Defects in a New Class of Sulfate/Anion Transporter Link ...defects in sulfur metabolism do not...

Defects in a New Class of Sulfate/Anion Transporter LinkSulfur Acclimation Responses to Intracellular GlutathioneLevels and Cell Cycle Control1[W][OPEN]

Su-Chiung Fang*, Chin-Lin Chung, Chun-Han Chen, Cristina Lopez-Paz, and James G. Umen*

Biotechnology Center in Southern Taiwan, Academia Sinica, Tainan County 741, Taiwan (S.-C.F., C.-L.C.,C.-H.C.); Agricultural Biotechnology Research Center, Academia Sinica, Taipei 115, Taiwan (S.-C.F., C.-L.C.,C.-H.C.); Institute of Marine Biotechnology, National Sun Yat-sen University, Kaohsiung 80424, Taiwan(C.-L.C.); and Donald Danforth Plant Science Center, St. Louis, Missouri 63132 (C.L.-P., J.G.U.)

We previously identified a mutation, suppressor of mating type locus3 15-1 (smt15-1), that partially suppresses the cell cycle defectscaused by loss of the retinoblastoma tumor suppressor-related protein encoded by the MAT3 gene in Chlamydomonas reinhardtii.smt15-1 single mutants were also found to have a cell cycle defect leading to a small-cell phenotype. SMT15 belongs to apreviously uncharacterized subfamily of putative membrane-localized sulfate/anion transporters that contain a sulfatetransporter domain and are found in a widely distributed subset of eukaryotes and bacteria. Although we observed that smt15-1has a defect in acclimation to sulfur-limited growth conditions, sulfur acclimation (sac) mutants, which are more severely defectivefor acclimation to sulfur limitation, do not have cell cycle defects and cannot suppress mat3. Moreover, we found that smt15-1,but not sac mutants, overaccumulates glutathione. In wild-type cells, glutathione fluctuated during the cell cycle, with highestlevels in mid G1 phase and lower levels during S and M phases, while in smt15-1, glutathione levels remained elevated during S andM. In addition to increased total glutathione levels, smt15-1 cells had an increased reduced-to-oxidized glutathione redox ratiothroughout the cell cycle. These data suggest a role for SMT15 in maintaining glutathione homeostasis that impacts the cell cycleand sulfur acclimation responses.

Cell cycle progression is coordinated with the cellularredox environment, which also undergoes periodic cycling.In budding yeast (Saccharomyces cerevisiae), DNA synthesisand mitosis occur during the reductive phase and celldivision initiates in the oxidative phase (Tu et al., 2005).Restriction of DNA replication to the reductive phase ofthe metabolic cycle is important to ensure genome in-tegrity (Chen et al., 2007). In mammals, low levels of re-active oxygen species stimulate cell cycle entry (Lee et al.,1998; Martindale and Holbrook, 2002; Boonstra and Post,2004) by activating cell cycle regulators (Shackelford et al.,

2000; Boonstra and Post, 2004; Macleod, 2008; Burhansand Heintz, 2009). Moreover, alterations in redox homeo-stasis can cause defects in cell cycle progression (Espositoet al., 1997; Reichheld et al., 1999; Alic et al., 2001; Menonet al., 2003; Markovic et al., 2009; Tsukagoshi et al., 2010).Glutathione is a thiol-containing tripeptide whose functionis not only important to maintain redox homeostasis whencoping with biotic and abiotic stresses (Cobbett et al., 1998;Ball et al., 2004; Rouhier et al., 2008; Foyer and Noctor,2009; Mhamdi et al., 2010; Dubreuil-Maurizi and Poinssot,2012; Shanmugam et al., 2012) but also acts as a redoxsignal or sensor for cell cycle control (Chiu et al., 2011;Chiu and Dawes, 2012). Therefore, defects in glutathione-mediated redox balance can lead to aberrant cell cycleprogression and, subsequently, defects in growth anddevelopment (Vernoux et al., 2000; Cairns et al., 2006; Jiaoet al., 2013). However, the molecular mechanism thatconnects glutathione-mediated cellular redox state andcell cycle regulation is not fully understood.

Retinoblastoma-related proteins (RBRs) are evolution-arily conserved cell cycle regulators with a central rolein controlling the initiation of DNA replication and cellcycle entry. The canonical retinoblastoma (RB) pathwayinvolves the cell cycle-regulated interaction of RBRs witha heterodimeric E2 promoter binding factor (E2F)/Dimerization partner (DP) transcription factor. TheRB-associated E2F/DP protein complex represses thetranscription of cell cycle genes, and this repression isreleased by the removal or modification of RBRs viaphosphorylation. Subsequently, E2F/DP-dependent

1 This work was supported by the American Cancer Society (Re-search Scholar grant no. RSG–05–196–01–CCG to J.G.U.), the NationalInstitutes of Health (grant no. R01 GM 078376 to J.G.U.), the NationalScience Council (grant nos. 99–2311–B–001–001, 100–2311–B–001–009,and 101–2311–B–001–030 to S.-C.F.), and the Biotechnology Center inSouthern Taiwan, Academia Sinica (to S.-C.F.).

* Address correspondence to [email protected] [email protected].

S.-C.F. and J.G.U. conceived and designed the experiments; S.-C.F.,C.-L.C., and C.-H.C. performed the experiments and analyzed thedata; C.L.-P. contributed to reagents and materials; S.-C.F. andJ.G.U. interpreted the data and wrote the article.

The author responsible for distribution of materials integral to thefindings presented in this article in accordance with the policy describedin the Instructions for Authors (www.plantphysiol.org) is: Su-ChiungFang ([email protected]).

[W] The online version of this article contains Web-only data.[OPEN] Articles can be viewed online without a subscription.www.plantphysiol.org/cgi/doi/10.1104/pp.114.251009

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transcription of cell cycle genes promotes S phase entryand cell cycle progression. Because of its central rolein controlling the transcription of cell cycle genes, RBserves as a convergence point for regulating the cell cyclein response to internal and external mitogenic signals(Nakagami et al., 2002; Stevaux et al., 2002; Cobrinik,2005; Dimova and Dyson, 2005; Wikenheiser-Brokamp,2006; Jullien et al., 2008; van den Heuvel and Dyson,2008; Borghi et al., 2010; Henriques et al., 2010; Johnstonet al., 2010; Chen et al., 2011; Gutzat et al., 2011, 2012;Weimer et al., 2012). Recent studies also provide evidencethat RB depletion causes metabolic reprogramming andsuggest that the RB pathway in animals exerts part of itseffect on cell proliferation through the control of Glnmetabolism (Nicolay et al., 2013; Reynolds et al., 2014).Chlamydomonas reinhardtii is a unicellular green alga

that proliferates using a multiple fission cell cycle. Itsmitotic cell cycle starts with a long G1 phase, duringwhich cells can grow manyfold in size. At the end ofG1, mother cells undergo n rapid rounds of alternatingS phase (DNA synthesis) and M phase (mitosis) toproduce 2n daughter cells. Two size checkpoints areintegrated into this mitotic cell cycle. In early/mid G1,cells pass commitment, the first size checkpoint, whichrequires cells to acquire sufficient mass to be able tocomplete the cell cycle. The second size checkpoint oc-curs during S/M, where mother cells, whose sizes canbe highly variable, undergo an appropriate number ofdivision cycles to produce uniformly sized daughters(Craigie and Cavaliersmith, 1982; Donnan and John,1983). Because C. reinhardtii mother cell division canoccur in the absence of concurrent growth, daughter cellsize can be conveniently used to assess the cell sizecheckpoint function during S/M phase (Umen, 2005).Our previous studies showed that the RB pathway

in C. reinhardtii is important for size checkpoint controland size-mediated cell division (Umen and Goodenough,2001; Fang et al., 2006). The C. reinhardtii RBR ho-molog is encoded by a single gene, MAT3. mat3 mu-tants pass commitment at a smaller size than normalbut remain in G1 for a normal period of time beforeentering S/M, where they undergo supernumerarycell divisions to produce tiny daughter cells. Mutationsin the C. reinhardtii E2F1 and DP1 genes could suppressthe mat3 size defect, indicating that the overall archi-tecture of the RBR pathway is conserved in C. reinhardtii,with MAT3/RBR serving as a negative regulator ofE2F- and DP-related proteins (Fang et al., 2006; Olsonet al., 2010).Besides mutations in E2F1 andDP1, several additional

extragenic suppressors of mat3 (smt mutants) were iso-lated that are weaker suppressors than e2f1 and dp1(Fang and Umen, 2008). smt15-1 mat3-4 double mutantsare larger than mat3-4 mutants but smaller than wild-type cells. This finding suggested that SMT15 might be apositive regulator of cell division. However, smt15-1 singlemutants had reduced daughter cell size compared withwild-type cells, indicating a potential negative regulatoryrole for SMT15 in controlling size-dependent cell division.These results suggest a nonlinear relationship between the

MAT3/RB pathway and the pathway(s) that are im-pacted by SMT15.

In this study, we identified the SMT15 gene andshowed that it encodes a member of a conserved butuncharacterized family of proteins with homology tosulfate/anion transporters. Although smt15 strains showeddefects in acclimation to sulfur limitation, canonical sulfuracclimation mutants did not show cell cycle defects andwere unable to suppress mat3, indicating that generaldefects in sulfur metabolism do not impact the cell cycle.Instead, we found that glutathione, an end product ofsulfur assimilation, overaccumulated in the smt15-1 mu-tant, which also showed attenuated induction of sulfuracclimation genes. These results identify a link betweenglutathione-mediated redox regulation and the cell cyclein C. reinhardtii and suggest a potentially newmechanismfor glutathione homoeostasis mediated through a con-served family of membrane transporters.

RESULTS

Characterization of the SMT15 Locus

smt15-1was isolated as a recessive suppressor ofmat3-4in a genetic screen using a paromomycin resistancemarker as an insertional mutagen. Linkage between theparomomycin marker and suppression of the mat3-4size phenotype was established previously (Fang andUmen, 2008). In this study, the flanking sequence ad-jacent to the inserted paromomycin resistance marker insmt15-1 was identified, and the insertion was located ina gene encoding a putative transporter (Fig. 1A). Becausethe genome assembly of sequences surrounding theSMT15 locus was incomplete, we used reverse tran-scription (RT)-PCR and RACE-PCR to isolate and deducethe structure of the SMT15 mRNA and the predictedSMT15 protein (see “Materials and Methods”). RT-PCRusing primers flanking the insertion site amplified aproduct of the predicted size for SMT15 complemen-tary DNA (cDNA) prepared from wild-type RNA butnot from smt15-1 (Fig. 1B).

The predicted SMT15 protein is homologous to afamily of sulfate/anion transporters comprising a sul-fate transporter domain (Pfam 00916), a sulfate trans-porter and anti-sigma factor antagonist (STAS) domain(Pfam 01740), and a cyclic nucleotide-binding domain(Pfam 00027; Fig. 1C). Like other transporters in thissuperfamily, SMT15 is predicted to encode an integralmembrane protein with 10 transmembrane helices(Supplemental Fig. S1; Sonnhammer et al., 1998). Aphylogenetic tree constructed from previously iden-tified sulfate transporters and representatives identi-fied in BLAST searches revealed that SMT15 belongsto members of tribe 1 of eukaryotic sulfate/aniontransporters, which includes plant Sulfate transporter(SULTR), metazoan Solute Carrier26 transporter (SLC26),and fungal Sulfate Permease (SUL) families (Takahashiet al., 2012). The sequence alignment of the sulfatetransporter domain (Pfam 00916) of tribe 1 of eukaryoticsulfate/anion transporters is shown in Supplemental

Plant Physiol. Vol. 166, 2014 1853

SMT15 Governs Sulfur Stress, Glutathione, and the Cell Cycle


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Figure S2. However, SMT15 and its orthologs are dis-tantly related to the major families of tribe 1 transportersand, therefore, are classified as a new group, family C(Fig. 1D). Members of family Cwere found in diverse taxa,including chlorophycean green algae such as Volvox,Chlorella, and Coccomyxa spp., but undetectable in the more

basal prasinophyte algae or in land plants (SupplementalFig. S3). However, family C homologs were foundoutside the green lineage in opisthokonts, includingchoanoflagellates and fungi (but not metazoans), andstramenopiles, including diatoms, brown algae, andoomycetes, as well as in dinoflagellates (Fig. 1D;

Figure 1. Molecular characterizationof SMT15. A, Schematic of smt15-1 in-sertion with chromosome number, coor-dinates, and genome version indicated.White boxes indicate exons. The in-verted triangle indicates the positionwhere insertion occurred. The loca-tions of RT-PCR primers used to assessthe expression of SMT15 in complementedstrains are depicted by the arrowheads.B, RT-PCR of SMT15 or internal controlmessage G protein Beta Subunit-LikeProtein (GBLP ) in the indicated strains.The primer locations used to amplifySMT15 are shown in A. wt, Wild type.C, Schematic representation of the pre-dicted SMT15 protein. Sulfate_transp,Sulfate transporter domain (Pfam 00916);STAS (Pfam 01740); CAP_ED, cyclicnucleotide-binding domain of theCatabolite Activator Protein family ofprokaryotic transcription factors (Pfam00027). D, Phylogenic relationship ofSMT15, SMT15 othologs, and othertribe 1 sulfate/anion transporters (Takahashiet al., 2012). The major lineages areboxed in orange for family C, in greenfor family P, in purple for family A1,and in blue for family A2. Bootstrapvalues of 50% or higher are shown foreach clade. Abbreviations of genus andspecies names of the given locus numbersor gene identifiers are as follows:An, Aspergillus niger; At, Arabidopsis;Cre, C. reinhardtii; Cs, Coccomyxasubellipsoidea; Ds, Dactylococcopsissalina; Es, Ectocarpus siliculosus; Hs,Homo sapiens; Mb, Monosiga brevicollis;Pi,Phytophthora infestans; Pt,Phaeodactylumtricornutum; Sc, S. cerevisiae; Ss, Spirulinasubsalsa; Tp, Thalassiosira pseudonana;and Vc, Volvox carteri.

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Supplemental Fig. S3). Among prokaryotes, SMT15homologs were found in cyanobacteria and a subset ofproteobacteria but not in archaea (Fig. 1D; SupplementalFig. S3). A sequence alignment of family C SMT15 ho-mologs is shown in Supplemental Figure S4. A notablediscordance in the family C lineage is the split affiliationof the opisthokont members, where the choanoflagellate(M. brevicollis) homolog groups with heterokonts and thefungal homologs group with bacteria (Fig. 1D). Thisdiscordance is not observed in the family A1 lineage,where the fungal, choanoflagellate, and metazoan trans-porters form a monophyletic clade, as expected fornormal vertical inheritance of A1 family members. Mostof the SMT15 homologs identified in this study containthe three conserved domains found in SMT15, but thereare exceptions, such as Emiliania huxleyi (EOD04916)and P. infestans (EEY60920 and EEY64284), two of whichare missing a detectable cyclic nucleotide-binding do-main (Supplemental Table S1).

Complementation of smt15

Complementation was used to confirm that disruptionof the SMT15 gene by insertion of the paromomycinresistance marker was responsible for the previouslyreported smt15-1 phenotypes (Fang and Umen, 2008).The smt15-1mutant strain was transformed with plasmidpSMT15.1 containing the wild-type SMT15 gene includ-ing its predicted promoter region and sequences down-stream of its 39 untranslated region (see “Materials andMethods”). Four hundred fifty-eight transgenic lineswere generated, and three transformants (lines 57, 62,and 64) showed noticeable restoration of growth tovarious degrees (Table I). RT-PCR showed that SMT15mRNA abundance was restored to 2.4%, 19.4%, and78.9% of wild-type levels in lines 57, 62, and 64, re-spectively (Fig. 2). Line 64 (smt15-1 pSMT15.1), with78.9% restored SMT15 RNA, showed rescue to nearwild-type growth rate (Table I) and was used for furtherexperiments. The nonlinear correlation between mRNAlevels and growth rate in lines 57 and 62 suggests thatvery low (line 57) to low (line 62) expression levels ofSMT15 may be enough to alleviate some of the growthdefect. Line 64 was crossed to amat3-4 strain to generatea population of smt15-1 mat3-4 pSMT15.1 and smt15-1mat3-4 progeny. All the smt15-1 mat3-4 progeny thatreceived the pSMT15.1 had mat3-like small cell sizedistributions, while progeny that did not receive the

complementing plasmid showed suppression of themat3-4 small-cell phenotype (Table II), as reported pre-viously for smt15-1 mat3-4 double mutants (Fang andUmen, 2008). Taken together, these data confirmed thatdisruption of SMT15 is responsible for suppression ofthe mat3-4 size phenotype and for the slow-growthphenotype of smt15-1 single mutants.

SMT15 mRNA Levels Are Light Regulated

To investigate how SMT15 mRNA was regulatedand whether its accumulation was controlled by thecell cycle-dependent or by diurnal/circadian rhythms,we used RT-PCR to monitor its expression in samplescollected from synchronous cultures. Wild-type cells weresynchronized under a 14-h-light/10-h-dark (14L:10D)regime, and the culture synchrony was assessed bymeasurements of cell size, mitotic index, and periodicexpression of the S/M phase marker gene Cyclin-Dependent Kinase B1 (CDKB1; Fang et al., 2006). SMT15mRNA abundance was measured under the 14L:10Dsynchrony regime or in portions of the culture that wereremoved and darkened at 10 h or left in continuouslight. In the 14L:10D culture, SMT15 mRNA accumu-lated steadily during the light phase and remained el-evated until the dark phase, when it declined (Fig. 3A).In the culture that was darkened at 10 h, SMT15mRNAlevels declined 4 h earlier than in the 14L:10D culture,indicating that maintenance of the highest SMT15 ex-pression is light dependent. On the other hand, con-tinuous light could not maintain high expression ofSMT15 after 14 h, although it stayed higher than in bothdark cultures (Fig. 3A). To more directly examine theinfluences of light on SMT15 mRNA accumulation, itslevels were monitored in asynchronous cultures beforeand after dark incubation. As shown in Figure 3B,SMT15mRNA levels in cells growing under continuousillumination (T0) were comparable to samples collectedat 12 h from synchronized cultures. Importantly, thelevels of SMT15 transcript declined within 2 h after

Figure 2. Complementation of the smt15-1 mutant. Expression levelsare shown for SMT15 mRNA in the wild type (wt), smt15-1, and threeindependently generated smt15-1 transformants that were complementedwith pSMT15.1 (lines 57, 62, and 64).

Table I. Growth rates of complemented smt15-1 strains

SE values were derived from four or five independent experiments.

Genotype Doubling Time

hWild type 6.1 6 0.2smt15-1 9.1 6 0.5smt15-1 pSMT15.1 line 57 7.5 6 0.4smt15-1 pSMT15.1 line 62 7.5 6 0.6smt15-1 pSMT15.1 line 64 6.9 6 0.7










switching to dark (T2 and T4) and increased again afterreillumination (T6, T8, and T10), indicating that SMT15expression is light regulated. No correlation was foundbetween transcript levels of SMT15 and the S/M phasemarker CDKB1 in unsynchronized cultures.

The Sulfur Acclimation Response Is Affected insmt15-1 Cells

Because SMT15 is predicted to encode a potentialsulfate/anion transporter and might play a role in sul-fur uptake or response to sulfur limitation (Pootakhamet al., 2010), we asked whether the smt15-1 mutantshowed any aberrant responses to sulfur limitationand whether SMT15 mRNA abundance was regulatedby the availability of sulfur. Viability tests of wild-typeand smt15-1 cultures during sulfur starvation (–S) con-ditions showed an enhanced loss of viability in smt15-1after 3 d, when less than 50% of the mutant cells werealive compared with more than 80% of the wild-typecells (Fig. 4A). To determine whether the loss of viabilitywas specific to –S, mutant and wild-type cells werestarved for nitrogen (–N) and phosphate (–P) as well.smt15-1 had decreased viability relative to wild-type cellsin –N, although the defect was not as severe as in –S,while the viability of the mutant was not significantlyaffected by –P (Supplemental Fig. S5). We next examinedwhether SMT15 mRNA levels respond to nutrient dep-rivation. As shown in Figure 4B, SMT15 mRNA wastransiently elevated severalfold after switching to –Sconditions but showed much less change in responseto –P conditions. SMT15 mRNA was slightly elevated2 h after –N treatment. Because –N responses can bevery rapid (Boyle et al., 2012; Blaby et al., 2013), we alsomonitored SMT15 mRNA with short time intervals.Our results showed that less than 3-fold change was ob-served within the first 6 h of –N treatment (SupplementalFig. S6).

The reduced ability of smt15-1 to cope with –S andthe early and transient induction of SMT15 mRNA by–S suggested that SMT15 might participate in thesulfur starvation acclimation responses. To further testthis idea, RNA sequencing (RNA-seq) was used to de-termine genome-wide transcript abundance in the wildtype and smt15-1 under sulfur-replete (+S) conditions orafter 6 h in –S. Normally, SULFURACCLIMATION (SAC)genes are up-regulated to allow cells to cope with sulfur

starvation stress (Zhang et al., 2004; González-Ballesteret al., 2010). We found that the induction of SAC geneswas either attenuated or decreased in smt15-1 (Table III).The defects of smt15-1 in inducing SAC-responsive geneswere verified using quantitative RT-PCR and comparedwith those of the sulfur acclimation mutant sac1, which isseverely impaired in its SAC response (Davies et al., 1994;Table III; Fig. 4C). While the transcriptional induction ofSAC markers was almost completely blocked in the sac1mutant, it was only attenuated in smt15-1 compared withits full induction in wild-type cells (Fig. 4C).

Among the SAC-responsive genes that were mis-regulated, most notable were the genes encoding sulfuruptake and assimilation proteins, such as arylsulfatases(ARS1 and ARS2), sulfate transporters (SULTR2, SLT1,and SLT2), and ATP sulfurylases (ATS1 and ATS2; fordetails, see Table III). Instead of being up-regulatedafter –S, transcripts of sulfite reductases (SIR1 andSIR2) and serine acetyl transferase (SAT1) of smt15-1were down-regulated at least 40-fold. In the case ofSAT1, its transcript levels were too low to be detectedby quantitative RT-PCR after –S (data not shown).Transcripts encoding enzymes involved in sulfur as-similation but not classified as SAC-responsive genes,such as ADENYLYLPHOSPHOSULFATE REDUCTASE1and ADENOSINE 59-PHOSPHOSULFATE KINASE1(González-Ballester et al., 2010), showed little or nochange in abundance between wild-type and smt15-1strains in either +S or –S conditions (SupplementalTable S2).

SMT15mRNA levels did not show significant changein the sac1mutant in either –S or +S medium, indicatingthat its transcriptional regulation is independent ofSAC1 signaling (data not shown). Because smt15-1 has acell size defect, we examined the expression of corecell cycle genes under +S and –S conditions comparedwith the wild type but found no significant alterations(Supplemental Table S2). Although not related to de-fects in smt15-1, there was one cyclin-dependent kinase-encoding gene, CDKG2, whose transcript showed astrong up-regulation in the wild type under –S condi-tions. Whether CDKG2 up-regulation is part of the SACresponse or possibly a general stress response remainsto be determined.

General Defects in Sulfur Acclimation Do Not Affect CellCycle Regulation

To assess whether sac mutants in general have cellcycle defects or can suppressmat3mutants, we examinedtheir cell size and also generated double mutants withmat3-4. SAC1 encodes a protein that shares similaritywith ion transporters (Pollock et al., 2005) and has beenshown to be the major positive regulator of SAC inC. reinhardtii (Davies et al., 1996). SAC3 encodes a Ser/Thrprotein kinase that is related to the Snf1p kinase ofbudding yeast and has been shown to play a negativerole in controlling sulfur acclimation responses (Davieset al., 1999). Unlike smt15-1, neither sac1 nor sac3 showed

Table II. Modal sizes and SE of complemented smt15-1 mat3-5 andrelated strains

SE values were derived from five independent smt15-1 mat3-5pSMT15.1 strains, three independent smt15-1 mat3-5 strains, andthree independent mat3-5 strains.

Genotype Daughter Cell Size

mm3

mat3-5 23.5 6 2.0smt15-1 mat3-5 37.6 6 1.6smt15-1 mat3-5 pSMT15.1 26.5 6 2.5


Fang et al.









any cell size defects as single mutants (SupplementalTable S3). Moreover, sac1 mat3-4 and sac3 mat3-4 strainshad size distributions indistinguishable frommat3-4 singlemutants, meaning that these two SAC regulators could notsuppress mat3-4 when mutated (Table IV; SupplementalFig. S7). These results show that defects in the –S accli-mation pathway in general do not affect cell sizecontrol or the MAT3/RB pathway in C. reinhardtii. Al-though smt15-1 is defective in sulfur stress responses,its effect on cell size regulation and its ability to sup-press mat3-4 must be unrelated to general sulfur stressresponses.

Elevated Glutathione Levels in smt15-1 Affect the SulfurAcclimation Response

Because a sulfur acclimation response was activatedin –S conditions in smt15-1 but did not reach fullstrength, we suspected that the mutant’s effect on SACmight be indirect. A literature search for sulfur-relatedmetabolites that influence the cell cycle led us to hy-pothesize that glutathione levels might be involved inthe smt15-1 cell cycle and sulfur stress phenotypes(Diaz Vivancos et al., 2010a; Markovic et al., 2011;Noctor et al., 2012; García-Giménez et al., 2013). It has

Figure 3. Regulation of SMT15 tran-script abundance. A, Relative expres-sion level of SMT15 or CDKB mRNA inlight is marked by light blue columns.Relative expression levels of SMT15 orCDKB mRNA in the dark period of14L:10D (L/D = 14h/10h) or 10-h-light/14-h-dark (L/D = 10h/14h) cycles ismarked by black or dark blue columns,respectively. The light and dark phasesare indicated by white and black barsabove the graphs, respectively. The syn-chrony of the cell culture was monitoredby the cell division marker CDKB1. Dataare from technical triplicates and pre-sented as mean relative RNA levels 6 SE.The expression level of SMT15 at timezero was arbitrarily set to 1. B, Relativeexpression levels of SMT15 mRNA inunsynchronized cultures exposed to 4 hof dark. Samples from synchronized cul-tures collected at 0 h (Syn 0) and 12 h(Syn 12) after light exposure were used forcomparison. Relative expression levels ofCDKB1 mRNA were used to estimaterelative portions of cells in the S/M state.RNA levels are presented in logarithmicscale.









been reported that glutathione, one of the end pro-ducts of the sulfur assimilation pathway, is capable ofrepressing sulfur assimilation and uptake in plants(Herschbach and Rennenberg, 1994; Lappartient et al.,1999; Vauclare et al., 2002; Buchner et al., 2004). Ad-ditionally, glutathione has been shown to play a role incell cycle progression in plants and animals (Thelanderand Reichard, 1979; Menon et al., 2003; Menon andGoswami, 2007; Diaz Vivancos et al., 2010b). Therefore,we speculated that SMT15 might connect sulfur accli-mation to cell cycle control through the misregulation ofglutathione homeostasis. To test this idea, we compared

glutathione levels in wild-type, smt15-1, and sac1 strainsin +S and –S conditions. Under sulfur-replete condi-tions, smt15-1 accumulated approximately 50% higherlevels of glutathione than the wild type (33 versus 20pmol mg21). After 16 h, –S wild-type cells and sac1 cellsboth had glutathione levels of approximately 1.5 pmolmg21,while smt15-1 had glutathione levels that were 2-foldhigher (approximately 3 pmol mg21; Fig. 5A; Table V).To verify linkage between the SMT15 lesion and theglutathione defect, we monitored glutathione levels inthe progeny of crosses between smt15-1 and wild-typestrains. The smt15-1 mutation is caused by the insertion

Figure 4. A, Viability of wild-type (wt)and smt15-1 strains following S depri-vation in liquid medium. B, Expressionpatterns of SMT15 mRNA in 2S, 2P,or 2N medium. C, Relative expressionlevels of sulfur acclimation genes in –Sconditions in wild-type, smt15-1, andsac1 strains.


Fang et al.



Tab

leIII.

Comparativetran

scriptab

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sulfurac

clim

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rge

nes

(Gonzalez-Ballester

etal.,2010)from

wild-typ

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t15-1

strainsgrownin

thepresence

(+S)

andab

sence

(2S)

ofsulfurin

RNA-seq

andquan

titative

RT-PCRan

alyses

wt2

S/wt+S,

Fold

chan

geoftheselected

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scriptsforthewildtypein

+San

d–S

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t15-12S/wt+S,

fold

chan

geoftheselected

tran

scriptsbetwee

nthesm

t15-1

strain

in2Smed

ium

andthewild-typ

ein

+Smed

ium;sm

t15-12S/sm

t15-1+S,

fold

chan

geoftheselected

tran

scriptsbetwee

nthesm

t15-1

strain

in–S

med

ium

andsm

t15-1

in+Smed

ium;sm

t15-1+S/wt+S,

fold

chan

geoftheselected

tran

scriptsbetwee

nthesm

t15-1

strain

in+Smed

ium

andthewild-typ

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+Smed

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ange

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scriptsunder

specified

conditionswas

determined

byRNA-seq

(R)orquan

titative

RT-PCR(Q

).Values

arefrom

cellsex

posedto

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conditionfor0h(R0orQ0),2h(Q

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,ex

ceptforSA

T1(Cre10.g466750.t1.3).Asterisks

indicatethat

the

RNA-seq

datashowed

sign

ificantdifference

sin

fold

chan

gefrom

thequan

titative

RT-PCRan

alyses.NT,

Nottested

.

Iden

tifier

inVersion5

(Version4)

Nam

ean

dDescription

Fold

Chan

gein

Expression

RPKM

Values

wt2

S/wt+S

smt15-12S/wt+S

smt15-12S/sm

t15-1+S

smt15-1+S/wt+S

wt+S

wt2

Ssm

t15-1+S

smt15-12S

R6(Q

2,Q6)

R6(Q

2,Q6)

R6(Q

2,Q6)

R0(Q

0)

Sulfuruptake

Cre16.g671400.t1.2

(Cre16.g671400.t1.1)

ARS1

,periplasm

icarylsulfatase

3,307(3,452,5,637)

1,407(887,3,093)

703(830,2,891)

2.0

(1.1)

0.3

992

0.6

422

Cre16.g671350.t1.2

(Cre16.g671350.t1.1)

ARS2

,periplasm

icarylsulfatase

5,010(1,348,4,235)

560(22,60)

280(38,102)

2.0

(0.6)

0.2

1,002

0.4

112

Cre17.g723350.t1.2

(Cre17.g723350.t1.1)

SULTR2,H

+/SO

422tran

sporter

type

69(72,114.2)

2.8

(5.5,13)

17(7.3,17)

0.2

(0.8)

12

830

233

Cre12.g502600.t1.2

(Cre12.g502600.t1.1)

SLT1,Na+/SO

422tran

sporter

type

1430(534,958)

229(230,547)

23(60,142)

10(3.9)

0.7

1,001

7160

Cre10.g445000.t1.2

(Cre10.g445000.t1.1)

SLT2,Na+/SO

422tran

sporter

type

313(312,761)

101(425,366)

19(457,394)

5.3

(0.9)

3940

16

302

Cre07.g348600.t1.3

(Cre07.g348600.t1.2)

SLP1,ch

loroplastsulfate

permease

9.3

(11.3,13.6)

2.0

(6.0,1.6)

2.4

(10.2,2.8)

0.8

(0.6)

656

512

Cre14.g616900.t1.2

(Cre14.g616900.t1.1)

SLP2,ch

loroplastsulfate

permease

5.8

(5.6,4.8)

2.0

(4.3,2.0)

2.2

(8.2,3.8)

0.9

(0.5)

23

134

21

47

Cre06.g257000.t1.2

(Cre06.g257000.t1.1)

SLP3,sulfate-bindingprotein

5.1

(3.6,3.0)

1.3

(1.7,0.5)

1.4

(3.5,0.9)

0.9

(0.5)

15

76

14

19

Cre06.g273750.t1.3

(Cre06.g273750.t1.2)

SUA1,ch

loroplastsulfate

tran

sporter

7.7

(5.2,7.7)

2.5

(4.4,6.4)

2.2

(6.0,8.9)

1.1

(0.7)

20

154

22

49

Cre02.g145750.t1.1

(Cre02.g145750.t1.1)

AOT2,am

inoac

idtran

sporter

12(85,43)

3.0

(38,23)

4.5

(31,19)

0.7

(1.2)

336

29

Cre06.g298750.t1.2

(Cre06.g298750.t1.1)

AOT4,am

inoac

idtran

sporter

15(20,33)

2.8

(30,22)

4.5

(30,22)

0.6

(1.0)

27

396

17

76

Sulfurassimilation

Cre03.g203850.t1.2

(Cre03.g203850.t1.1)

ATS1

,ATPsulfurylase

5.4

(3.2,3.8)

1.5

(2.2,1.6)

2.8

(5.4,4.0)

0.5

(0.4)

292

1,584

155

429

Cre02.g107450.t1.2

(Cre02.g107450.t1.1)

ATS2

,ATPsulfurylase

10(7.4,12)

2.6

(5.8,3.3)

2.0

(6.3,3.5)

1.3

(0.9)

27

279

34

69

Cre16.g15615.t1

(Cre16.g693150.t1.2)

SIR1,ferredo

xin-sulfitereductase

4.9

(4.4,6.0)

,0.05(,

0.05,,0.05)

2.0

(,0.05,,0.05)

,0.05(0.9)

41

201

12

Cre16.g8568.t1

(Cre16.g693050.t1.2)

SIR2,ferredo

xin-sulfitereductase

3.3

(3.8,4.2)

,0.05(,

0.05,,0.05)

1.0

(,0.05,0.1)

,0.05(0.4)

53

173

0.1

0.1

Cre16.g685550.t1.2

(Cre16.g685550.t1.1)

ASL4,O-ace

tyl-Se

r(thiol)-

lyase/Cys

synthesis

9.2

(7.4,17)

3.1

(5.1,6.2)

4.0

(6.0,7.2)

0.8

(0.9)

140

1,294

109

437

Cre10.g466750.t1.3

(Cre10.g466750.t1.2)

SAT1,Se

rac

etyl

tran

sferase

33(26,42)

3.1*(,

0.05,,0.05)

6.0*(,

0.05,,0.05)

0.5

(1.0)

17

568

953

(Tab

leco

ntinues

onfollowingpag

e.)





of a paromomycin resistance marker in the SMT15 locus(Fig. 1A; Fang and Umen, 2008), and the paromomycinresistance phenotype was used to identify six smt15-1progeny along with six wild-type progeny that wereparomomycin sensitive. Elevated glutathione levels seg-regated with the paromomycin resistance phenotype inall 12 progeny, confirming that the smt15-1 mutation in-creases glutathione levels (Fig. 5B). Moreover, glutathionelevels decreased and became closer to those of the wildtype in the complemented smt15-1 strains under +S con-ditions (Fig. 5A; Table V). Restoration of glutathionelevels was more modest in the complemented lines versusthe parental smt15-1 strain, possibly due to amore stringentrequirement for SMT15 function under –S conditions.

Glutathione Levels Cycle in Synchronous Cultures

Glutathione levels have been reported to be cell cycleregulated in plants and animals (Menon and Goswami,2007; Diaz Vivancos et al., 2010a; García-Giménez et al.,2013). To investigate whether glutathione levels are cellcycle regulated in C. reinhardtii, we monitored glutathionecontent in synchronized cultures. In a synchronous cul-ture, cells normally divide at the end of the light period orthe beginning of the dark period, making it difficult touncouple the effects of light/dark transitions from cellcycle transitions. In order to circumvent this issue, wefirst synchronized cultures in a 12-h/12-h light/darkcycle and then released the synchronized cells intocontinuous light. We sampled the cultures starting attime zero, which corresponds to the end of the lastdark period, and continued through the completion ofthe cell cycle in continuous light. Culture synchronywas evaluated by scoring passage through commit-ment and mitotic index (Fig. 6A) and by measuring theperiodic expression of the cell cycle marker genesCDKB1 and Proliferating Cell Nuclear Antigen (PCNA;Fig. 6B). In synchronized wild-type cultures, glutathi-one concentration doubled (from approximately 13 toapproximately 28 pmol mg21) during the first fewhours in the light, reached its peak around the time ofcommitment, and then dropped gradually for the re-mainder of the cell cycle, reaching basal levels just beforeS/M (Fig. 6C). In synchronized smt15-1, glutathione ac-cumulation followed a similar pattern to the wild typeduring the first 8 h in the light, corresponding to early

Tab

leIII.(Continued

from

previouspage.)

Iden

tifier

inVersion5

(Version4)

Nam

ean

dDescription

Fold

Chan

gein

Expression

RPKM

Values

wt2

S/wt+S

smt15-12S/wt+S

smt15-12S/sm

t15-1+S

smt15-1+S/wt+S

wt+S

wt2

Ssm

t15-1+S

smt15-12S

R6(Q

2,Q6)

R6(Q

2,Q6)

R6(Q

2,Q6)

R0(Q

0)

Sulfurrecycling

Cre13.g569400.t1.2

(Cre13.g569400.t1.1)

TAUD1,taurindioxy

genase

72(60,223)

31(149,141)

19(69,66)

1.6

(2.2)

15

1,072

24

463

Cre13.g569500.t2.1

(Cre13.g569500.t1.1)

TAUD2,taurindioxy

genase

18(N

T)

8.0

(NT)

6.3

(NT)

1.3

(NT)

32

580

41

257

Cre07.g352550.t1.2

(Cre07.g352550.t1.1)

RDP3,putative

rhodan

ese

domainphosphatase

111(47,25)

42(23,16)

4.1

(8.7,6.2)

10.2

(8.7,2.6)

5553

51

209

Table IV. Modal daughter cell sizes and SE of mat3-4, smt15-1 mat3-4,sac1 mat3-4, and sac3 mat3-4 strains

SE values were derived from four independent experiments forsmt15-1 mat3-4 and sac3 mat3-4 strains and five independent ex-periments for mat3-4 and sac1 mat3-5 strains.

Genotype Daughter Cell Size

mm3

mat3-4 23.1 6 1.1smt15-1 mat3-4 38.4 6 3.4sac1 mat3-4 23.7 6 1.5sac3 mat3-4 19.6 6 1.3


Fang et al.



and mid G1 phases, but its levels did not decreasetoward the onset of cell division and instead remainedelevated during the time that glutathione levels droppedin wild-type cultures during S/M (Fig. 6C). These dataindicate that rhythmic glutathione accumulation insmt15-1 is defective and shows its greatest departurefrom the wild type during cell division (S/M).Glutathione can exist in an oxidized dimeric state

(GSSG) or a reduced monomeric state (GSH). To inves-tigate whether GSH-GSSG redox ratios are diurnally orcell cycle regulated, we measured them in synchronizedwild-type and smt15-1 cultures. In wild-type cultures, theGSH-GSSG ratio was approximately 16 in early G1 and

declined during the light period to between 5 and 6 at thetime of cell division (S/M; Fig. 6D). The temporal changesin GSH-GSSG ratios in synchronous smt15-1 cultures weresimilar to those in the wild type, except that the valueswere about double that of the wild type at every timepoint tested (Fig. 6D). Taken together, our data show thatsmt15-1 has both elevated total glutathione levels andelevated GSH-GSSG ratios compared with the wild type.

DISCUSSION

SMT15 Belongs to a Distinct Subfamily of PutativeSulfate/Anion Transporters

smt15-1 was isolated as a suppressor of the small cellsize defect of mat3 (Fang and Umen, 2008). Here, wecharacterized the smt15-1 mutant and verified thatdisruption of SMT15, which encodes a novel sulfate/anion transporter family member, caused growth andsize defects in C. reinhardtii. Phylogenetic analysis in-dicates that SMT15 belongs to a distinct subfamily ofsulfate/anion transporters whose origins are difficultto discern because of its patchy distribution amongeukaryotes and prokaryotes (Supplemental Fig. S3;Takahashi et al., 2012). However, the general phylogeneticcoherence we observed among prokaryotic and eukaryoticmembers suggests limited amounts of horizontal transferof SMT15-like/family C genes between kingdoms,with the only exception being the split affiliation ofthe opisthokont members, where fungal homologs areclosest to the eubacterial group while the choanoflagellatehomolog is closest to other eukaryotic members (Fig. 1D).Overall, the phylogeny of family C is consistent with theearly acquisition of SMT15/family C genes at the base ofthe eukaryotic lineage (Supplemental Fig. S2), perhapsthrough an endosymbiotic event, followed by multipleindependent losses.

Few members of tribe 1 superfamily transportershave been characterized. Some of them in the SLC26family function as Cl2/HCO3

2 transporters and othersare SO4

22 transporters (Satoh et al., 1998; Melvin et al.,1999; Soleimani et al., 2001; Wang et al., 2002). In-triguingly, down-regulation of a diastrophic dysplasiasulfate transporter (SLC26A2) is tightly associated withhigh rates of proliferation in colon cancer cells (Yusaet al., 2010), suggesting a connection between this familyof transporters and the control of cell proliferation. Plant

Figure 5. A, Normalized glutathione content (GSH + GSSG) of the wildtype (wt), smt15-1, sac1, and three independent complemented strains inthe presence of sulfur (+S) or after sulfur depletion for 16 h (2S). Strains thatshowed significant differences compared with the wild type (P , 0.05) arelabeled with single asterisks; complemented strains that showed significantdifferences (P , 0.05) compared with smt15-1 are labeled with two as-terisks. B, Normalized glutathione content (GSH + GSSG) of segregatedwild-type and smt15-1 progeny. MT+, Mating type plus; MT2, matingtype minus.

Table V. Glutathione concentrations of the complemented smt15-1 strains

SE values were derived from at least three independent experiments.

Genotype Total Glutathione Content (+S) Total Glutathione Content (2S)

pmol mg21

21gr (wild type) 20.0 6 0.7 1.5 6 0.4smt15-1 33.5 6 2.8 3.0 6 0.1sac1 17.5 6 1.1 1.2 6 0.1smt15-1 pSMT15.1 line 57 24.3 6 3.1 2.5 6 0.9smt15-1 pSMT15.1 line 62 23.7 6 2.3 2.3 6 0.3smt15-1 pSMT15.1 line 64 20.6 6 2.6 2.2 6 0.7







Figure 6. A, Graph showing passage through com-mitment (dashed lines) and mitotic index (solid lines)of synchronous cultures. Wild-type (wt) culture en-tered S/M phase at approximately 12 h; smt15-1 cul-ture entered S/M phase at approximately 10 h. Thecell cycle phases are indicated by bars above thegraph. The wild type is labeled in blue and smt15-1 islabeled in red. Because daughter cells of wild-type(6145-Y1) and smt15-1 strains failed to hatch fromthe mother cells in the first few hours after cultureswere switched to light, the cell size and commitmentassays were omitted at the early time points. Syn-chronized cultures were maintained in 12-h-light/12-h-dark cycles. Cultures were kept under light duringsampling. B, Expression of S/M phase markers PCNAand CDKB1 determined by quantitative RT-PCR, withSE shown by error bars. C, Total glutathione levels ofthe synchronized wild-type and smt15-1 strains withSE indicated. D, GSH-GSSG ratios of synchronizedwild-type and smt15-1 strains with SE indicated.


Fang et al.



transporters in family P are mainly SO422 transporters.

The rest of tribe 1 transporters remain uncharacterized.Our study represents, to our knowledge, the first phe-notypic characterization of a family C member outside ofbudding yeast, whose homolog is YGR125W, a sulfatetransporter domain (Pfam 00916) containing membraneprotein. Like smt15-1, YGR125W mutants have growthdefects as measured by competition assays (Breslowet al., 2008). Moreover, YGR125W interacts geneticallywith mutations in SUL1 and SUL2 that encode sulfatetransporters, but its specific roles in budding yeast insulfate metabolism or other aspects of cell physiologyhave not been investigated.

SMT15 and SAC Responses

The reduction of viability of smt15-1 cells in –S andits inability to activate a full SAC response indicates arole for SMT15 in sulfur starvation, but this role is likelyto be indirect. We found that smt15-1 overaccumulatesglutathione (Fig. 5A), whose elevated levels are known tosuppress the SAC response (Herschbach and Rennenberg,1994; Lappartient et al., 1999; Vauclare et al., 2002;Buchner et al., 2004). It is likely, therefore, that elevatedlevels of glutathione in smt15-1 strains attenuate theSAC response in –S conditions. Our RNA-seq dataset showed that mRNAs of g-GLUTAMYLCYSTEINESYNTHETASE and GLUTATHIONE SYNTHETASE,two major enzymes required for glutathione biosynthesis,were not altered significantly in smt15-1 (SupplementalTable S2), suggesting that SMT15 affects glutathionehomeostasis by a posttranscriptional mechanism.C. reinhardtii encodes several –S-inducible sulfate

transporters whose functions have been partiallycharacterized (Pootakham et al., 2010). SMT15 mRNAlevels were induced under –S conditions, but not to theextent of the messages for known sulfur transporters,such as SULTR2 and SLT1/2 (Table III). The trans-porters responsible for sulfate uptake under +S con-ditions are not known, but if the growth defects ofsmt15-1 in +S were due to inadequate sulfate uptake,it would be expected to show a constitutive SAC re-sponse under +S conditions, which is not the case(Fig. 4C; Table III). A plausible transport function forSMT15 might be in maintaining metal ion homeostasisthrough its function as a cotransporter (Lee et al., 2014;Srinivasan et al., 2014). Glutathione binds to and helpsdetoxify heavy metals and typically accumulates inresponse to elevated metals (Cobbett and Goldsbrough,2002; Jozefczak et al., 2012; Zagorchev et al., 2013), so itselevated levels in smt15-1 could reflect a response toaltered metal ion levels. An alternative possibility is thatSMT15 is not a transporter but a sensor/signalingprotein linked to sulfur metabolism, glutathione me-tabolism, and/or metal stress. Precedent for such afunction comes from the Sac1 protein of C. reinhardtii,which has homology to transporters but likely serves asa sensor to activate SAC responses (Davies et al., 1996;Pollock et al., 2005). Future work aimed at determining

the substrate(s) of SMT15 should help clarify its role insulfur metabolism and ion transport.

SMT15, Glutathione, and Cell Cycle Control

Our findings lead to the question of what substrates,if any, are transported by SMT15 and why defects inits function cause glutathione overaccumulation andcell cycle defects? Although smt15-1 has a clear sulfuracclimation defect, this defect is unlikely to be linkedto its effect on cell cycle control, because mutants withmore severe defects in SAC responses had no detect-able impact on the cell cycle (Table IV; SupplementalFig. S7). Our finding that glutathione overaccumulatesin smt15-1 but not in sac mutants is a possible clue forunderstanding the cell cycle defects in this mutant.Glutathione levels and subcellular localization havebeen linked to cell cycle control in both plants andanimals (Markovic et al., 2007; Pallardó et al., 2009;Pellny et al., 2009; Diaz Vivancos et al., 2010a, 2010b),although its specific impact on cell cycle-related pro-cesses is not completely understood. Changing the redoxstate of cell cycle regulators through glutathionylationhas also been shown to influence the cell cycle (Chiuand Dawes, 2012). A particularly striking finding inour study was the cell cycle-regulated fluctuations inglutathione levels and their disruption in smt15-1. Insynchronized wild-type cells, glutathione levels peakedin mid G1 and then declined during cell division,while in smt15-1, they remained elevated (Fig. 6C).The glutathione redox ratio (GSH-GSSG) also fluctuatedduring the wild-type cell cycle, reaching its lowestlevels around the time of cell division (Fig. 6D). Whilethis cyclical pattern was mirrored in smt15-1, the mu-tant GSH-GSSG ratio was consistently around 2-foldhigher than in the wild type, suggesting that a morereducing cellular environment occurs in smt15-1 cellsfor reactions that are in equilibrium with glutathione.Based on these findings, we speculate that aberrantglutathione levels and redox homeostasis cause the in-creased division and small-cell phenotypes observed inthe smt15-1 mutant. Supporting this hypothesis arenumerous reports on the close association between cellproliferation and elevated glutathione levels and highGSH-GSSG ratios (Mauro et al., 1969; Kosower andKosower, 1978; Suthanthiran et al., 1990; Sánchez-Fernández et al., 1997; May et al., 1998; Nkabyo et al.,2002). We have not been able to examine temporal redoxcontrol at a finer scale in wild-type and smt15-1 cells,as has been done in budding yeast, where redox andrespiratory activity cycle with a period of about 40 min(Tu et al., 2005). It may be revealing to examineC. reinhardtii cells at these time scales to see if thereare shorter scale periodicities in its redox metabolism.

In Arabidopsis (Arabidopsis thaliana), glutathione levelswere found to cycle in synchronized cell cultures butappeared to peak during cell division, which is differentfrom what we found for C. reinhardtii, where glutathionelevels were near their lowest levels during cell division









(Fig. 6C). However, the increased glutathione duringS and G2 phases occurred in a partially synchronouscell population generated from starvation and refeeding(Diaz Vivancos et al., 2010b). This raises the question ofwhether the peak of glutathione seen was due to cellcycle phasing or is a general response to proliferationinduced by refeeding starved cultures. A similar caveathas been raised for measurements of glutathione in syn-chronous mammalian cell culture experiments (Markovicet al., 2010). In our hands, starvation and refeedingof C. reinhardtii also caused a transient increase inglutathione levels that are unrelated to cell cycle phasing(Supplemental Fig. S8). Therefore, it is premature to con-clude that our results conflict with those from Arabidopsisor other organisms in which glutathione cycling hasbeen examined with respect to the cell cycle.

Although the established connections betweenglutathione-mediated redox state and cell cycle controlmake redox defects a plausible explanation for the cellcycle phenotypes of smt15-1, we cannot exclude the pos-sibility that other defects account for its cell size phenotype.It is clear that the mutant does not cause the misregulatedexpression of core cell cycle genes (Supplemental Table S2)and is most likely causing a posttranscriptional changein cell cycle control. The molecular mechanism con-necting glutathione flux and cell cycle control remainsto be determined.

Even though the accumulation of SMT15 mRNA wasin sync with cell cycle phasing (Fig. 3A), it is also lightregulated. It is well known that light reactions of pho-tosynthesis generate abundant reactive oxygen species(Balsera et al., 2014; Schmitt et al., 2014) and may acti-vate the transcription of SMT15 in order to maintainglutathione homeostasis and thereby mitigate cellulardamage and stress caused by reactive oxygen species.Equally possible is that glutathione levels are directlycoupled to the glutathionylation/deglutathionylation ofproteins required for photosynthetic activities as cellsgrow (Zaffagnini et al., 2012; Michelet et al., 2013).

Recent studies on cancer cell metabolism indicatethat cancer cells are metabolically different from normalcells. The increased demand of glutathione is importantfor RB-defective cells to cope with oxidative stress(Nicolay et al., 2013; Reynolds et al., 2014). At least inthe mouse system, increases in g-glutamyl-Cys ligasepartly contribute to increased glutathione levels in RB2/2

cancer cells (Reynolds et al., 2014). The connection be-tween increased cell division and defective glutathionecontrol in smt15-1 and tumor cells suggests that cellcycle-dependent cellular redox regulation may be similarin both systems, and smt15-1 may provide a unique toolfor understanding redox-regulated cell cycle control.

MATERIALS AND METHODS

Strains and Growth Conditions

Chlamydomonas reinhardtii strains 21gr (CC1690, mating type plus [MT+]),6145-Y1 (CC1691-Y1, mating type minus [MT2]), smt15-1 (Fang and Umen, 2008),sac1 (CC3801, MT+; Davies et al., 1996), and sac3 (CC3799, MT+; González-Ballester

et al., 2008) were used. sac1 and sac3 were ordered from the C. reinhardtii stockcenter (http://chlamycollection.org/). 6145 (CC1691) was backcrossed to 21grtwice to remove a y1 mutation (Bednarik and Hoober, 1985; White and Hoober,1994) and was then designated as 6145-Y1 (CC1691-Y1). 6145-Y1 was used in theexperiment presented in Figure 6. 21gr or mutants derived from the 21gr strainbackground were used in the majority of experiments. sac1 and sac3 werebackcrossed to 6145-Y1, and MT2 sac progeny were backcrossed to 21gr togenerate sac1-g and sac3-g progeny of both mating types. sac1-g and sac3-g wereused for the experiments in this study.

For segregation analysis, bulk meiotic progeny were germinated on high-saltmedium (HSM) plates, resuspended and serially diluted in Tris-acetate-phosphate(TAP), and plated for single colonies to obtain meiotic segregants.

Cells were grown in HSM (Sueoka, 1960) or TAP (Gorman and Levine, 1965)under illumination at 250 to 300 mmol photons m22 s21 aerated with 0.5% (v/v)CO2. Culture synchronization was induced by growth in 12-h-light/12-h-darkcycles for several weeks. The cultures were then transferred to 14L:10D cycles tooptimize synchrony. Synchronized cultures were maintained in 14L:10D cyclesunless mentioned otherwise.

For –S treatment, the sulfate salts in HSM was replaced with chloride salts(HSM-S). Sulfur-free trace elements were prepared as described by the C. reinhardtiiResource Center (http://chlamycollection.org/hutners-trace-elements-recipe).For sulfur deprivation experiments, the cells were grown to logarithmic phasein +S medium, washed twice with 250 mL of HSM-S, and then resuspended inHSM-S.

Identification of Genomic DNA Sequences Flanking thesmt15-1 Insertion

The isolation of smt mutants was described previously (Fang and Umen,2008). The following protocol for identifying insertion sites was provided bySteve Pollock (personal communication) with details to be published elsewhere.Briefly, genomic DNA was digested with SmaI to generate blunt-ended fragments.A blunt-ended asymmetric adaptor consisting of a 48-bp DNA oligonucleotideand a 10-bp oligonucleotide was then ligated to the digested genomic DNA. Aninsert-specific primer, IMP3-1 (59-CGATTTCGGCCTATTGGTTA-39), and anadaptor primer, AP1 (59-GTAATACGACTCACTATAGAGT-39), were used toamplify the genomic flanking region adjacent to the smt15-1 insertion. The insert-specific primer IMP3-2 (59-ATTTCCATTCGCCATTCAGG-39) and the adaptorprimer AP2 (59-ACTATAGAGTACGCGTGGT-39) were used for nested PCR. PCRfragments were amplified by Taq DNA polymerase in a final volume 20 mL in thepresence of 13 ExTaq buffer (Takara Bio), 1 mM primers, 80 mM deoxyribonucleotidetriphosphate (dNTP), and 2% (v/v) dimethyl sulfoxide (DMSO). PCR conditionswere as follows: 96°C for 3 min, followed by 36 cycles of 94°C for 30 s, 60°C for30 s, and 72°C for 2 min. PCR products were gel purified and sequenced.

Strain Genotyping

One microliter of genomic DNA prepared as described (http://www.chlamy.org/methods/quick_pcr.html) was used for PCR amplification. PCRfragments were amplified using Taq DNA polymerase in a final volume of20 mL in the presence of 13 ExTaq buffer (Takara Bio), 1 mM primers, 80 mM dNTP,0.5 M betaine, and 3% (v/v) DMSO. Primer pairs used for PCR-based genotypingare listed in Supplemental Table S4. PCR conditions were as follows: 96°C for2 min, followed by 42 cycles of 94°C for 30 s, 65°C for 30 s, and 72°C for 45s.

RACE-PCR and Isolation of the Full-Length SMT15 cDNA

Total RNA was isolated as described previously (Fang et al., 2006). Fivemicrograms of total RNA was used for cDNA synthesis. cDNA was synthesizedwith a mixture of oligo(dT) and random primers (9:1 ratio) at 55°C for 70 minusing the ThermoScript RT-PCR kit (Invitrogen) following the manufacturer’s in-structions. PCR was used to amplify different parts of the SMT15 cDNA under thefollowing conditions: 20-mL RT-PCRwith 0.5 mL of cDNA, 4 mL of 53 Phusion HFbuffer, 200 mM dNTPs, 0.5 mM primers, 3% (v/v) DMSO, 0.5 M betaine, and 0.4 unitPhusion High-Fidelity DNA polymerase (New England Biolabs). PCR amplifica-tion conditions were as follows: 98°C for 1 min, and then 40 cycles of 98°C for 10 s,65°C for 20 s, and 72°C for 35 s. An approximately 1.78-kb SMT15 cDNA fragmentwas amplified using primers 59-ATGCCGCCTCAGCTAACGCAC-39 and59-CTGGAATGCGAAGTAGAGTG-39. An approximately 1.32-kb SMT15 cDNAfragment was amplified using primers 59-AAGCTGTGGGAGCTGTTCAA-39 and59-TTGGCGAAGATGACGTTGAC-39. The amplified cDNA fragments were thencloned into pGEM-T easy vector (Promega) separately and sequenced.


Fang et al.




http://chlamycollection.org/

http://chlamycollection.org/hutners-trace-elements-recipe

http://www.chlamy.org/methods/quick_pcr.html

http://www.chlamy.org/methods/quick_pcr.html



The 39 RACE-PCR was carried out using the GeneRacer Kit with SuperScriptIII reverse transcriptase according to the manufacturer’s instructions(Invitrogen). 59-ATCTCACGGGACTGGCTCAT-39 and GeneRacer 39 primers(provided by the manufacturer) were used to amplify the 39 end of SMT15 underthe following conditions: 40 cycles of 98°C for 10 s, 65°C for 20 s, and 72°C for 50s. The three overlapping SMT15 cDNA fragments were assembled by restrictionenzyme digestion followed by DNA ligation to obtain a full-length SMT15cDNA. The assembled SMT15 cDNA was sequenced to verify its accuracy.RACE-PCR failed to amplify the 59 end of the SMT15 cDNA. However, a cDNAfragment containing the predicted start codon of SMT15 gene model g809.t1(based on the C. reinhardtii version 5 assembly at http://www.phytozome.net/chlamy.php) was amplified using primers 59-ATGAGTTTGGGCAAGCGCTCGT-39and 59-CGGCATCAGGCTTCCTACAAC-39 and verified by sequencing.

Phylogenetic Tree Construction

To construct the phylogenic tree in Figure 1D, the SMT15 protein sequence(KF709427) was used for a BLASTP query of the National Center for Bio-technology Information (NCBI). The high-scoring positives (E-value of 1e-25was set as an arbitrary cutoff) were selected to generate Figure 1D. An inde-pendent BLAST search on the Phytozome database (http://www.phytozome.net/) was conducted to check for the presence of SMT15 homologs in othergreen algae. The locus numbers or gene identification numbers in Figure 1Dare indicated according to Joint Genome Institute (http://www.jgi.doe.gov/),Phytozome (http://www.phytozome.net/), Saccharomycetes Genome Data-base (http://yeastgenome.org/), The Arabidopsis Information Resource(http://www.arabidopsis.org/), or NCBI (http://www.ncbi.nlm.nih.gov/protein/). Locus numbers or gene identification numbers used to generatephylogenetic trees are listed in Supplemental Table S1. Members of familiesA1, A2, and P from tribe 1 sulfate/anion transporters (Takahashi et al., 2012),including Aspergillus niger, Arabidopsis (Arabidopsis thaliana), C. reinhardtii,Coccomyxa subellipsoidea, Monosiga brevicollis, Saccharomyces cerevisiae, andVolvox carteri, were included as an outgroup for tree construction. For thefamily C group, proteins containing both the sulfate permease domain (Pfam00916) and the STAS domain (Pfam 01740 or cd07042) were selected for treeconstruction. Full-length protein sequences were aligned by ClustalW. Theresulting alignments were used to construct phylogenetic trees in MEGA 5.22(Tamura et al., 2011). The neighbor-joining method was used to generatephylogenetic trees, and 1,000 replicates were used for bootstrapping. Bootstrapvalues of 50% or higher were shown for each clade. The evolutionary distanceswere computed using the JTT matrix (Jones et al., 1992).

To search for distant SMT15 homologs in different phyla (SupplementalFig. S2), SMT15 or its homologs were BLAST searched against nonredundantprotein sequences, ESTs, and transcriptome assemblies in NCBI. E values of 1e-15 orless were set as an arbitrary cutoff, and candidates containing both thesulfate permease and STAS domains were used to map the phylogeneticdistribution of family C members in Supplemental Figure S2. The tree inSupplemental Figure S2 is based on findings by Baldauf (2003) and Gupta(2005).

Complementation of smt15-1

Bacterial artificial chromosome clone pTQ5801 (generated by Dr. PeterLefebvre at the University of Minnesota and available from the ClemsonUniversity Genomics Institute) was digested with BstBI andNheI, and a 13.675-kbDNA fragment containing the predicted SMT15 gene was gel purified and ligatedto BstBI- and NheI-digested pSP124 plasmid that had been modified tocontain BstBI and NheI restriction enzyme sites (Lumbreras et al., 1998) togenerate pSMT15.1. BstBI-linearized pSMT15.1 was transformed into smt15-1 byelectroporation (http://nutmeg.easternct.edu/~adams/ChlamyTeach/experiments.html), and transformants were selected on TAP containing 5 mg mL21 zeocin(Invitrogen).

Growth Rate Measurements

Liquid cultures were grown in continuous light, and cell density wasmaintained between 105 to 106 cells mL21 by dilution into fresh HSM. Culturesat around 1 to 2 3 105 cells mL21 were used for the initial sampling point, andadditional samples were collected every 3 h for 12 h. Chlorophyll content wasused as a measure of culture density as described previously (Harris, 1989),and each growth experiment was repeated at least three times.

Dark-Shift Experiments and Cell-Size Measurements

Dark-shift experiments and daughter cell size measurements were per-formed as described previously (Fang and Umen, 2008). Briefly, liquid cultureswere grown under continuous light in HSM, and cell density was maintainedbetween 105 and 106 cells mL21 before dark incubation. After incubatingcultures in the dark for 16 to 18 h, cells were fixed with a final concentration of0.2% (v/v) glutaraldehyde and 0.005% (v/v) Tween 20. Cell size distributionswere measured using a Coulter Counter (MULTISIZER 3; Beckman-Coulter)set to count at least 300 events in the modal channel. Modal daughter cell sizewas determined from at least three independent cultures as described previ-ously (Fang and Umen, 2008).

Cell Viability Analysis

A total of 1 3 106 cells were collected by brief centrifugation at 2,000g in amicrocentrifuge tube. Cells were resuspended in 1 mL of HSM with 5 mM ofthe vital fluorescent dye carboxymethylfluorescein diacetate (CMFDA; Invitrogen;Johnson et al., 2013). Cells were kept in the dark at room temperature for 30to 40 min and then washed two times with phosphate-buffered saline (PBS).Fluorescence and differential interference contrast images were taken on aZeiss Axio Scope A1 fluorescence microscope equipped with an AxioCamHRc camera (Zeiss). Viability was measured as (number of fluorescent cells)/(number of total cells)3 100%. Glutaraldehyde-treated or heat-treated cells wereused as negative controls and showed no signal after CMFDA staining. Theviability data we obtained with CMFDA staining was consistent with what wasdescribed previously using a different method (Chang et al., 2005).

RNA-seq

Approximately 30 mg of DNA-free RNA samples isolated from wild-type(21gr) and smt15-1mutant cells 6 h after being transferred to2S or +S mediumwas submitted to Genomics BioSci & Tech, Taiwan (http://www.genedragon.com.tw/site_ngs.php#menu03). Total RNA was converted into a librarytemplate for sequencing using an mRNA sequencing sample preparation kit(catalog no. RS-930-1001; Illumina). Briefly, mRNA was isolated by Sera-MagMagnetic Oligo(dT) Beads, washed, and then fragmented using divalent cat-ions under elevated temperature. First and second strand cDNAs were syn-thesized and end repaired to convert overhangs into blunt ends. The 39 endswere then adenylated for ligation of the Illumina adapters. cDNA templateswere purified by gel isolation for a size selection of approximately 2006 25 bpand amplified via PCR. Products were isolated by the QIAquick PCR Purifi-cation Kit (Qiagen) and added to the flow cell for sequencing. IlluminaHiSeq2000 technology was used. The raw sequencing data were filtered toremove low-quality sequences, including ambiguous nucleotides, adaptor se-quences, and repeat sequences. Processed sequence files from the Illuminapipeline output were aligned against the version 4.1 assembly of the C. reinhardtiigenome using Bowtie software (Langmead et al., 2009). One mismatch wasallowed during the alignment. More than 76% of the reads were mapped toannotated transcripts of the version 4.1 C. reinhardtii genome. The statistics onread analyses are summarized in Supplemental Table S5. Expression estimatesfor each sample were provided in units of reads per kilobase of exon per millionaligned reads (RPKM; Mortazavi et al., 2008). Two-fold or 4-fold differenceswere used as cutoffs for identifying differentially expressed genes. Genes withdifferential expression patterns were subjected to Gene Ontology and KyotoEncyclopedia of Genes and Genomes pathway analyses using the online AlgalFunctional Annotation Tool (Lopez et al., 2011; http://pathways.mcdb.ucla.edu/algal/index.html).

Quantitative PCR

One or 3 mg of total RNA was used for cDNA synthesis. DNA-free RNAwas reverse transcribed in the presence of a mixture of dT and randomprimers (9:1 ratio) using Thermoscript reverse transcriptase (Invitrogen)according to the manufacturer’s instructions. Ten-microliter RT-PCR con-tained 2.5 mL of 1:20 diluted cDNA, 0.2 or 0.25 mM primers, and 5 mL of 23KAPA SYBR FAST master mix (KAPA Biosystems). Real-time PCR was per-formed using a CFX96 Real-Time PCR detection system (Bio-Rad). Quantita-tive analysis was calculated based on the comparative cycle threshold method22ΔΔCT using G protein Beta Subunit-Like Protein as an internal standard by CFXManager Software following the manufacturer’s instructions (Bio-Rad).Primers used for quantitative PCR are listed in Supplemental Table S4.




http://www.phytozome.net/chlamy.php

http://www.phytozome.net/chlamy.php

http://www.phytozome.net/


http://www.jgi.doe.gov/


http://yeastgenome.org/

http://www.arabidopsis.org/

http://www.ncbi.nlm.nih.gov/protein/

http://www.ncbi.nlm.nih.gov/protein/






http://nutmeg.easternct.edu/~adams/ChlamyTeach/experiments.html

http://nutmeg.easternct.edu/~adams/ChlamyTeach/experiments.html

http://www.genedragon.com.tw/site_ngs.php#menu03

http://www.genedragon.com.tw/site_ngs.php#menu03


http://pathways.mcdb.ucla.edu/algal/index.html

http://pathways.mcdb.ucla.edu/algal/index.html



Glutathione Measurements

For total glutathione measurements, 2 3 107 cells were collected fromC. reinhardtii cultures and washed with 13 PBS twice. The cell pellet was thenresuspended in 500 mL of cold 5% (w/v) metaphosphoric acid and homogenizedby vortexing in the presence of 300 mL of zirconium beads (BioSpec Products) at4°C for 30 min. The suspension was transferred to a 1.5-mL tube and centrifugedat 12,000g to 14,000g for 5 min at 4°C. The supernatant was transferred into aclean 1.5-mL tube and stored at 280°C before analysis. Total glutathionewas measured enzymatically using a glutathione detection kit (Enzo LifeSciences). Briefly, glutathione reductase was added to reduce GSSG to GSH.Total GSH reacts with 5,59-dithiobis-2-nitrobenzoic acid and produces ayellow 5-thio-2-nitrobenzoic acid that absorbs at 405 nm. GSH content wasextrapolated based on a standard curve generated by GSSG with a knownconcentration (provided in the kit). Total glutathione was normalized tototal protein content. For protein content measurement, 1 3 107 cells werecollected and heated in 2% (w/v) SDS solution at 100°C for 10 min. A 2-folddilution of the crude protein extract was used for protein content measurement.Protein content was measured with a detergent-compatible DC protein assay kit(Bio-Rad) following the manufacturer’s instructions. To determine the ratio ofGSH to GSSG, total glutathione and GSSG were measured. A total of 7.5 3 108

cells were collected and washed with 13 PBS twice. Cell pellets were resus-pended in 1,000 mL of cold 5% (w/v) metaphosphoric acid (Sigma) andhomogenized by vortexing in the presence of 500 mL of zirconium beads at4°C for 45 min. The suspension was transferred to a 1.5-mL tube andcentrifuged at 12,000g to 14,000g for 5 min at 4°C. The supernatant wastransferred to a clean 1.5-mL tube and stored at 280°C before analysis. Formeasurement of GSSG, 1 mL of 2 M 4-vinylpyridine (Sigma) was added per50 mL of sample before measurement. 4-Vinylpyridine blocks free thiolspresent in the reaction, thus eliminating any contribution of GSH. Sampleswithout 4-vinylpyridine treatment were used for measurement of totalglutathione. Total glutathione and GSSG were measured using a glutathionedetection kit (Enzo Life Sciences) as described previously. The ratio of GSHto GSSG was deduced from (total glutathione 2 GSSG)/GSSG.

Supplemental Data

The following materials are available in the online version of this article.

Supplemental Figure S1. Prediction of transmembrane topology of SMT15protein by TransMembrane prediction using Hidden Markov Models(TMHMM 2.0).

Supplemental Figure S2. ClustalW alignment of the sulfate transporterdomain (Pfam 00916) of listed proteins in Figure 1.

Supplemental Figure S3. Diverse distribution of C family SMT15 homologsin eukaryotic and prokaryotic phyla.

Supplemental Figure S4. ClustalW alignment of parts of Family C sulfate/anion transporters.

Supplemental Figure S5. Viability of wild-type and smt15-1 strains followingN deprivation or P deprivation in liquid medium.

Supplemental Figure S6. Expression patterns of SMT15 mRNA undernitrogen-deplete conditions.

Supplemental Figure S7. Size distributions of dark-shifted cells from sac1mat3-4, sac3 mat3-4, and mat3-4.

Supplemental Figure S8. Glutathione concentration and mitotic index ofwild-type cells after being diluted into fresh medium.

Supplemental Table S1. Accession number, locus number, or gene identities,and structure organization of representative proteins used for constructingthe phylogenic tree.

Supplemental Table S2. Comparative quantification of transcript abundance(RPKM) of selected cell cycle genes and genes involved in S assimilationand glutathione biosynthesis by RNA-seq.

Supplemental Table S3. Daughter cell size distribution of wild-type, sac1,and sac3 strains.

Supplemental Table S4. List of primer pairs used for quantitative RT-PCR.

Supplemental Table S5. Statistics of RNA-seq reads.

ACKNOWLEDGMENTS

We thank Dr. Wirulda Pootakham (Carnegie Institution for Science) fortechnical advice on sulfur acclimation responses, Dr. Stéphane Lemaire (CentreNational de la Recherche Scientifique, Université Pierre et Marie Curie) fortechnical advice on glutathione extraction, and Dr. Kuo-Chen Yeh (AgriculturalBiotechnology Research Center, Academia Sinica) for comments on the article.We also thank Yen-Ju Lin and Chung-Ya Liao (Industrial Technology ResearchInstitute) for assistance in the analysis of RNA-seq data.

Received September 25, 2014; accepted October 29, 2014; published October31, 2014.

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Defects in a New Class of Sulfate/Anion Transporter Link ...defects in sulfur metabolism do not...

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