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Modulation of CHT7 Complexes during Light/Dark- and Nitrogen-Mediated Life Cycle Transitions of Chlamydomonas

Tomomi Takeuchi, Yang-Tsung Lin, Nicholas Fekaris, James Umen, Barbara B. Sears, Christoph Benning

Published December 2020. DOI: https://doi.org/10.1104/pp.20.00864

Abstract

The Chlamydomonas reinhardtii Compromised Hydrolysis of Triacylglycerols7 (CHT7) protein has been previously implicated in the regulation of DNA metabolism and cell-cycle-related gene expression during nitrogen (N) deprivation, and its predicted protein interaction domains are necessary for function. Here, we examined impacts of the cht7 mutation during the cell division cycle under nutrient deficiency in light-dark synchronized cultures. We explored the potential mechanisms affecting CHT7 complex activities during the cell cycle and N starvation, with a focus on the possible interaction between CHT7 and the C. reinhardtii retinoblastoma tumor suppressor (RB) protein homolog MAT3. Notably, the absence of CHT7 did not negatively impact the synchrony of cell division and cell cycle progression during diel growth. Although the majority of CHT7 and MAT3/RB proteins were observed in separate complexes by blue native-PAGE, the two proteins coimmunoprecipitated both during synchronized growth and following N deprivation, suggesting the presence of low abundance subcomplexes containing CHT7 and MAT3/RB. Furthermore, we observed several phosphorylated isoforms of CHT7 under these conditions. To test the potential role of phosphorylation on the structure and function of CHT7, we performed site-directed mutagenesis of previously identified phosphorylated amino acids within CHT7. These phosphorylated residues were dispensable for CHT7 function, but phosphorylated variants of CHT7 persisted, indicating that yet-unidentified residues within CHT7 are also likely phosphorylated. Based on the interaction of CHT7 and MAT3/RB, we postulate the presence of a low-abundance or transient regulatory complex in C. reinhardtii that may be similar to DREAM-like complexes in other organisms.

Many of the life cycle decisions of the photosynthetic unicellular alga Chlamydomonas reinhardtii are dictated by the diurnal signals present in the environment, such as the daily cycles of alternating light and dark periods. Under photoautotrophic diel cycles, the vast majority of the biological processes, including cellular growth and division, become temporally ordered or synchronized in C. reinhardtii cells (Zones et al., 2015). C. reinhardtii cells divide using a modified cell cycle termed multiple fission, where cells increase in volume during the day and undergo successive rounds of S/M (synthesis/mitosis) phases and divisions during the night (Spudich and Sager, 1980; Cross and Umen, 2015). These growth and division events likely became temporally segregated such that flagella-mediated phototaxis and energy-production by photosynthesis are maximized when light is available, while mitosis and cytokinesis can proceed in the dark when loss of flagellar motility has a lesser impact on fitness (Spudich and Sager, 1980; Bišová and Zachleder, 2014; Cross and Umen, 2015; Zones et al., 2015). In multiple fission, cells undergo a prolonged G1 phase, where they enlarge many times their initial volume without the initiation of mitosis (Umen and Goodenough, 2001). Upon reaching a critical volume, cells pass a size-regulated checkpoint called Commitment and complete at least one round of division even when there is no further increase in volume (Spudich and Sager, 1980; Craigie and Cavalier-Smith, 1982; Donnan and John, 1983; Umen and Goodenough, 2001). To ensure equally sized daughters are produced, the number of S/M phases and divisions that the mother cells undergo are tightly linked to their cell sizes in C. reinhardtii (Craigie and Cavalier-Smith, 1982; Donnan and John, 1983).

In many organisms, a conserved transcriptional regulatory module, termed DREAM (DP, RB, E2F, and Myb-MuvB), whose activities are determined by the combinatorial presence of distinct components, is responsible for mediating the transcriptional regulation of cell cycle genes in the context of development, metabolic adjustment, or response to the environment (Korenjak et al., 2004; Lewis et al., 2004; Harrison et al., 2006, 2007; Georlette et al., 2007; Litovchick et al., 2007; Pilkinton et al., 2007; Schmit et al., 2007; Tabuchi et al., 2011; Kobayashi et al., 2015). Orthologous DREAM-like complexes have also been identified in Arabidopsis (Arabidopsis thaliana), and they function to regulate cell cycle gene expression during plant growth and development (Kobayashi et al., 2015). C. reinhardtii also has a homolog of the retinoblastoma tumor suppressor (RB) protein, called MAT3, in addition to the E2F1 transcriptional activator and its dimerization partner, DP1 (Bišová et al., 2005; Olson et al., 2010). These RB pathway proteins are also involved in the control of cell size and cell division in C. reinhardtii (Umen and Goodenough, 2001; Fang et al., 2006; Olson et al., 2010). The mat3 mutant cells are smaller in size than wild-type cells since they pass the commitment point at a smaller volume and also undergo more rounds of the S/M phases than the wild type (Umen and Goodenough, 2001; Fang et al., 2006). A class of cyclin-dependent kinase, CDKG1, has been identified as one of the regulators responsible for coupling the mother cell size to the number of subsequent divisions by phosphorylating MAT3 in a cyclin d-dependent manner (Li et al., 2016). Furthermore, the conserved cyclin-dependent kinase and CDK1 ortholog, CDKA1, has recently emerged as an essential regulator of critical cell size and commitment in C. reinhardtii, by mechanisms distinct from those utilized by the RB pathway proteins (Cross, 2020).

Another group of proteins that the previously characterized DREAM complexes of both animals and plants share is a family of proteins that contains two Cys-rich motifs hereafter collectively referred to as a CXC domain. CXC domains are reported to bind DNA in a sequence-specific manner in metazoans (Beall et al., 2002; Schmit et al., 2009; Fauth et al., 2010; Tabuchi et al., 2011; Zheng et al., 2014; Marceau et al., 2016), but CXC domain proteins are much less characterized in plants. One CXC domain-containing protein in C. reinhardtii with a characterized function in the nutrient-responsive regulation of cell division is the Compromised Hydrolysis of Triacylglycerols7 (CHT7) protein (Takeuchi et al., 2020). The cht7 mutant was initially isolated due to its defects in the remobilization of triacylglycerols (TAGs) and delay in the resumption of growth following a period of nitrogen (N) deprivation and refeeding (Tsai et al., 2014). The original cht7 mutant was isolated in a cell-wall-less parental strain, dw15, and exhibited gene expression phenotypes during mixotrophic N-replete growth and N refeeding after N deprivation (Tsai et al., 2014; Tsai et al., 2018).

Because cell-wall-less strains are more susceptible to stress, they may accumulate additional genetic changes that could alter their stress response phenotypes and/or cell cycle regulation compared with wild-type walled strains. To better understand the impacts of the CHT7 mutation, the cht7 mutant was previously backcrossed to two walled strains, 21gr and 6145c, and recharacterized (Takeuchi et al., 2020). Notably, in a walled strain background and in the absence of N, CHT7 was necessary for the full repression of cell-cycle-dependent genes as cells entered quiescence, or a reversible and temporary arrest of growth and cell division, and for the survival of cells under N starvation (Takeuchi et al., 2020). Many cht7 mutant cells failed to cease division following the withdrawal of N, and they were unable to properly resume cell division when N was again provided in the medium (Takeuchi et al., 2020). Furthermore, the C-terminal disordered region with the predicted protein interaction domains, and not the presumed CXC domain implicated in DNA-binding, was essential for the function of CHT7 (Takeuchi et al., 2020). A CHT7 complex of constant Mr was observed both during N-replete growth and following N deprivation by blue native-PAGE (BN-PAGE; Tsai et al., 2014), and both CHT7 and MAT3 were observed in the nucleus (Olson et al., 2010; Tsai et al., 2014). However, how CHT7 function is mediated at the molecular level and whether a complex orthologous to DREAM exists in C. reinhardtii remain unknown.

Here, we assessed the impact of the cht7 mutation on the cell division cycle using N-replete synchronized cht7 cells cultured under 12 h light:12 h dark diel cycles. We explore the potential mechanisms by which the transcriptional or gene regulatory activity of CHT7 is controlled at the levels of the protein, both during N-replete light-dark synchronized growth and following N deprivation. In particular, we investigated the interactions between CHT7 and MAT3/RB, as well as their bulk complexes, and the physiological functions of CHT7 phosphorylation. The results here suggest the potential existence of DREAM-like complexes in C. reinhardtii.

RESULTS

Synchronization of Cell Division in the cht7 Mutant and Its Complemented Line under N-Replete Alternating Light-Dark Cycles

To test whether the absence of CHT7 affects the synchrony of cell division, cells from the wild-type 6145c, cell-walled cht7 (1) (hitherto referred to as cht7; Takeuchi et al., 2020) and its C terminally HA-tagged CHT7 complemented line, CHT7-HA:cht7, were grown photoautotrophically in high-salt (HS) medium under alternating 12 h light:12 h dark cycles in environmental photobioreactors (ePBRs; Lucker et al., 2014). All cultures were maintained in steady state at a density equivalent to midlog growth using a turbidostat prior to the start of the experiment, and the extent of cell division synchrony for each line was assessed qualitatively by microscopy (Fig. 1A) and quantitatively by measuring the cell size distribution (Fig. 1B) at different times during the diel cycle. Beginning at the start of the light period (Zeitgeber 0 [ZT 0]), the microscopic images showed the cells from all three lines, including the cht7 mutant, enlarging in volume with a progression of the day (ZT 6). By the start of the dark period (ZT 12), clear signs of division were evident by the increased number of dividing mother cells (Fig. 1A). By the end of the dark period (ZT 24), nearly all daughter cells were observed as hatched single cells outside of the mother cell wall (Fig. 1A).

Figure 1.
Figure 1.

N-replete photoautotrophic synchronized growth of the cht7 mutant under the 12 h light:12 h dark diel cycle. A, Microscopic images of wild-type 6145c, cht7, and CHT7-HA:cht7 cells at 0, 6, 12, and 24 h into the diel cycle. The light and dark periods are represented by white and black boxes, respectively. Scale bars = 10 μm. The cell size distributions (B) and the median cell sizes (C) of wild-type 6145c, cht7, and CHT7-HA:cht7 cultures at 0, 6, 12, and 24 h into the diel cycle. For C, numbers represent the averages and sds of two to four independent experiments.

These observations were also supported by the representative cell size distributions shown in Figure 1B, although a subpopulation of cht7 cells was observed to be larger than the wild-type or complemented cells at both ZT 0 and ZT 6, as indicated by the wider cell size distributions of cht7 cells. The median cell diameters of the cht7 mutant calculated from three independent experiments through the diel cycle were indistinguishable from those of the wild type or the complemented lines (Fig. 1C). Thus, the vast majority of cht7 cells grew and divided synchronously as observed for the wild-type and the complemented cells (Fig. 1). These results indicate that the loss of CHT7 does not have a major impact on cell cycle progression during N-replete photoautotrophic diel growth.

Levels of CHT7 Transcripts and CHT7 Proteins during the N-Replete Diel Cycle

The previous coexpression analysis of CHT7 using the Algae Gene Coexpression Database (Aoki et al., 2016) showed that CHT7 is highly coexpressed with genes involved in cell cycle regulation (Takeuchi et al., 2020). The coexpression of CHT7 with the cell-cycle-related genes was also observed by Zones et al. (2015). To assess whether the transcripts of CHT7 oscillate according to a distinct phase of the cell division cycle or the time of day, the expression levels of CHT7 were analyzed by reverse transcription quantitative PCR (RT-qPCR) in the wild-type and CHT7-HA:cht7 complemented lines during the light and dark synchronized growth (Fig. 2A). The CHT7 transcript abundance showed a strong diurnal oscillation, with its peak between 10 and 12 h after the start of a light period when many cells are entering the S/M phase of the cell division cycle and coincident with many cell cycle genes (Fig. 2A). When we examined the high-resolution transcriptomic data of synchronized wild-type C. reinhardtii cells (CC-5152, cw+, mt−; a progeny from a cross between 21gr and 6145c) during 12 h light:12 h dark cycle obtained by Zones et al. (2015), CHT7 and the C. reinhardtii RB pathway genes (MAT3, E2F1 and DP1) as well as other cell cycle genes showed a similar expression pattern (Supplemental Fig. S1). The abundance of CHT7 proteins also oscillated under the diel cycle when the levels of CHT7 were assessed in two wild-type strains (21gr and 6145c) and the CHT7-HA:cht7 complemented line by immunoblots using the CHT7 antibody (Fig. 2, B–D). The abundance data for CHT7 is consistent with a stable pool of protein being made during S/M phase when the CHT7 transcript level peaks, followed by dilution as cells grow, such that its concentration on a per-cell basis decreases during G1 phase. We note that if CHT7 remains chromatin-bound during G1, its effective concentration in cells and its function as a regulator of gene expression may not be affected by this dilution effect. Hence, the observed oscillation may not be a predictor of CHT7 function, rather the consequence of cell size changes and dilution effects occurring during the cell cycle.

Figure 2.
Figure 2.

Abundance of CHT7 transcripts and CHT7 proteins during N-replete light-dark synchronized growth. A, Levels of CHT7 transcripts assessed in wild-type 6145c and the CHT7-HA:cht7 complemented line cultured under 12 h light:12 h dark cycles. Target gene expression was normalized to the CBLP gene (Schloss, 1990). B to D, CHT7 protein levels and their quantification in wild types, 21gr (B), and 6145c (C), in addition to the CHT7-HA:cht7 complemented line (D) throughout the diel cycle. Equal amounts of total protein (25 μg) were separated by SDS-PAGE and followed by immunoblotting. The blots were probed with CHT7 antibody and later stained with Coomassie. For the quantification, values represent the averages and the sds of two to three biological replicates harvested at respective time points.

CHT7 and MAT3 Coimmunoprecipitate Together during N-Replete Synchronized Growth and following N Deprivation

The most abundant CHT7 complexes observed in dw15 (cw−) cells during mixotrophic N-replete growth and following N deprivation by BN-PAGE are similar in size (Tsai et al., 2014). When the BN-PAGE was performed using N-replete synchronized wild-type 21gr cells harvested during the 12 h light:12 h dark cycle and subjected to immunoblotting using CHT7 antibody, the majority of CHT7 was again observed in a protein complex of ∼242 kD (Fig. 3). Thus, we conclude that the Mr of the CHT7 complex observable by BN-PAGE remains constant regardless of the cell cycle stages or N availability.

Figure 3.
Figure 3.

Assessment of CHT7 complex size during N-replete light-dark synchronized growth. CHT7 complex visualized by BN-PAGE followed by immunoblotting in wild type 21gr at indicated times during 12 h light:12 h dark cycle. Equal amounts of total protein (25 μg) prepared under native nondenaturing conditions were subjected to BN-PAGE, and the blots were probed using CHT7 antibody.

Since CXC domain-containing proteins in other organisms are found in a conserved multisubunit protein complex with RB-related proteins (Korenjak et al., 2004; Lewis et al., 2004; Harrison et al., 2006; Litovchick et al., 2007; Pilkinton et al., 2007; Schmit et al., 2007; Tabuchi et al., 2011; Kobayashi et al., 2015), we tested whether CHT7 and MAT3/RB associate and whether their association is cycle or nutrient dependent, using coimmunoprecipitation (co-IP) assays. Nondenatured cell lysates for immunoprecipitation (IP) were prepared using mat3-4 cht7 double mutant cells with complementing fusion constructs producing HA-MAT3 and CHT7-GFP proteins during N-replete light-dark synchronized photoautotrophic growth (Fig. 4, A and B) and following 48 h of N deprivation under mixotrophic conditions (Fig. 4C), with the HA-MAT3:mat3-4 complemented line as a negative control. Although we did not include a line expressing GFP by itself to rule out direct interaction between the GFP tag and MAT3, we think this interaction is unlikely due to lack of protein interaction domains in GFP. We chose three cell cycle times for co-IP experiments: early-/mid-G1 phase (ZT 5) when cells were mostly precommitment stage, late-G1 phase (ZT 10) when cells were mostly postcommitment stage, and S/M phase (ZT13) when most cells were dividing (Fig. 4, A and B). Prior to IP, cells were treated with dithiobis-succinimidyl propionate to cross link proteins, and following sonication, the lysates were incubated with ethidium bromide on ice to eliminate chromatin association (Lai and Herr, 1992).

Figure 4.
Figure 4.

Co-IP of HA-MAT3 with CHT7-GFP during N-replete synchronized growth and following N deprivation. A, Microscopy images of the mat3-4 cht7 double mutant cells producing HA-MAT3 and CHT7-GFP proteins during 12 h light:12 h dark N-replete synchronized growth at indicated times. Scale bars = 10 μm. Co-IP of HA-MAT3 with CHT7-GFP assessed at indicated times during the N-replete diel cycle (B) or after 48 h of N deprivation (C). For both B and C, the IP was performed using the GFP antibody against the GFP-tag fused to CHT7. Twenty-five micrograms of total protein was loaded for input and flow through (FT). Ten percent volume of immunoprecipitated sample was used to check for the enrichment of CHT7-GFP in the IP fraction, and the rest was used to check whether HA-MAT3 coimmunoprecipitates with CHT7-GFP. In B, the blots to the left were probed with the CHT7 antibody, while those shown to the right were probed with the HA antibody. In C, the blots were probed with GFP antibody or HA antibody as indicated. A longer exposure of the immunoblot performed against the HA epitopes of MAT3 is shown to the right to clearly show the presence of HA-MAT3 in the IP fraction.

GFP-tagged CHT7 was pulled down with a GFP antibody, and different fractions were subjected to immunoblotting, then probed with the CHT7 antibody to check for the efficiency of IP (Fig. 4B, left blots) or with HA antibody to check for the co-IP of HA-MAT3 with CHT7-GFP (Fig. 4B, right blots). While HA-MAT3 coimmunoprecipitated with CHT7-GFP at all three cell cycle stages, the signals for HA-MAT3 pulled down with CHT7-GFP appeared the strongest at ZT13, in cells undergoing S/M phase (Fig. 4B). Furthermore, association between the two proteins was also detectable when the reverse co-IP experiments were performed by immunoprecipitating HA-MAT3 proteins with HA antibody and probing blots with CHT7 or GFP antibodies, though this experiment was more challenging due to higher background signal on the blots (Supplemental Fig. S2). In addition, when GFP-tagged CHT7 proteins were pulled down with GFP antibody using extracts prepared from N-deprived cells, HA-MAT3 was also observed in the IP fraction (Fig. 4C). However, the signals for coimmunoprecipated HA-MAT3 proteins were much weaker in comparison to those observed for N-replete synchronized conditions, which could be the result of slightly different experimental conditions in the independent experiments or of a changing strength of interaction between the different proteins under the different conditions. In summary, specific interaction between HA-MAT3 and CHT7-GFP was highly reproducible in our experiments, though we cannot completely rule out the unlikely possibility that the GFP or HA moieties happen to fortuitously interact with MAT3 or CHT7 proteins, respectively.

To test the impact of N deprivation on the levels of CHT7 or MAT3, their protein abundance was quantified in total protein extracts from two wild types, 21gr and 6145c, and the HA-MAT3 complemented line during N-replete growth or following 48 h of N deprivation by immunoblotting (Supplemental Fig. S3). Following a period of N starvation, the levels of both CHT7 and HA-MAT3 were reduced approximately by half on an equal protein basis compared to those observed during N-replete growth (Supplemental Fig. S3). Taken together, these results suggest the presence of complex(es) that contain both CHT7 and MAT3 in C. reinhardtii and their decreased abundance is at least partially responsible for the reduced co-IP signal for the two proteins following N deprivation.

The Majority of CHT7 and RB Proteins Exist in Different Complexes

Since the results of our co-IP assays showed that CHT7 and MAT3 coimmunoprecipitate together in vivo, we chose to test whether MAT3 is part of the stable main CHT7 complex observed by BN-PAGE. To this end, we first assessed the C. reinhardtii HA-MAT3 complex size using total protein extracts from the HA-MAT3:mat3-4 complemented cells during N-replete mixotrophic growth (Fig. 5A, left) and following 48 h of N deprivation (Fig. 5A, right) with BN-PAGE followed by immunoblotting with HA antibody. Regardless of the N conditions, HA-MAT3 proteins existed as part of a complex with a molecular mass of ∼480 kD, although the abundance of the HA-MAT3 complex decreased following N deprivation (Fig. 5A). Thus, presumably the most abundant form of the HA-MAT3 complex detected using BN-PAGE was substantially larger than that of the CHT7 complex (Fig. 5A), which has a molecular mass of ∼242 kD (Fig. 3; Tsai et al., 2014).

Figure 5.
Figure 5.

The bulk of CHT7 and MAT3 proteins exist in complexes of different sizes. A, HA-MAT3 complex sizes in wild-type 21gr and HA-MAT3:mat3-4 complemented line during N-replete mixotrophic growth (left) and following 48 h of N deprivation (right) as observed by BN-PAGE and subsequent immunoblotting with HA antibody. B, Comparison of CHT7 complex sizes in wild type 21gr and the mat3-4 mutant by BN-PAGE and immunoblotting with the CHT7 antibody. C, Antibody supershift assay followed by BN-PAGE and immunoblotting. The native protein extract from the HA-MAT3:mat3-4 complemented line was incubated with 10 μg of HA antibody, separated on a native gel, and subjected to immunoblotting. The blots were probed with HA antibody (left blot) or CHT7 antibody (right). In all cases, equal amounts of total nondenatured protein (25 μg) were used for gel electrophoresis. In B and C, protein extracts were prepared from cells cultured under N-replete mixotrophic conditions.

We then assessed the CHT7 complex size in the mat3-4 mutant and wild type 21gr with BN-PAGE followed by immunoblotting with the CHT7 antibody. Because the sizes of both CHT7 and HA-MAT3 complexes did not change in response to N availability, all the subsequent experiments were performed using cells grown under N-replete mixotrophic conditions for simplicity. The CHT7 complex detected in the mat3-4 mutant was similar in size to that of wild type, and no downward shift in their apparent molecular mass was observed for the mutant (Fig. 5B), indicating that MAT3 is not a part of the major CHT7 complex visualized by BN-PAGE. These findings were further corroborated by an antibody supershift assay followed by BN-PAGE, where the total protein extract from the HA-MAT3:mat3-4 line was incubated with HA antibody to allow for the formation of an antibody bound HA-MAT3 complex prior to native-PAGE and immunoblotting (Fig. 5C). As expected, the antibody coupled HA-MAT3 complex showed an increase in its apparent molecular mass as compared to the HA-MAT3 complex without the antibody coupling when the blot was probed with HA antibody (Fig. 5C, left). However, when the same blot was stripped and probed with the CHT7 antibody, the apparent size of the CHT7 complex in the HA antibody-coupled sample remained similar to that without the antibody coupling (Fig. 5C, right), further strengthening the conclusion that HA-MAT3 is indeed not a component of the major CHT7 complex that is visualized by BN-PAGE. These results indicate that the majority of CHT7 and HA-MAT3 proteins exist in complexes of different sizes during normal growth and following N deprivation.

Phosphorylation Status of CHT7 during Life Cycle Transitions

Previously, structure-function analyses of CHT7 showed that the predicted protein-interaction domains at its highly disordered C-terminal end are indispensable for CHT7 function (Takeuchi et al., 2020). Posttranslational modifications play a key role in the regulation of proteins with intrinsically disordered regions (Oldfield and Dunker, 2014). While a variety of posttranslational modifications may play a role, a C. reinhardtii phosphoproteomic study by Wang et al. (2014) indicated that CHT7 is phosphorylated at S84 and S90 during N-replete mixotrophic growth. Thus, the phosphorylation status of CHT7 was initially assessed in the CHT7-HA:cht7 complemented line during N-replete light-dark synchronized growth to determine whether CHT7 is phosphorylated in vivo and if this phosphorylation is regulated in a cell cycle stage-dependent manner (Fig. 6, A–C).

Figure 6.
Figure 6.

Phosphorylation status of CHT7-HA during N-replete synchronized growth and following N deprivation. A, Microscopic images of CHT7-HA:cht7 complemented cells during 12 h light:12 h dark N-replete synchronized growth at indicated times. Scale bars = 10 μm. B, Detection of phosphorylated CHT7-HA isoforms at indicated times during N-replete diel synchronized growth by phos-tag SDS-PAGE followed by immunoblotting with HA antibody. One sample under each condition was treated with calf intestinal phosphatase (CIP; indicated by the plus signs) to identify the nonphosphorylated isoform of CHT7-HA. Equal amounts of total protein (10 μg) were loaded. C, Quantification of phosphorylated CHT7-HA signals (per total CHT7-HA signals detected) at indicated times during the diel cycle by phos-tag acrylamide gels followed by immunoblotting with HA antibody. The values represent averages and sds of three independent experiments. D, Detection of phosphorylated CHT7-HA isoforms during N-replete mixotrophic growth and following 48 h of N deprivation by phos-tag SDS-PAGE followed by immunoblotting with HA antibody. Samples treated with CIP are indicated with plus signs as above.

We used phos-tag SDS-PAGE gels (Kinoshita et al., 2006; Kinoshita and Kinoshita-Kikuta, 2011) followed by immunoblotting using an HA antibody with extracts prepared from synchronized populations enriched in presumed pre- (ZT 4) or postcommitment (ZT9) cells, in addition to the S/M phase cells (ZT 12 and ZT 13) during the diel cycle (Fig. 6, A and B). In all cases, we compared migration of untreated samples to control samples that were pretreated with CIP to remove phosphoryl groups. The immunoblots probed by HA antibody revealed the presence of hypophosphorylated CHT7-HA and at least two phosphorylated isoforms of CHT7-HA that increase in abundance as the cell cycle progresses, which appeared to peak during the S/M phase when cells are undergoing division (ZT 12 and ZT 13; Fig. 6B). Results of the quantified percentages of phosphorylated CHT7-HA at indicated times from immunoblots across three independent experiments also supported this observation (Fig. 6C).

We then tested whether the phosphorylation status of CHT7 changes in response to N availability by performing the phos-tag experiments during N-replete mixotrophic growth and following 48 h of N deprivation (Fig. 6D). Although nonphosphorylated CHT7-HA as well as at least two phosphorylated isoforms of CHT7-HA were observed independently of N conditions, all nonmodified and modified forms of CHT7-HA were more abundant during N-replete growth, when cells have sufficient nutrients to support the cell division cycle, than following a period of N starvation (Fig. 6D). This is likely explained by the overall decrease in the levels of CHT7 proteins following N deprivation (Supplemental Fig. S3). These results indicated that CHT7 is indeed phosphorylated in vivo and that the proportion of overall phosphorylated CHT7 is higher in cells that are nearing or undergoing mitosis.

The Impact of CHT7 Phosphosilent and Phosphomimic S84 and S90 Mutations on Its Activity

The previous C. reinhardtii phosphoproteomics study by Wang et al. (2014) identified two phosphorylated Ser residues, S84 and S90, on the N-terminal end of CHT7 (Fig. 7A) during N-replete mixotrophic growth in tris-acetate phosphate (TAP) medium. To determine whether the sites of CHT7 phosphorylation potentially correspond to these residues and to better understand their contributions to the regulation of CHT7 function, CHT7-HA constructs containing phosphosilent (A84/A90) or phosphomimic (D84/D90) mutations of the two serines were introduced into the cht7 mutant, and the presence of the respective protein products was confirmed by immunoblotting with the HA antibody (Fig. 7B). To test whether the amino acid substitutions of S84 and S90 abolish the presence of the phosphorylated CHT7 isoforms, phos-tag SDS-PAGE followed by immunoblotting with HA antibody was performed using these phosphomutant lines (Fig. 7C). Nonphosphorylated and phosphorylated forms of the mutated CHT7-HA persisted in both CHT7-HA A84/A90 and D84/D90 lines, although the signals for the CHT7-HA proteins containing the respective mutations were always weaker than those observed for the wild-type CHT7-HA proteins (Fig. 7C). No obvious differences in abundance between the two CHT7-HA versions were detectable when the denatured proteins were separated by SDS-PAGE and subjected to immunoblotting using HA antibody (Fig. 7B).

Figure 7.
Figure 7.

Characterization of putative CHT7-HA phosphorylation sites using phospho-silent and phospho-mimic CHT7-HA mutants. A, Schematic representation of the full-length CHT7-HA protein sequence showing the putative CHT7 phosphorylation sites, S84 and S90, identified in the C. reinhardtii phosphoproteomic studies by Wang et al. (2014). The CXC domain (AA107-226) is also indicated by the red box outline. B, An immunoblot of CHT7-HA, phospho-silent CHT7-HA A84/A90, and phospho-mimic CHT7-HA D84/D90 proteins probed using HA antibody. Equal protein amounts were loaded and separated by SDS-PAGE. The blot was subsequently stained by Coomassie Brilliant Blue. C, Detection of phosphorylated CHT7-HA isoforms during N-replete mixotrophic growth in the CHT7-HA:cht7 complemented line and phospho-silent CHT7-HA A84/A90:cht7 and phospho-mimic CHT7-HA D84/D90:cht7 lines by phos-tag acrylamide gels followed by immunoblotting with HA antibody. Samples treated by CIP are indicated by the plus signs. Equal amounts of total protein were used. D, Growth curve of wild type 21gr, cht7, in addition to the full-length CHT7-HA and its clones with either phospho-silent (A84/A90) or phospho-mimic (D84/D90) mutations in the cht7 mutant background during N refeeding (NR) after 48 h of N deprivation. E, TAG content of the same set of lines after 24 h of N refeeding following 48 h of N deprivation. Three independent cultures were analyzed, and sd is shown.

To determine the contributions of potential S84 and S90 phosphorylations on CHT7 function, we assessed the ability of the mutated CHT7-HA to rescue the growth and TAG degradation delay observed in the cht7 mutant during N refeeding and found that these phenotypic defects were fully restored in cht7 cells producing CHT7-HA with phosphosilent (A84/S90) or phosphomimic (D84/D90) mutations (Fig. 7, D and E). These results suggest that while these two Ser residues of CHT7 may be phosphorylated, they do not significantly contribute to the physiological function of CHT7 under the conditions tested.

DISCUSSION

In our previous study, we showed that the presence of CHT7 was essential for the repression of a number of S/M phase-specific genes and for the proper coordination of cell division during N deprivation-induced quiescence in C. reinhardtii (Takeuchi et al., 2020). Here, we investigated the potential molecular mechanisms modulating CHT7 activity during N-replete light-dark synchronized photoautotrophic growth and following a period of N starvation, primarily at the protein level. We began by examining the role of CHT7 in the control of the cell division cycle by testing whether CHT7 is required for the synchronization of cells by the alternating light-dark cycles and observed no defects in the kinetics of cell cycle progression in cht7 cells during the N-replete diel cycle (Fig. 1). Therefore, given the previously observed phenotypic defects of the cht7 mutant during N starvation, including the loss of viability and the presence of multiple nuclei (Takeuchi et al., 2020), the physiological impact of the loss of CHT7 appears to be more apparent when N availability is limiting. In addition, although the CHT7 transcripts and CHT7 proteins both showed patterns of abundance resembling those for MAT3/RB complex proteins and other cell cycle genes/proteins, (Fig. 2; Supplemental Fig. S1), the sizes of the CHT7 complex visualized by BN-PAGE remained constant regardless of the cell cycle stage (Fig. 3).

To gain insights into mechanisms governing CHT7 complex activity during synchronized growth and following N deprivation, we initially tested whether CHT7 interacts with MAT3, a homolog of the RB protein in C. reinhardtii, in a cell cycle- or N-dependent manner by co-IP (Fig. 4). In Arabidopsis, a CXC domain protein, TCX5, and the RB ortholog, RBR1, represent the two core components present in both the repressor and activator forms of the DREAM-like complexes, although the transcriptional regulatory activities of these complexes have not been attributed directly to TCX5 in plants (Fischer and DeCaprio, 2015; Kobayashi et al., 2015; Fischer and Müller, 2017). Our results showed that CHT7 and MAT3 coimmunoprecipitate regardless of the cell cycle stage or N availability (Fig. 4). However, during the N-replete synchronized growth, the interaction between the two proteins appeared to be most prevalent in mitotic cells, potentially due to the increasing abundance of these proteins around this stage (Fig. 4).

We then tested whether MAT3 is part of the CHT7 complex visualized by BN-PAGE followed by immunoblotting and found that MAT3 is not a constituent of the bulk CHT7 complex (Fig. 5). These results at first glance appear to contradict the results of the co-IP experiments. However, in the BN-PAGE experiments, only microgram amounts of nondenatured total cell lysates could be loaded, and the protocol is performed without the prior enrichment of CHT7 or MAT3/RB proteins and without the use of cross linkers. Therefore, it is possible that only the most abundant, robust, or stable CHT7 or MAT3 core complexes are detectable by BN-PAGE. Using the more sensitive co-IP approach, a lower abundance or less stable CHT7 and MAT3 interaction that could be direct or indirect was detected. Because the native lysate was treated with ethidium bromide during the co-IP procedures, it is unlikely that the observed association is mediated through chromatin interactions. Taken together, these results point to the presence of a low abundance or unstable subcomplex that contains both CHT7 and MAT3/RB. Finally, it is worth noting that mat3 and dp1 mutants showed altered cell cycle progression but no transcriptional defects in cell cycle genes (Fang et al., 2006; Cross, 2020), while cht7 mutants showed derepression of cell cycle genes (Takeuchi et al., 2020), indicating that the influence on the cell cycle of the most abundant complexes containing either CHT7 or MAT3/RB, respectively, may be through different mechanisms. The nature of the interaction between MAT3/RB and CHT7 components and the significance of this interaction remain to be determined.

Through an attempt to better understand the role of phosphorylation in mediating CHT7 activity, we identified at least two phosphorylated species of CHT7 through phos-tag SDS-PAGE followed by immunoblotting (Fig. 6), suggesting CHT7 as a potential target of a yet-unspecified protein kinase-based signaling pathway. While we observed phosphorylated CHT7 proteins both during N-replete light-dark synchronized growth and following N deprivation (Fig. 6, B and D), the sites, numbers, and combinations of such phosphorylations as well as their potential functional significance remain unknown. Indeed, the mutations of S84 and S90, identified as the putative phosphorylation sites of CHT7 by Wang et al. (2014), neither fully eliminated the phosphorylated CHT7 isoforms observed by phos-tag SDS-PAGE followed by immunoblotting nor compromised the function of CHT7 (Fig. 7).

Thus, the identification of CHT7 complex components and the localization of CHT7 phosphorylation during the cell division cycle and N deprivation-induced quiescence are now critical next steps in gaining deeper mechanistic insights into the regulation of CHT7 activities. This is especially true given the previous findings that the highly disordered C-terminal end of CHT7 with predicted protein-interaction domains, and not its CXC domain, is essential for the function of CHT7 (Takeuchi et al., 2020). Furthermore, since the putative phosphorylation sites of CHT7 identified to date are located on the dispensable N terminus of the protein, whether any posttranslational modifications exist on the largely unstructured C terminus of CHT7 and if they are relevant to its function remain to be explored. Various mutated versions of CHT7 should serve as useful tools in future mass spectrometry endeavors to determine CHT7 complex members and the sites of CHT7 phosphorylation, in addition to gaining further mechanistic insights into CHT7 function during N-replete synchronized growth and especially N deprivation-induced quiescence.

MATERIALS AND METHODS

Generation and Validation of Lines

The cell-wall-less Chlamydomonas reinhardtii parental line, dw15 (cw15, nit1, cw−, mt+) was obtained from Arthur Grossman, Department of Plant Biology, Carnegie Institution for Science, Stanford, California. The cell-walled wild-type strains 21gr and 6145c (cw+, mt+, and mt−, respectively), in addition to the HA-MAT3:mat3-4 complemented line (cw+, mt+; Olson et al., 2010) were obtained from James Umen, Donald Danforth Plant Center, Olivette, Missouri. Generation of the original cell-wall-less cht7 mutant in the dw15 background (cw−, mt+; Tsai et al., 2014) and the ninth generation backcrossed cell-walled cht7 (cw+, mt−; Takeuchi et al., 2020) were described previously. Generation of the C terminally GFP-tagged CHT7 complementation constructs, pMN24 CHT7-GFP, and its transformation into the cell wall-deficient cht7 mutant were described previously (Tsai et al., 2014).

To produce the HA-MAT3 CHT7-GFP:mat3-4 cht7 (cw+, mt+) line used in this study, pMN24 CHT7-GFP:cht7 (cw−, mt+) was first crossed to cw+ CC198 (er-u-37, str-u-2-60, cw+, mt−), and a cw+ cht7 progeny producing CHT7-GFP was further crossed to the HA-MAT3:mat3-4 complemented line. The resulting meiotic progenies were initially screened based on their resistance to hygromycin B (aphv7, linked to the cht7 mutation) and paromomycin (aphv8, linked to the pSL18 HA-MAT3 complementation construct), in addition to their ability to grow using nitrate (NIT1, linked to the pMN24 CHT7-GFP complementation construct). Since the mat3-4 mutation is known to be linked to the mt+ locus (Armbrust et al., 1995), mt+ lines were selected by mating type PCR using the published primers (Table 1; Zamora et al., 2004). Primers published in Tsai et al. (2014), APH7-F and APH7-R, in addition to CHT7-F and CHT7-R, were used for genotyping PCR to validate the presence of the aphv7 insert and the CHT7 gene, respectively (Table 1). The presence of CHT7-GFP and HA-MAT3 proteins was verified by SDS-PAGE followed by immunoblotting using a 1:1000 dilution of GFP antibody (Abcam, ab6673) and 1:10,000 donkey α-goat-HRP secondary antibody (Santa Cruz Biotechnolgy, sc2020), or using a 1:1000 dilution of HA-HRP antibody (Sigma, catalog #12013819001), respectively. CC-198 used for the cross described above, and the mat3-4 mutant (cw+, mt+, CC3994) were obtained from the Chlamydomonas Resource Center (http://www.chlamycollection.org).

View this table:
Table 1. Primers used for site-directed mutagenesis (SDM), genotyping PCR, and RT-qPCR

The generation of pSL18 (−AscI) CHT7-3xHA and pGEM CHT7 constructs, which served as final destination vector and template for CHT7 phosphomimic (D84/D90) and -silent (A84/A90) SDM PCRs, respectively, was described previously (Takeuchi et al., 2020). SDM PCRs were carried out using pGEM CHT7 as a template, with SDM primer pairs provided in Table 1. The presence of the desired mutations and the correctness of the PCR-amplified sequences were confirmed by sequencing. To make pSL18 (−AscI) CHT7-3xHA D84/D90 and A84/A90 constructs, pGEM CHT7 constructs containing the respective mutations were digested with AscI and BsrGI-HF and ligated into pSL18 (−AscI) CHT7-3xHA digested with the same enzymes. Generation and validation of the ninth-generation cw+ cht7 mutant transformed with the pSL18 (−AscI) CHT7-3xHA complemented line were described previously (Takeuchi et al., 2020). Introduction of the mutated pSL18 (−AscI) CHT7-3xHA D84/D90 and A84/A90 constructs linearized with NcoI and AhdI, respectively, into the cw+ cht7 mutant were performed according to Takeuchi et al. (2020). The production the CHT7-3xHA proteins (with or without the respective mutations) in cht7 was verified by SDS-PAGE followed by immunoblotting, using a 1:1000 dilution of HA-horseradish peroxidase (HRP) antibody (Sigma, catalog no.12013819001).

Growth Conditions

For synchronization, the cells from respective lines were initially inoculated into HS medium (Sueoka, 1960) in 125-mL Erlenmeyer flasks, and the cultures were bubbled with an equal volume of mixed gas (95% air and 5% [v/v/] CO2) every hour, with constant stirring using a magnetic bar at 200 rpm. The cultures were kept under a 12 h light:12 h dark cycle and 250 μmol m−2s−1 of light (5600K; YUJILEDS VTC series high CRI LED 2835) at ambient room temperature (∼24°C) until reaching log phase growth. The ePBRs (Phenometrics, PBR102-F model) were inoculated to a final cell density of approximately 1 × 105 cells mL−1 in 330 to 350 mL of HS media, and the cultures were grown under 12 h light:12 h dark cycles with 2000 μmol m−2s−1 of light (FSL200 full spectrum light; https://www.phenometricsinc.com). The reactors were kept at a constant temperature of 28°C with stirring at 200 rpm. The ePBR cultures were injected with air mixed with 5% (v/v) CO2 for 30 s every 15 min at a flow rate of 0.36 L min−1. Each ePBR was normalized to approximately 3 μg mL−1 of total chlorophyll by measuring the chlorophyll levels at ZT 6 and setting the automated dilution using a turbidostat. The cultures were maintained under a steady-state growth at midlog concentration for at least 2 d prior to the start of the experiment. More information on the ePBR designs and parts can be found in Lucker et al. (2014) and the manufacturer’s website (https://www.phenometricsinc.com).

To obtain N-deprived cultures, cells were grown mixotrophically in Tris acetate phosphate medium (Harris, 1989) to a midlog phase under 100 to 120 mmol m−2s−1 of constant light (4200K; SYLVANIA F24T12 CW/HO - 35W) at 22°C and were N deprived as previously described (Takeuchi et al., 2020). For growth curves during N refeeding, N was resupplied from a 100× NH4Cl solution after 48 h of N deprivation, and the growth was assessed by measuring the OD750 of respective cultures at indicated times.

Assessment of Total Chlorophyll Content

For normalization of each ePBR culture based on chlorophyll content, cells collected from 1 mL of culture at ZT 6 were resuspended in 1 mL of 80% (v/v) acetone, centrifuged at 15,000g for 5 min at room temperature, and the absorbance of supernatant was assessed at 646 and 663 nm. Total chlorophyll per mL was calculated according to Porra (2002).

Microscopy and Cell Size Analysis

At indicated times during the 12 h light:12 h dark cycle, cells were fixed in 0.2% glutaraldehyde in phosphate-buffered saline (PBS) for 1 h at room temperature or overnight at 4°C. The fixed cells were centrifuged at 3000g for 5 min at room temperature, washed once, and resuspended in PBS. All microscopy was performed using the Leica DMi 800 inverted microscope (Leica Microsystems) and a 63× oil immersion objective (HC PL APO 63×/1.40 oil CS2), and the images were collected using Leica Application Software (https://www.leica-microsystems.com). The cell size distributions were analyzed by a Beckman Coulter Counter Z2 and AccuComp Z2 software (https://www.beckman.com).

RT-qPCR

RNA extraction and RT-qPCR analyses were performed as previously described (Takeuchi et al., 2020). Sequences of the RT-qPCR primers used are provided in Table 1.

Immunoblotting

Preparation of the total denatured cell lysates, SDS-PAGE and immunoblotting using CHT7 antibody were performed as described previously (Tsai et al., 2014). The fused epitopes (HA and GFP tags) were probed by a 1:1000 dilution of HA-HRP antibody (Sigma, catalog no. 12013819001) or 1:1000 dilution of GFP antibody (Abcam, ab6673) followed by 1:10,000 donkey α-goat-HRP secondary antibody (Santa Cruz Biotechnolgy, sc2020).

BN-PAGE

BN-PAGE was performed as described in Tsai et al. (2014). For antibody supershift assay followed by BN-PAGE, approximately 100 μg of total nondenatured protein was incubated with 10 μg of HA antibody (BioLegend, 16B12, catalog no. 901503) for 1 h on ice, and equal protein amounts (25 μg, prior to the antibody addition) were loaded onto native gels prior to carrying out the rest of BN-PAGE protocol.

Co-IP Assays

For the preparation of input proteins, 200 to 250 mL of cultures grown under the respective conditions were centrifuged at 3000g for 5 min at 4°C in the presence of a 1:4000 dilution of 20% (v/v) Tween 20 and resuspended in 1 mL of resuspension buffer (PBS, 1:100 plant protease inhibitor cocktail [Sigma-Aldrich], 1:100 phosphatase inhibitor cocktail [Halt; Thermo Scientific], and 1:100 phenylmethylsulfonyl fluoride [PMSF; Sigma-Aldrich, 93482]). Proteins were cross linked by the addition of freshly prepared dithiobis-succinimidyl propionate cross linker to cells (Thermo Scientific) to a final concentration of 1 mm and incubation on ice for 30 min. The cross linking reaction was quenched by the addition of 1 m Tris-HCl, pH 7.5, to a final concentration of approximately 100 mm and incubation on ice for 15 min. When the lysate preparation was not performed on the same day, cells were centrifuged at 3000g for 5 min at 4°C, resuspended in 1 mL of storage buffer (resuspension buffer containing 20% [v/v] of glycerol), and kept at −80°C upon freezing in liquid N. Prior to sonication, samples were thawed on ice, and the storage buffer was again replaced by 1 mL of resuspension buffer before proceeding. The sonication was performed using a Misonix Inc S-3000 sonicator (Cole-Parmer) with a one-sixteeth-inch microtip probe at 4°C. Samples were kept on watery ice, and the sonication program (total process time of 2 min with 0.5 s on and off pulses at the power output setting of 2.0) was repeated six times with inversion of tubes after the end of each cycle. To eliminate DNA-dependent interactions, the lysate was incubated with 50 μg mL−1 (final concentration) of ethidium bromide on ice for 30 min, and the supernatant was obtained by centrifuging for 30 min at 20,000g and 4°C. The protein concentration was assessed using Bradford reagent (Bio-Rad), and 3 to 5 mg of total protein extracts adjusted with resuspension buffer to a final volume of 1 mL were used for IP after preclearing the lysate with 20 μL of prewashed uncoupled protein G dynabeads (Thermo Scientific) for 45 min at 4°C with rotation. For the pull-down, protein G dynabeads were coupled with either HA (BioLegend, 16B12, catalog no. 901503) or GFP antibody (Abcam, catalog no. ab1791), and the IP was performed by rotating 50 μL of antibody-coupled beads and the aforementioned precleared lysate at 4°C overnight (∼14 h). The antigen-bound beads were washed five times with 1 mL of cold washing buffer 1 (PBS, additional 150 mm of NaCl, 0.05% [v/v] Tween 20 and 1:100 PMSF [Sigma-Aldrich, 93482]) and once with 1 mL of washing buffer 2 (PBS, 1:00 PMSF [Sigma-Aldrich, 93482]). The bound-proteins were eluted from beads in 50 μL of 2× Laemmli sample buffer (4% [w/v] SDS, 20% [v/v] glycerol, 125 mm Tris-HCl, pH 6.8, 0.004% [v/v] bromophenol blue) with β-mercaptoethanol (5%; v/v) and dithiothreitol (at a final concentration of 100 mm) added immediately before use and by incubating the samples for 10 min at 95°C prior to SDS-PAGE and immunoblotting.

Phos-tag SDS-PAGE

For sample preparation, 10 to 15 mL of cultures grown under the respective conditions were centrifuged at 3000g for 5 min at 4°C in the presence of 1:4000 dilution of 20% (v/v) Tween-20, and the cells were resuspended to a final volume of 250 μL with 1× NEB buffer 3 (New England Biolabs, NEBuffer 3) containing 1:100 dilutions of plant protease inhibitor cocktail (Sigma-Aldrich) and PMSF (Sigma-Aldrich, 93482). For the CIP treatment, cells were sonicated using a Misonix S-3000 sonicator (Cole-Parmer) with a one-sixteeth inch microtip probe at 4°C as follows. Samples were kept on watery ice and were sonicated for a total process time of 2 min, with 10 s on and off pulses at the power output setting of 1.0. CIP (New England Biolabs) was added to a final concentration of 0.8 U μL−1 of sample, and the tubes were incubated on ice for 15 min and at 37°C for 15 min. Reactions were stopped by the addition of an equal volume (250 μL) of 2× Laemmli sample buffer (4% [w/v] SDS, 20% [v/v] glycerol, 125 mm Tris-HCl, pH 6.8, 0.004% [v/v] bromophenol blue, 10% [v/v] β-mercaptoethanol, 200 mm dithiothreitol) and by incubating the samples for 5 min at 95°C. For non-CIP treatment samples, the cells were resuspended to a final volume of 250 μL in the 1× NEB buffer 3 mixture described above followed by an equal volume of 2× Laemmli sample buffer and then heated for 5 min at 95°C. The protein concentration was assessed using RC DC protein assay (Bio-Rad), and equal amounts of proteins were separated in Bis-Tris (pH 6.8, final concentration of 357 mm) acrylamide gels containing 75 μm of phos-tag (NARD Institute, LTD, AAL-107) and 150 μm of zinc nitrate. Phos-tag SDS-PAGE was performed as described in Kinoshita and Kinoshita-Kikuta (2011) and followed by immunoblotting using 1:1000 dilution of the HA-HRP antibody (Sigma, catalog no. 12013819001). Signal quantifications of the blots were performed by Bio-Rad Image Lab software (https://www.bio-rad.com).

Lipid Analysis

The quantification of TAGs was performed as previously described in Takeuchi et al. (2020).

Accession Numbers

The following accession numbers are found in the JGI-Phytozome 13 C. reinhardtii v5.5 database: MAT3/RB (Cre06.g255450); CHT7 (Cre11.g481800); E2F1 (Cre01.g052300); and DP1 (Cre07.g323000).

Supplemental Data

The following supplemental materials are available.

  • Supplemental Figure S1. High-resolution expression profiles of CHT7 and RB pathway genes (MAT3, E2F1, and DP1) of wild type cultured under 12 h light:12 h dark cycles obtained from the RNA-seq datasets generated by Zones et al. (2015).

  • Supplemental Figure S2. Co-IP of CHT7-GFP with HA-MAT3 during N-replete light-dark synchronized growth and under N-replete mixotrophic growth.

  • Supplemental Figure S3. Abundance of CHT7 and HA-MAT3 proteins during N-replete growth and following N deprivation.

Acknowledgments

The contents of this study are solely the responsibility of the authors and do not necessarily represent the official views of the National Institute of General Medical Sciences or National Institutes of Health.

Footnotes

  • The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Christoph Benning (benning{at}msu.edu).

  • T.T. designed and conducted the protein experiments and microscopy and analyzed the data; Y.-T.L. performed the synchronization of cht7 mutant cultures and RT-qPCR analysis; T.T. and Y.-T.L. contributed to harvesting of samples; N.F. conducted cell size measurements and assisted in the maintenance of photobioreactors; J.U. designed experiments and interpreted data; B.B.S. designed experiments, interpreted data, and performed the crosses for strain construction; C.B. designed and coordinated the study and interpreted data; and T.T. provided the draft of the manuscript, which was edited by B.B.S. and C.B. with feedback provided by all other authors.

  • www.plantphysiol.org/cgi/doi/10.1104/pp.20.00864

  • ↵1 This work was supported by the National Science Foundation (grant nos. MCB–1515169 to C.B., and MCB–1515220 and IOS 1755430 to J.U.), the Chemical Sciences, Geosciences, and Biosciences Division, Office of Basic Energy Sciences, Office of Science, U.S. Department of Energy (grant no. DE–FG02–91ER20021 to C.B.), the Michigan State University AgBioResearch (grant no. MICL02357 to C.B.), the National Institutes of Health (grant no. R01GM126557 to J.U.), the U.S. Department of Energy, Office of Science, Office of Biological and Environmental Research, Genomic Science Program (grant no. DE–SC0020400 to J.U.), and a predoctoral training award from the National Institute of General Medical Sciences of the National Institutes of Health (grant no. T32–GM110523 to T.T.).

  • ↵3 Senior author.

  • ↵[OPEN] Articles can be viewed without a subscription.

  • Received July 6, 2020.
  • Accepted September 21, 2020.
  • Published October 1, 2020.

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Modulation of CHT7 Complexes during Light/Dark- and Nitrogen-Mediated Life Cycle Transitions of Chlamydomonas
Tomomi Takeuchi, Yang-Tsung Lin, Nicholas Fekaris, James Umen, Barbara B. Sears, Christoph Benning
Plant Physiology Dec 2020, 184 (4) 1762-1774; DOI: 10.1104/pp.20.00864
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