular and cisternal profiles (referred to as ER profiles) and the number of branch points (three types; see Fig. 2 C), which is a direct indicator of network formation (Lee and Chen, 1988; Waterman-Storer and Salmon, 1998; Dreier and Rapoport, 2000; Uchiyama et al., 2002). Confocal section stacks spanning cells were deconvoluted to subtract background noise, and the ER network was then converted into a skeleton model (Fig. 5 A, a–c; one optical section shown). The skeleton is a one-pixel-thick 2D model of the network in an optical section and allows direct measurement of the above parameters and their relation to the area occupied by the ER (ER area) in the same section. To obtain a reliable average value for each cell, we selected three to five sections at regular intervals to cover the whole cell depth. The minimum length of the ER profiles was set to 0.2 μm, which is the theoretical resolution limit of the diffraction limited LM.

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Figure 5.

Morphometric analysis of ER branch points and profile lengths. Quantification was done from confocal images of Hsp47-GFP (B and C) and from EM images of ssHRP-KDEL (D and E) expressing cells. (A) Stacks of optical sections (a, prophase section shown) were deconvoluted (b) and a skeleton model (c) of the ER network was formed by ImagePro software showing branch points in red and ER profiles in white. Bar, 5 μm. (B) The number of branch points per ER area increased during mitosis and reached a peak value in metaphase. The branch point numbers were statistically different from the interphase values (P < 0.01) in all phases of mitosis. (C) Mitotic cells showed a clear increase in short 0.2–0.4 and 0.4–0.8 μm ER profiles (P < 0.05 except for telophase 0.2–0.4 μm) as compared to interphase cells. INTER = interphase (n = 11 cells), PRO = prophase to prometaphase (n = 16), META = metaphase (n = 9), ANA = anaphase (n = 9), TELO = telophase (n = 7). (D) The number of branch points per ER area increased from interphase to metaphase and returned to the interphase level after mitotic exit. The branch point number in metaphase was statistically different from the interphase value (P < 0.001). (E) Mitotic cells had a significantly increased proportion of short ER profiles (0.040–0.400 μm) and reduced proportion of long profiles (>1.000 μm) as compared to interphase cells (P < 0.001). This pattern was most pronounced in metaphase cells. INTER (n = 22), PRO (n = 18), META (n = 15), ANA (n = 14), TELO (n = 18). The values are presented as the mean ± SEM.

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).

The quantification of ssGFP-KDEL–positive ER showed 36% increase in branch points in metaphase (132 ± 20, n = 5) as compared to interphase cells (97 ± 10, n = 3). In other phases of mitosis the change from the interphase was not as high as with the other marker protein. The number of ER profiles changed only during metaphase, in which there was an increased number of short profiles (unpublished data).

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).

The distribution of ER membranes in different length categories revealed that the two shortest groups had the highest number of profiles throughout the cell cycle and changed most during mitosis (Fig. 5 E, Table II). The largest proportion of profiles was taken up by the group of 0.150–0.400 μm profiles: 33% during interphase, around 40% in the groups of prophase and prometaphase and telophase, and more than 45% during meta- and anaphase. The increase in short ER profiles was balanced by decrease in long ER profiles. The largest decrease was observed in long 1.000–2.500-μm profiles: their proportion was reduced by more than 90% during metaphase and anaphase. As expected, the longest profiles starting from 2.500 μm and consisting almost entirely of the NE profiles were absent during meta- and anaphase and close to the interphase level during telo- and prophase. These results indicate that ER membranes are converted from long membrane structures into a large number of shorter highly branched structures in mitotic cells.

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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Figure 6.

Electron tomographic analysis of the ER in interphase and metaphase cells. Successive semi-thick sections (200 nm) from ssHRP-KDEL/CHO-K1 cells were subjected to ET. The modeled ER in the interphase cell appeared mostly as sheets both in sections cut parallel (A, combined from 3 sections) and perpendicular (B and Video 2, combined from 4 sections) to the coverslip. The structure of the ER in metaphase cell appeared tubular and lacked sheets (C and Video 3, combined from 2 sections). The insets are EM micrographs at low magnification to indicate the phase of the cell cycle. The illustration shows the depth of the modeled volume of cells. Note that A and B are from separate cells. Bars are 500 nm, and in insets 2 μm.

On the other hand, a modeled metaphase ER illustrated the dramatic changes in structure as it nearly completely lacked sheets (Fig. 6 C; an animated ER model of a metaphase cell is included as Video 3, available at http://www.jcb.org/cgi/content/full/jcb.200705112/DC1). ET revealed that mitotic ER is mostly tubular in structure, and that this tubular network remains continuous. Similar results, continuous tubular network with greatly reduced sheet structures, were obtained from a modeled ER of an anaphase cell (unpublished data). Thus, 3D ET analysis showed the continuity of ER during different phases of the cell cycle and indicated the morphological transformation from sheet structures of interphase cells into tubular structures in dividing cells.

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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Figure 7.

EM analysis of the NE throughout the cell cycle. Cytochemically stained LBR-HRP/CHO-K1 cells during interphase (a) and early prophase (b, boxed area enlarged in c), prometaphase (d and e), metaphase (f), anaphase (g), and telophase (h). The NE marker dispersed to the reticular ER after the NE breakdown and started to accumulate back to the reforming NE during late anaphase. Unstained ER tubules (arrowheads) were frequently observed next to or continuous with stained ER tubules (arrows) during all phases of mitosis. Cross-sections of nuclear pores (open arrows), chromosomes (Chr), NE, and centrioles (*) are indicated. Bars are 0.5 μm in high (c, d–h) and 5 μm in low magnification images.

The behavior of the LBR label during late anaphase or early telophase was essentially the reversal of early mitosis: the label accumulated on the reforming NE and was retained there (Fig. 7, g and h). The NE seemed to be rebuilt directly from bits of the surrounding reticulum and established its thin NE-like appearance only after sufficiently long membranes had formed on the chromosome surface.

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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Figure 8.

Electron tomographic analysis of the NE in mitotic cells. Successive semi-thick sections (200 or 250 nm) from LBR-HRP/CHO-K1 cells were subjected to ET. (A) In the prometaphase cell, the modeled ER (combined from 3 sections) showed mostly tubular structure, however, the remnants of the NE could be observed as darkly stained profiles (arrowheads in inset A). (B) During metaphase, the NE marker protein could be found dispersed in the tubular ER network (combined from 3 sections). (C) The telophase ER still had mostly tubular structure (combined from 4 sections), although the NE marker started to accumulate in the small sheet structures around the chromosomes. ER is modeled in yellow, darkly stained membranes in red, and chromosomes in light brown. The insets are EM micrographs indicating the cell cycle phase. Bars are 500 nm, and in insets 2 μm.

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 2× 15 min on ice before fixation with PFA and immunostaining with S6-antibody (Jackson ImmunoResearch Laboratories). The cells were imaged with a UPlanApo 100×/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 63×/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 63×/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 11,500× 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 46,000× 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, ​,8,8, and ​and1010 were collected with a 1k × 1k Multiscan 794 CCD camera (Gatan Corp.) at 3,500 or 5,000×, providing a pixel size of 5.42 nm or 3.68 nm, respectively, and for Fig. 2 with a 4k × 4k Ultrascan 4000 CCD camera (Gatan Corp.) at 5,000×, providing a 2× 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 × 1, 3 × 1, or 2 × 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.

We thank Hisao Kondo for the CHO-K1/Hsp47-GFP cell line and Jan Ellenberg for the pLBR-GFP construct, Jussi Jäntti and Dave Shima for critical reading of the manuscript, Catherine Rabouille and Hisao Kondo for helpful discussion, Mervi Lindman for expert technical assistance in specimen preparation for ET, and the personnel of EM unit for technical support.

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.