It is generally accepted that global translation varies during the cell cycle and is low during mitosis. However, addressing this issue is challenging because it involves cell synchronization, which evokes stress responses that, in turn, affect translation rates. Here, we have used two approaches to measure global translation rates in different cell-cycle phases. First, synchrony in different cell-cycle phases was obtained involving the same stress, by using temperature-sensitive mutants. Second, translation and DNA content were measured by flow cytometry in exponentially growing, single cells. We found no major variation in global translation rates through the cell cycle in either fission yeast or mammalian cells. We also measured phosphorylation of eukaryotic initiation factor-2α, an event that is thought to downregulate global translation in mitosis. In contrast with the prevailing view, eIF2α phosphorylation correlated poorly with downregulation of global translation and ectopically induced eIF2α phosphorylation inhibited global translation only at high levels.

It is one of the basic principles of cell proliferation that there is a link between general cell growth (protein synthesis) and cell-cycle regulation. Such a link is logical and has been hypothesized to exist, but its nature has been elusive. Protein synthesis is one of the most energy-demanding cellular processes and is, therefore, carefully regulated. It is a generally accepted view that global translation is considerably reduced during mitosis (reviewed by Sivan and Elroy-Stein, 2008). The reduction is thought to result from altered phosphorylation state of translation initiation factors. In particular, phosphorylation of the eukaryotic translation initiation factor 2 alpha (eIF2α) is induced after a number of different stresses and is thought to be the main reason for repressed translation. Cell-cycle-dependent downregulation of translation in G2/M phase was also attributed to increased eIF2α phosphorylation (Datta et al., 1999; Kim et al., 2014; Silva et al., 2015; Tinton et al., 2005).

Early translation measurements in synchronized mammalian cells revealed a 70% reduction of the global translation rate during mitosis (Fan and Penman, 1970). More recent studies using different synchronization methods suggested that the magnitude of the translation reduction depends on the method of synchronization (Coldwell et al., 2013; Shuda et al., 2015b). Also, studies using budding yeast indicated that the rate of protein synthesis is constant during the cell cycle (Elliott and McLaughlin, 1978; Elliott et al., 1979). More recent studies (in mammalian cells) have reported conflicting results regarding the level of translational reduction during mitosis (Shuda et al., 2015a; Stumpf et al., 2013; Tanenbaum et al., 2015), and the question of whether and to what extent global translation is downregulated during mitosis remains unanswered.

Measurement of translation in different cell-cycle phases is challenging because it often involves cell-cycle synchronization, which – in itself – can evoke stress responses that, in turn, will affect translation rates. Thus, the exact contribution of the synchronization method versus cell-cycle progression to any observed change in translation rates or the phosphorylation state of translation initiation factors is difficult to assess. Here, we use novel approaches to measure global translation rates during the cell cycle and whether they depend on eIF2α phosphorylation.

Global translation in synchronized cells

First, we utilized temperature-sensitive fission yeast mutants that arrest at different phases of the cell cycle. We synchronized the cells by shifting to the restrictive temperature before release into the cell cycle, achieving synchrony at different cell-cycle phases in response to the same treatment (i.e. temperature shift). Samples for analysis of DNA content, translation rate and eIF2α phosphorylation were taken every 20 min for 160 or 220 min after release from cell-cycle arrest. DNA content and translation rate were measured in single cells, by flow cytometry. Translation was assayed by pulse-labeling with the methionine analogue L-homopropargylglycine (HPG) (Knutsen et al., 2011), which is incorporated into growing polypeptide chains. It should be noticed that our assay addresses the regulation of global translation rather than the well-established translational regulation of individual proteins. To reveal small differences in signal intensity, the samples were barcoded (see Translation assays in Materials and Methods) and processed together in the very same solution. Barcoding involves staining each sample with an amino-reactive fluorescent dye that binds to amine functional groups present primarily on lysine side chains and at the N-terminus of a protein. Since the amount of dye that has labeled the protein is covalently attached to the cells, excess dye can be washed away. By staining cell samples with different concentrations of the dye, samples are imparted with unique dye-intensity distributions (Krutzik and Nolan, 2006). Barcoded samples were then pooled and processed together to measure translation rates. Using this approach eliminates differences due to technical issues and samples can be compared with great accuracy. The cdc10-M17 mutant was used to synchronize cells in G1, cdc25-22 was used to synchronize cells in G2 and nda3-KM311 was used to arrest the cells in mitosis. Phosphorylation of eIF2α was assessed by immunoblot analysis.

The rate of translation changed as the cells progressed from the block and through the cell cycle, apparently consistent with cell-cycle-dependent translation. However, the changes in translation rate followed the same pattern after release from the cell-cycle arrest regardless of when in the cell cycle the cells were synchronized (Fig. 1A–D, Fig. S1). At early time points the translation rate was low and after release it gradually increased to a rate above that measured before the shift. At late time points translation rates became similar to those measured in exponentially growing cells. There was no correlation between any particular cell-cycle phase and increase or decrease in translation rates. These results strongly suggest that global translation is not regulated in a cell-cycle-dependent manner and that the variations observed are caused by the synchronization.

Fig. 1.

Global translation in cells synchronized in the cell cycle. Cells of the indicated strains were grown exponentially at 25°C (A,B,E–G) or 30°C (C,D,H,I), incubated at 36°C or 20°C for one generation time and then shifted back to 25 and 30°C, respectively. Samples were taken at the indicated times after the shift. (A,C) Median intensities of the AF647 (HGP) signal normalized to that of exponentially growing cells. Average of three biological repeats and ± standard errors (±s.e.) are shown. (B,D) Illustrations of cell-cycle progression in the respective mutants. Fig. S1 shows the cell-cycle distributions. (E–I) Quantification of phosphorylated eIF2α normalized to tubulin in the indicated strains. Average and ±s.e. of three independent experiments are shown. Representative immunoblots are shown in Fig. S2.

Fig. 1.

Global translation in cells synchronized in the cell cycle. Cells of the indicated strains were grown exponentially at 25°C (A,B,E–G) or 30°C (C,D,H,I), incubated at 36°C or 20°C for one generation time and then shifted back to 25 and 30°C, respectively. Samples were taken at the indicated times after the shift. (A,C) Median intensities of the AF647 (HGP) signal normalized to that of exponentially growing cells. Average of three biological repeats and ± standard errors (±s.e.) are shown. (B,D) Illustrations of cell-cycle progression in the respective mutants. Fig. S1 shows the cell-cycle distributions. (E–I) Quantification of phosphorylated eIF2α normalized to tubulin in the indicated strains. Average and ±s.e. of three independent experiments are shown. Representative immunoblots are shown in Fig. S2.

To test the effects of a temperature shift, wild-type fission yeast cells were subjected to the same changes in temperature as employed to synchronize the cell-cycle mutants. Interestingly, translation rates followed the same pattern in wild-type cells and the cell-cycle mutants described above (Fig. 1A,C), demonstrating that the observed changes are due to the temperature shift rather than to the stage during the cell-cycle at which the particular mutant arrests. Furthermore, in itself, the change in temperature from 25 to 36°C and back to 25°C induced a transient delay of G2 onset (Fig. S1G), which is probably due to the previously described Rad3ATR-Rad9-dependent mechanism (Janes et al., 2012). Curiously, a shift from 30 to 20°C and back to 30°C also induced a cell-cycle delay, but in G1/S (Fig. S1H).

Phosphorylation of eIF2α was high at the early time points in the heat-sensitive mutants, then gradually diminished (Fig. 1E,F), regardless of where in the cell cycle the particular mutant was arrested. There was no correlation between eIF2α phosphorylation and any particular cell-cycle phase. We used α-tubulin as loading control, since quantification of the total eIF2α is difficult in fission yeast due to the lack of a good antibody. To exclude the possibility that the ratio of eIF2α to α-tubulin changes during the temperature-shift experiments and/or during the cell cycle, leading us to incorrect conclusions, we tested whether the amount of eIF2α varies as compared to α-tubulin. There was no change in the eIF2α:α-tubulin ratio (Fig. S2A,B), which allowed us to normalize the eIF2α-P signal to α-tubulin.

As a control, and to assess synchrony achieved in the above experiments, we followed expression of the G1 cyclin Cig2 by immunoblotting. The previously reported cell-cycle-dependent regulation was obvious in all three strains (Fig. S1), showing that the synchrony achieved in the above experiments allows us to detect cell-cycle-dependent changes in protein levels. Furthermore, the temperature shift resulted in increased levels of phosphorylated eIF2α also in the wild-type cells (Fig. 1G), confirming that such temperature shifts, routinely employed in cell-cycle synchronization experiments, invoke a stress response.

When cells were shifted from 20 to 30°C, changes in the amount of phosphorylated eIF2α were much less pronounced, be it in wild-type cells or the cold-sensitive nda3 mutant (Fig. 1H,I). Notably, the nda3 mutant arrests in metaphase, the very cell-cycle phase during which levels of phosphorylated eIF2α are thought to increase and contribute to a downregulation of translation. Furthermore, the biggest change in translation rate was observed in those cells shifted from 20 to 30°C, both for wild-type cells and the nda3 mutant (Fig. 1C), although this treatment resulted in the smallest change in the amount of phosphorylated eIF2α (Fig. 1H,I). These results directly contradict the prevailing view that eIF2α phosphorylation correlates with and is the reason for downregulation of global translation.

To assess the contribution of eIF2α phosphorylation to the observed changes in translation rates, strains carrying non-phosphorylatable eIF2α-S52A were used. Cell-cycle synchronization experiments and translation measurements were performed as above. Surprisingly, translation rates followed exactly the same pattern in the absence of eIF2α phosphorylation as in its presence; low immediately after the temperature shift, then recovering (Fig. 1A,C). Furthermore, in the heat-sensitive mutants translation was much more downregulated when eIF2α could not be phosphorylated (Fig. 1A).

We conclude that the changes in translation rates during the cell-cycle synchronization experiments were not due to cell-cycle-specific regulation of translation, but to the temperature shift itself. Furthermore, phosphorylation of eIF2α is not cell-cycle regulated and is not required for the downregulation of global translation after temperature shift.

Global translation in exponentially growing cells

Having seen no evidence of cell-cycle-dependent regulation of translation in synchronized cells, we set out to measure translation rates in different cell-cycle phases in non-synchronized cells. To this end, we measured HPG incorporation and DNA content in exponentially growing cells by flow cytometry. Cells in each cell-cycle phase were gated on two-parametric DNA cytograms (Knutsen et al., 2011) and HPG incorporation per cell was quantified in each cell-cycle phase. There were no significant differences in the rate of translation in the different cell-cycle phases (Fig. 2A,C). It should be noted that this method does not allow us to distinguish cells in mitosis from those in G1. Thus, a high translation rate in G1 cells might compensate for a reduced translation rate in the mitotic cells so that the relative translation rate for the mixed M-G1 population appears to be unchanged. However, in such a scenario the distribution of the HPG intensities in the M-G1 population would be broad, but this is not the case (Fig. 2A,C), which argues against this explanation. Another concern is that a low number of mitotic cells in the population would conceal a low translation rate in mitotic cells. To address this issue, cells of the M-G1 population were sorted onto microscopy slides and the microtubules were stained. At least 20% of the cells clearly contained a mitotic spindle (data not shown), demonstrating that the translation rates measured in the M-G1 population reliably represent those of mitotic cells. In addition, we analyzed exponentially growing fission yeast cells grown in a medium with isoleucine as sole nitrogen source. Under these conditions G1 is longer and cytokinesis occurs in G1 (Carlson et al., 1999), which allows us to distinguish a G1 population containing 1C DNA from mitotic cells. Also under these conditions, translation rates were similar in the different cell-cycle phases (Fig. 2B,D). These results obtained in non-synchronized, exponentially growing cells confirm that global translation does not vary significantly through the cell cycle.

Fig. 2.

Global translation in exponentially growing cells. (A,B) Two-parametric flow cytometry plots of fission yeast cells grown in EMM (A) or isoleucine-minimal medium (B). (C,D) Average of median intensity of the AF647 signal normalized to G2 (C) or G1 (D) from at least three biological repeats with ±s.e. Gating is shown in Fig. S2.

Fig. 2.

Global translation in exponentially growing cells. (A,B) Two-parametric flow cytometry plots of fission yeast cells grown in EMM (A) or isoleucine-minimal medium (B). (C,D) Average of median intensity of the AF647 signal normalized to G2 (C) or G1 (D) from at least three biological repeats with ±s.e. Gating is shown in Fig. S2.

Basic cellular processes, such as regulation of translation through the cell cycle are expected to be conserved in evolution, but the extent of such regulation might vary from organism to organism. Therefore, we investigated whether the level of global translation varies during the cell cycle in human cells. To this end, we measured translation rates in different cell-cycle phases in three different human cell lines. To measure translation, non-synchronized cells were pulse-labeled with the puromycin analogue O-propargyl-puromycin (OPP) and analyzed by flow cytometry. Cells in G1, S and G2 were identified on the basis of their DNA content and mitotic cells were identified using the mitotic marker histone 3 (H3) phosphorylated at serine residue 10 (phospho-S10-histone H3). The cell lines investigated were normal epithelial RPE cells immortalized by telomerase expression, the osteosarcoma-derived U2OS cells and cervix carcinoma-derived HeLa cells. There is a wide distribution of the intensity of the OPP signal in the G1 population, indicating that there are significant differences in translation rates among G1 cells. This feature is particularly obvious in normal RPE cells but less pronounced in the two cancer cell lines (Fig. 3). The G1 cells with lower translation rates might represent cells that have not yet passed the restriction point. There is a gradual increase in translation from G1 phase through S to G2 in all three cell lines, and a somewhat lower rate in mitotic cells. However, the rate of protein synthesis in mitotic cells is higher or similar to that in G1 cells and the extent of reduction from G2 to M ranges from 40% (RPE) to 15% (U2OS). Cells of each cell line were also fixed for microscopy after pulse labeling with OPP. Mitotic cells were identified by tubulin staining. Consistent with the results of flow cytometry, there are no major differences in the intensity of the OPP signal between mitotic and interphase cells (Fig. 3G).

Fig. 3.

Global translation through the cell cycle in human cells. (A–C) Two-parametric flow cytometry plots of the indicated cell lines. Yellow lines represent the mean intensity of AF647 (OPP) for each cell-cycle phase. (D–F) Bar graphs representing mean AF647 (OPP) intensity ±s.d. (G) Microscopic images of RPE, HeLa and U2OS cells labeled with OPP. Tubulin and DNA staining are shown for the identification of mitotic cells (arrowheads). (H) Quantification of phosphorylated eIF2α normalized to eIF2α in the indicated cell-cycle phases. Exponentially growing HeLa cells were fixed, and stained for H3-P and DNA content to identify cells in each cell-cycle phase. Then 50,000 cells from each phase were sorted to measure the level of phosphorylated eIF2α. Average and ±s.e. of three independent experiments are shown. Representative immunoblots are shown in Fig. S3. (I) Two-parametric flow cytometry plots of asynchronously growing and Nocodazole-arrested cells and cells 4 h after release from the Nocodazole block. (J) Bar graphs representing mean AF647 (OPP) intensity ±s.d. after Nocodazole block and release. Levels of phosphorylated eIF2α versus eIF2α phosphorylation are shown in Fig. S3.

Fig. 3.

Global translation through the cell cycle in human cells. (A–C) Two-parametric flow cytometry plots of the indicated cell lines. Yellow lines represent the mean intensity of AF647 (OPP) for each cell-cycle phase. (D–F) Bar graphs representing mean AF647 (OPP) intensity ±s.d. (G) Microscopic images of RPE, HeLa and U2OS cells labeled with OPP. Tubulin and DNA staining are shown for the identification of mitotic cells (arrowheads). (H) Quantification of phosphorylated eIF2α normalized to eIF2α in the indicated cell-cycle phases. Exponentially growing HeLa cells were fixed, and stained for H3-P and DNA content to identify cells in each cell-cycle phase. Then 50,000 cells from each phase were sorted to measure the level of phosphorylated eIF2α. Average and ±s.e. of three independent experiments are shown. Representative immunoblots are shown in Fig. S3. (I) Two-parametric flow cytometry plots of asynchronously growing and Nocodazole-arrested cells and cells 4 h after release from the Nocodazole block. (J) Bar graphs representing mean AF647 (OPP) intensity ±s.d. after Nocodazole block and release. Levels of phosphorylated eIF2α versus eIF2α phosphorylation are shown in Fig. S3.

Phosphorylation of eIF2α was investigated in HeLa cells. Non-synchronized cells were fixed, cell-cycle stage was analyzed as above and cells were collected by fluorescence-activated cell sorting (FACS). Phosphorylation of eIF2α was investigated in the different populations by immunoblotting. There were no significant changes in the levels of eIF2α phosphorylation during the cell cycle (Fig. 3H).

The above results strongly suggest that the previously observed, apparently cell-cycle-dependent, variation in translation rates was instead a result of synchronization. In order to directly address this, we synchronized HeLa cells by using Nocodazole and mitotic shake-off, and measured the translation rates. Consistent with previous studies, translation rates changed dramatically in Nocodazole-treatated cells (Fig. 3I,J) and levels of phosphorylated eIF2α increased upon Nocodazole-induced arrest (Fig. S3). However, in light of our results above, these dramatic changes are unlikely due to cell-cycle regulation but, rather, to the stress response following treatment with Nocodazole.

These findings strongly suggest that global translation rates are not dramatically downregulated in mitotic cells and that earlier studies overestimated the extent of variation through the cell cycle.

Phosphorylation of eIF2α and global translation

Surprisingly poor correlation was observed between the levels of phosphorylated eIF2α and global translation in the temperature-shift experiments. Previous work demonstrated that eIF2α phosphorylation can attenuate the translation of mRNAs (Harding et al., 2003; Hinnebusch, 1994). However, this is not the only consequence of eFI2α phosphorylation and the primary role might be to upregulate the translation of a specific class of mRNAs (reviewed by Dever, 2002).

To directly address the importance of eIF2α phosphorylation on global translation rates, we expressed eIF2α kinase 2 (EIF2AK2, hereafter referred to as PKR), one of the four human eIF2α kinases, in fission yeast and measured phosphorylated eIF2α and global translation rates. PKR expression was controlled by the regulatable nmt1 promoter, which is induced upon thiamine removal from the medium (Basi et al., 1993; Maundrell, 1993). We used two different versions of the promoter, resulting in two different expression levels of PKR. Cells were grown exponentially with the promoter repressed before PKR expression was induced, and global translation rates as well as eIF2α phosphorylation were measured during the first 24 h (in six generations) after induction. PKR expression was detected at 13 h after induction and levels of phosphorylated eIF2α reached their maximum at 16–19 h (Fig. 4A,B, Fig. S4). The extent of eIF2α phosphorylation induced by PKR driven by the weaker promoter was comparable to that induced by milder stresses (Fig. 4C, Fig. S4) but, surprisingly, we did not see any significant decrease in global translation rates in this case. The rate of translation remained similar to that before induction of PKR expression. (Fig. 4D). However, in cells that expressed PKR via the full-strength nmt promoter, translation was strongly reduced and, consistently, these cells did not form colonies when the promoter was derepressed (not shown). These results are consistent with previous findings, suggesting that extreme and lasting eIF2α phosphorylation can inhibit global translation and is lethal (Dever et al., 1993; Zhan et al., 2002). We conclude that the extent of eIF2α phosphorylation is crucial for its effect on downregulation of global translation. Very high levels of phosphorylated eIF2α block translation, but intermediate levels seem to have little influence on global translation.

Fig. 4.

Phosphorylation of eIF2α and global translation. Cells carrying the indicated plasmids were grown exponentially with the promoter repressed and one sample was taken to measure translation. The promoter was induced for the indicated times. (A,B) Quantification of phosphorylated eIF2α (eIF2α-P) phosphorylation normalized to α-tubulin at the indicated time points when PKR is expressed from the two different promoters. Notice the different scales on the y-axes. Representative immunoblots are shown in Fig. S4. (C) Quantification of phosphorylated eIF2α normalized to tubulin after the indicated stresses. Average and ±s.e. of three independent experiments are shown. Representative immunoblots are shown in Fig. S4. (D) Median intensities of the AF647 (HGP) signal normalized to that of exponentially growing cells (promoter repressed). Average of three biological repeats and ±s.e. are shown.

Fig. 4.

Phosphorylation of eIF2α and global translation. Cells carrying the indicated plasmids were grown exponentially with the promoter repressed and one sample was taken to measure translation. The promoter was induced for the indicated times. (A,B) Quantification of phosphorylated eIF2α (eIF2α-P) phosphorylation normalized to α-tubulin at the indicated time points when PKR is expressed from the two different promoters. Notice the different scales on the y-axes. Representative immunoblots are shown in Fig. S4. (C) Quantification of phosphorylated eIF2α normalized to tubulin after the indicated stresses. Average and ±s.e. of three independent experiments are shown. Representative immunoblots are shown in Fig. S4. (D) Median intensities of the AF647 (HGP) signal normalized to that of exponentially growing cells (promoter repressed). Average of three biological repeats and ±s.e. are shown.

Global translation rate changes little during the cell cycle

Many recent studies dispute the generally accepted view that global translation varies in a cell-cycle-dependent manner and is low during mitosis. Our results suggest that the discrepancies arise from experimental challenges. Studies of cell-cycle-related events often involve synchronization of cell cultures. In our work here, we employed temperature-sensitive yeast mutants. It should be noticed that studies on heat stress generally employ higher temperatures (>40°C) and the temperatures we used are close to those common in the natural environment of fission yeast cells. However, here we show that even the temperature shifts routinely used to synchronize the temperature-sensitive S. pombe mutants invoke a cellular stress response by themselves and influence global translation rates, supporting the idea that previously reported cell-cycle-dependent changes in translation rates are caused by the method of synchronization. Using the same stress to synchronize cells in different cell-cycle phases allowed us to separate the effects of cell-cycle progression from temperature shift on global translation rates. It is possible that, in our experiments, modest cell-cycle-dependent variations in global translation rates are concealed by imperfect synchrony. However, the synchrony achieved in the block-and-release experiments (Fig. S1) should have allowed us to observe the dramatic changes described previously. Furthermore, using flow cytometry to measure translation in exponentially growing cells allowed us to investigate global translation rates in different cell-cycle phases in non-stressed cells.

One caveat of analyzing the cell cycle of fission yeast by flow cytometry is that mitotic cells can only be identified after separation of the daughter nuclei, but cells in the early phases of mitosis cannot be distinguished from cells in G2. Thus, a reduction of global translation rates in metaphase would not be detected when using asynchronously growing cells and flow cytometry alone, although it would have been detected in the block-and-release experiments. Collectively, these data demonstrate that global translation is not significantly different between any of the cell-cycle phases in fission yeast cells.

In the human cell lines we also saw only small changes in the translation rate, consistent with recent studies reporting only minor variations. Mitotic cells were identified on the basis of phosphorylated histone H3, a mitotic marker that is present both in metaphase and anaphase. Notably, our approach did not involve any synchronization method, exposure to chemicals or changes in the cellular environment, which makes our results less subject to artifacts and technical problems. Furthermore, when we synchronized the cells, we also observed the previously reported variations, confirming the notion that the changes in translation are due to the synchronization-induced stress rather than cell-cycle progression.

Physiological levels of phosphorylated eIF2α do not significantly repress global translation

Under stressful conditions cells reduce the rate of global translation to conserve resources (Holcik and Sonenberg, 2005). At the same time, synthesis of proteins necessary to survive the stress is maintained or even increased. Many different forms of stress result in phosphorylation of eIF2α in eukaryotic cells (Clemens, 2001; Sonenberg and Hinnebusch, 2009), which it is thought to be required for both responses – downregulation of global translation and upregulation of translation of certain mRNAs. In addition, phosphorylation of eIF2α is also implicated in the cell-cycle-dependent regulation of translation. Here, we find that increased levels of phosphorylated eIF2α do not correlate with any particular cell-cycle phase but, rather, with the stress involved in synchronization, be it temperature shift or exposure to Nocodazole. We conclude that eIF2α phosphorylation is not regulated in a cell-cycle-dependent manner.

There is compelling evidence that eIF2α phosphorylation can attenuate the translation of mRNAs (Harding et al., 2003; Hinnebusch, 1994). The regulation of eIF2α phosphorylation is relevant for a number of diseases, such as neurodegenerative disorders, cancer and autoimmune diseases (Fullwood et al., 2012; Koromilas, 2015; Marchal et al., 2014; Ohno, 2014; Ravindran et al., 2016; Way and Popko, 2016). In all these fields, increased levels of phosphorylated eIF2α has commonly been taken to be a readout of reduced global translation. However, the two parameters have rarely been measured in the same experiment. Furthermore, at least under some stress situations other initiation factors can substitute for eIF2, as recently shown for eIF5B under hypoxia (Ho et al., 2018). Our results also suggest poor correlation between eIF2α phosphorylation and repressed global translation. First, eIF2α phosphorylation is clearly not required for the temperature-shift-induced downregulation of translation (Fig. 1), consistent with previous findings after UVC irradiation, oxidative stress and ER stress (Hamanaka et al., 2005; Knutsen et al., 2015; Shenton et al., 2006). Second, in the absence of eIF2α phosphorylation translation is repressed not less but rather more dramatically after temperature shift (Fig. 1). Third, ectopically induced eIF2α phosphorylation did not noticeably downregulate global translation in unstressed fission yeast cells, unless it was induced to high levels (Fig. 4). We suggest that the impact of phosphorylated eIF2α on global translation has been overestimated in the literature and that eIF2α phosphorylation cannot be used as a marker of downregulated translation. Our results demonstrate that the amount of phosphorylated eIF2α is crucial to determine whether it impacts on global translation and it has only a minor effect on the global translation at levels observed after mild stresses, mimicked by the expression of PKR from the weaker promoter in this study. In contrast, for the typical studies of stress responses involving the eIF2α kinases, often extreme conditions are employed, resulting in a severe block to global translation initiation. Under physiological conditions cells probably rarely experience these massive insults. Collectively, and as also suggested previously (Dever, 2002), these results imply that the main physiological role of eIF2α phosphorylation is not the downregulation of global translation but, most likely, the translation of certain mRNAs.

Cells and cell handling

All fission yeast strains used in this study are derivatives of S. pombe L972 h wild-type strain (Leupold, 1950). Strains used in this study are listed in Table S1.

Cells were maintained and cultured as previously described (Moreno 1991). The cells were grown in liquid Edinburgh minimal medium (EMM) with appropriate supplements at 25°C (or at 30°C for nda3-KM311cells) to a cell concentration of 2–4×106/ml. The cells were synchronized in G1 or G2 phase by incubating cdc10-M17 or cdc25-22 cells, respectively, at 36°C for 4 h (or 5 h for cdc10-M17 eIF2alphaS52A strain) before release into the cell cycle at 25°C; in M phase by incubating nda3-KM311 cells at 20°C for 4 h before release into the cell cycle at 30°C; and in early G2 by centrifugal elutriation as previously described (Hagan et al., 2016).

To obtain a population of mononuclear G1 cells, cultures were maintained at 30°C in minimal medium where NH4Cl was replaced with 20 mM L-isoleucine (Carlson et al., 1999).

Cultures of S. pombe transformants (together with a wild-type control culture) were grown to a cell concentration of 8×106/ml (OD595=0.4) in minimal medium where NH4Cl was replaced with 3.75 g/l L-glutamic acid, monosodium salt (Pombe Minimal Glutamate medium, PMG). To induce human PKR expression, cells cultured in PMG containing 5 µg/ml thiamine (Sigma-Aldrich) were harvested by centrifuging for 3 min at 1800 g, washed three times with PMG without thiamine, and resuspended in PMG lacking thiamine for the induction of nmt1 and nmt41 promoters.

Human U2OS and HeLa cells were cultivated in DMEM (Dulbecco's Modified Eagle's Medium) (Invitrogen) supplemented with 10% fetal bovine serum (FBS) (Gibco) and 1% penicillin/streptomycin (P/S) (Gibco). Human retinal pigment epithelium (RPE) cells immortalized with hTERT were cultivated in DMEM/F12 Glutamax supplement (Invitrogen) supplemented with 10% FBS, 1% P/S and 0.01 mg/ml hygromycin B (Sigma). Cells were tested for contamination once every two months.

Translation assays

To label newly synthesized proteins, 50 µM of L-homopropargylglycine (HPG, Thermo Fisher Scientific) was added to 1 ml samples of the main yeast culture taken out 10 min before the indicated time points. To stop translation, 0.1 mg/ml of cycloheximide (CHX) was added after 10 min. Cells were fixed in ice-cold methanol or 70% ethanol, washed in 0.5 ml TBS and barcoded using up to five different concentrations (450, 124.8, 31.2, 6.24 and 0.78 ng/ml) of Pacific Blue dye (PB; Thermo Fisher Scientific P10163) for 30 min in the dark at room temperature. Stained samples were then washed three times in 0.5 ml TBS and pooled. The samples were permeabilized with 0.5 ml 1% Triton X-100 in TBS and blocked with 1% BSA in TBS. To detect HPG, Alexa Fluor 647 was linked to the incorporated HPG in a ‘click’ reaction (Liang and Astruc, 2011) using the Click-iT cell reaction buffer kit (Thermo Fisher Scientific C10269) following the manufacturer's protocol to ligate the HPG alkyne with a fluorescent azide. Incorporation was quantified by using flow cytometry (LSR II flow cytometer, BD Biosciences). SYTOX Green dye (Thermo Fisher Scientific) was used to stain the DNA. Cell doublets were excluded from the analysis as described previously (Knutsen et al., 2011). Samples without HPG were used as negative controls. O-propargyl-puromycin (OPP), (Thermo Fisher Scientific) was added to 4 µM for 20 min, the cells were then trypsinized and fixed in 70% ethanol. To detect incorporated OPP, the fixed cells were washed once in PBS with 1% FBS. OPP was ligated with Alexa Fluor 647 in a ‘click’ reaction following the manufacturer's instructions. The samples were incubated for 5 min in detergent buffer [0.1% Igepal CA-630, 6.5 mM Na2HPO4, 1.5 mM KH2PO4, 2.7 mM KCl, 137 mM NaCl, 0.5 mM EDTA (pH 7.5)] containing 4% non-fat milk to block non-specific binding. The cells were incubated for 1 h with primary antibody against phospho-S10-histone H3 (1:500, Millipore 06-570) in detergent buffer containing 2% non-fat milk, washed once in PBS with 1% FBS and incubated for 30 min with Alexa Fluor 488-linked secondary antibody (1:500, Thermo Fisher Scientific A-11034) in detergent buffer. All incubations were carried out in the dark at room temperature. The cells were washed once in PBS with 1% FBS and stained with 1.5 µg/ml of Hoechst 33258 (Sigma) in PBS. The samples were analyzed using flow cytometry (LSR II flow cytometer, BD Bioscience, San Jose, CA) and data were analyzed using the FlowJo software (https://www.flowjo.com).

Fluorescence-activated cell sorting

Exponentially growing cells were fixed with 70% ethanol and stained for phospho-S10-histone H3 as described above and detected using Alexa Fluor 647-coupled secondary antibody. DNA was stained with 8 µg/ml propidium iodide. 50,000 cells from each cell-cycle phase were harvested using a FACS Aria II cell sorter.

UVC irradiation

Fission yeast cells were irradiated with 254 nm UV light (UVC) in a suspension in EMM (or PMG) medium under continuous stirring to ensure equal irradiation dose (Nilssen et al., 2003). The incident dose was measured with a radiometer (UV Products). A surface dose of 1100 J/m2 (at a dose rate of ∼250 J/m2/min) induces a checkpoint response but results in >90% cell survival. Samples for protein analysis were taken immediately after irradiation.

H2O2 treatment

Cells grown in PMG medium were treated with H2O2 at the indicated concentrations for 15 min before samples were taken.

Leucine starvation

An auxotroph strain was grown in PMG medium supplemented with leucine. The cells were washed with PMG medium three times and incubated in medium not containing leucine for the indicated times.

Immunoblotting

Total protein extracts of yeast cells were obtained using a low-salt buffer (25 mM MOPS pH 7.1, 60 mM β-glycerophosphate, 15 mM p-nitrophenyl phosphate, 15 mM MgCl2, 15 mM EGTA pH 8.0, 1 mM DTT, 0.1 mM Na3VO4, 1% Triton X-100) supplemented with protease inhibitors (Roche). Cell debris was removed by centrifugation at 14,000 g for 15 min at 4°C. The extracts were mixed with 4× LDS sample buffer (Thermo Fisher Scientific) and 50 mM DTT. Human cells were lysed in Laemmli sample buffer.

Cell extracts were separated on polyacrylamide gels, transferred onto PVDF membranes and probed with antibodies against phosphorylated eIF2α (1:750, CST 3398), eIF2α (1:1000, Santa Cruz sc-11386) PKR (1:3000, Abcam 32052), α-tubulin (1:30,000, Sigma-Aldrich T5168) and γ-tubulin (1:30,000, Sigma T6557). Signal intensities were quantified using ImageJ software.

We thank L. Lindbergsengen and M. O. Haugli for excellent technical assistance. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Author contributions

Conceptualization: B.G.; Methodology: V.S., B.G.; Validation: V.S.; Formal analysis: V.S.; Investigation: V.S., B.G.; Writing - original draft: B.G.; Writing - review & editing: V.S., E.B., B.G.; Supervision: B.G.; Funding acquisition: E.B., B.G.

Funding

We are grateful to Kreftforeningen (the Norwegian Cancer Society), Helse Sør-Øst RHF (the Norwegian South-Eastern Health Authority) and Radiumhospitalets Legater for funding.

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Competing interests

The authors declare no competing or financial interests.

Supplementary information