Cytochrome cM Decreases Photosynthesis underPhotomixotrophy in Synechocystis sp. PCC 68031[CC-BY]

Daniel Solymosi,a Lauri Nikkanen,a Dorota Muth-Pawlak,a Duncan Fitzpatrick,a Ravendran Vasudevan,b

Christopher J. Howe,b David J. Lea-Smith,b,c and Yagut Allahverdiyevaa,2,3

aLaboratory of Molecular Plant Biology, Department of Biochemistry, University of Turku, Turku FI–20014,FinlandbDepartment of Biochemistry, University of Cambridge, Cambridge CB2 1QW, United KingdomcSchool of Biological Sciences, University of East Anglia, Norwich NR4 7TJ, United Kingdom

ORCID IDs: 0000-0003-3655-3654 (D.S.); 0000-0002-7192-9322 (L.N.); 0000-0001-7005-2007 (D.M.-P.); 0000-0001-9116-1392 (D.F.);0000-0001-9195-1116 (R.V.); 0000-0002-6975-8640 (C.J.H.); 0000-0003-2463-406X (D.J.L.-S.); 0000-0002-9262-1757 (Y.A.).

Photomixotrophy is a metabolic state that enables photosynthetic microorganisms to simultaneously perform photosynthesisand metabolism of imported organic carbon substrates. This process is complicated in cyanobacteria, since many, includingSynechocystis sp. PCC 6803, conduct photosynthesis and respiration in an interlinked thylakoid membrane electron transportchain. Under photomixotrophy, the cell must therefore tightly regulate electron fluxes from photosynthetic and respiratorycomplexes. In this study, we demonstrate, via characterization of photosynthetic apparatus and the proteome, thatphotomixotrophic growth results in a gradual inhibition of QA

2 reoxidation in wild-type Synechocystis, which largelydecreases photosynthesis over 3 d of growth. This process is circumvented by deleting the gene encoding cytochrome cM(CytM), a cryptic c-type heme protein widespread in cyanobacteria. The DCytM strain maintained active photosynthesis overthe 3-d period, demonstrated by high photosynthetic O2 and CO2 fluxes and effective yields of PSI and PSII. Overall, thisresulted in a higher growth rate compared to that of the wild type, which was maintained by accumulation of proteins involvedin phosphate and metal uptake, and cofactor biosynthetic enzymes. While the exact role of CytM has not been determined, amutant deficient in the thylakoid-localized respiratory terminal oxidases and CytM (DCox/Cyd/CytM) displayed a phenotypesimilar to that of DCytM under photomixotrophy. This, in combination with other physiological data, and in contrast to aprevious hypothesis, suggests that CytM does not transfer electrons to these complexes. In summary, our data suggest thatCytM may have a regulatory role in photomixotrophy by modulating the photosynthetic capacity of cells.

Switching between different trophic modes is anadvantageous feature that provides great metabolicflexibility for cyanobacteria. For a long time, these pho-tosynthetic prokaryotes were considered to be predom-inantly photoautotrophic organisms (Smith 1983; Staland Moezelaar 1997). Lately, accumulating evidencemarks the physiological and ecological importance of

trophic modes involving organic carbon assimilation,e.g. photomixotrophy (Zubkov and Tarran 2008; Moore,2013). Dissolved organic carbon, most notably mono-saccharides, including Glc and Fru, accumulates in theenvironment mainly during phytoplankton blooms(Ittekkot et al., 1981; Teeling et al., 2012). During pho-tomixotrophy, photosynthetic organisms must balancethe consumption of organic carbon sources with photo-synthesis and carbon fixation.

In the model cyanobacterium Synechocystis sp. PCC6803, photomixotrophy is further complicated by theoperation of anabolic and catabolic processes occurringin the same cellular compartment and by the presenceof an interlinked thylakoid membrane-localized elec-tron transport pathway involved in both photosynthe-sis and respiration (Vermaas, 2001; Mullineaux, 2014;Lea-Smith et al., 2016a). In Synechocystis, photosyn-thetic linear electron flow is similar to other oxygenicphotoautotrophs. In PSII and PSI, the energy of theharvested photons induces charge separation. Electronsfrom the PSII primary donor P680 pass via pheophytinand the primary quinone QA to the secondary quinone,QB. Oxidized P6801, which driveswater splitting on theluminal side of PSII, is the strongest biological oxidizingmolecule. When QB is doubly reduced, it binds two

1This work was supported by the Academy of Finland (project no.315119 to Y.A. and Finnish Center of Excellence project no. 307335),the NordForsk Nordic Center of Excellence “NordAqua” (no. 82845),and the Waste Environmental Education Research Trust (C.J.H.).

2Author for contact: allahve@utu.fi.3Senior author.The author responsible for distribution of materials integral to the

findings presented in this article in accordance with the policy de-scribed in the Instructions for Authors (www.plantphysiol.org) is:Yagut Allahverdiyeva (allahve@utu.fi).

D.S. and Y.A. designed the research; D.S. performed the majorityof the experiments; D.M.P. and D.S. performed and analyzed proteo-mics data; L.N. performed Cytf kinetic measurements and immuno-blotting; D.J.L.-S. constructed the mutant strains; all authorscontributed to analysis of the data; D.S., Y.A., and D.J.L.-S wrotethe article; and all authors revised the manuscript.

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protons from the cytosol, converting plastoquinone(PQ) to plastoquinol (PQH2), which then diffuses intothe membrane PQ pool. Cytochrome (Cyt) b6f receivestwo electrons from PQH2 and transfers an electron tothe mobile small protein, plastocyanin (Pc) or cyto-chrome c6 (Cyt c6). An electron is subsequently trans-ferred to PSI, replacing a newly excited electron that istransferred from the PSI reaction center P7001 viaseveral co-factors to ferredoxin (Fed). Lastly, electronsare transferred from Fed to NADP1 by Fed-NADP1reductase (FNR) to generate NADPH. In the respiratoryelectron transfer pathway, PQ is reduced by NAD(P)Hdehydrogenase-like complex I (NDH-1) and succinatedehydrogenase (SDH), using electrons ultimately de-rived from Fed (Schuller et al., 2019) and succinate,respectively. Electrons from the PQ pool can be trans-ferred to a thylakoid-localized respiratory terminaloxidase (RTO), cytochrome bd-quinol oxidase (Cyd), or,via Cyt b6f and Pc/Cyt c6 to a second RTO, an aa3-typecytochrome c oxidase complex (Cox). How Synecho-cystis regulates electron input from PSII and the NDH-1 and SDH complexes into the photosynthetic electrontransport chain and to RTOs under photomixotrophicconditions is not fully understood. Moreover, Synecho-cystis encodes four isoforms of the flavodiiron proteins(FDPs), Flv1 to Flv4, which likely utilize NAD(P)H(Vicente et al., 2002; Brown et al., 2019) or reduced Fed(Santana-Sanchez et al., 2019). These proteins functionin light-induced O2 reduction as heterooligomers con-sisting of Flv1/Flv3 and/or Flv2/Flv4 (Helman et al.,2003; Allahverdiyeva et al., 2013, 2015; Mustila et al.,2016; Santana-Sanchez et al., 2019).In Synechocystis, the water-soluble Cyt c6 (formerly

referred to as Cyt c553) can substitute for Pc underconditions of copper deprivation (Durán et al., 2004).Cyt c6 belongs to the Cyt c family, whose members arecharacterized by a covalently bound c-type heme co-factor. c-type Cyts are further classified into groupssuch as the Cyt c6-like proteins, Cyt c555, Cyt c550, andCytM (Bialek et al., 2008). Apart from the well-established role of Cyt c6 in electron transfer (Kerfeldand Krogmann, 1998) and the role of Cyt c550 (PsbV) instabilizing the PSII water splitting complex (Shen andInoue, 1993), most of the Cyt c proteins remainenigmatic.Cyt cM (CytM) is conserved in nearly every se-

quenced cyanobacterium, with the exception of theobligate symbionts Candidatus acetocyanobacterium tha-lassa and Candidatus Synechococcus spongiarum(Supplemental Fig. S1; Bialek et al., 2016). In Synecho-cystis, CytM is encoded by sll1245 (Malakhov et al.,1994). Nevertheless, its subcellular location is ambigu-ous. An early study localized CytM to the thylakoidand plasma membranes in “purified” membrane frac-tions (Bernroitner et al., 2009). However, cross con-tamination between membranes, which has been anissue in studies using similar separation techniques,was not determined (Sonoda et al., 1997; Schultze et al.,2009). In later proteomics studies, CytM has not beendetected or localized using membranes purified by

either two-phase aqueous polymer partitioning orsubcellular fractionation (Baers et al., 2019). However,the structure of the hydrophobic N terminus resemblesa signal peptide, which suggests that CytM is targetedto a membrane. Sequence similarity to the N terminuscleavage site of Synechocystis Cyt c6 suggests that the Nterminus is processed and the mature 8.3-kD protein isinserted into the lumen (Malakhov et al., 1994). How-ever, cleavage does not seem to occur in vivo, as theprotein extracted from various cyanobacterial species,including Synechocystis, Synechococcus elongatus PCC6301, and Anabaena sp. PCC 7120, was found to bearound 12 kD (Cho et al., 2000; Bernroitner et al., 2009),implying that the hydrophobic N terminus remains onthe protein and serves as a membrane anchor. Thesubcellular location of CytM, and whether it is mem-brane anchored, is therefore still unknown.It has been suggested that CytM may play a role in

respiratory or photosynthetic electron transfer (Mannaand Vermaas, 1997; Bernroitner et al., 2009). In Syn-echocystis, CytMwas shown to reduce the CuA center ofCox in vitro with efficiency similar to that of Cyt c6(Bernroitner et al., 2009). However, given the midpointpotential of CytM (1150 mV), electron transfer fromCytf (1320 mV) to CytM would be energetically uphill(Cho et al., 2000). Notably, CytM is unable to reduce PSIin vitro (Molina-Heredia et al., 2002). Thus, it is difficultto see how the protein would substitute for Cyt c6 or Pc.Importantly, CytM is not detected under photoauto-trophic conditions (Baers et al., 2019), and deletion ofthe gene does not affect net photosynthesis or darkrespiratory rates (Malakhov et al., 1994) under theseconditions. Cold, high light, and salt stress, however,induce gene expression and the stress-induced cotran-scriptional regulation between cytM (CytM), petJ (Cytc6), and petE (Pc) suggests a stress-related role in elec-tron transfer (Malakhov et al., 1999).Besides environmental stresses, CytM has been

linked to organic carbon-assimilating trophic modes. Adark-adapted variant of Leptolyngbya boryana wasfound to grow faster than thewild type in heterotrophy.Genome resequencing revealed that the fast-growingstrain harbored a disrupted cytM (Hiraide et al.,2015). In line with this, the cytM deletion mutant ofSynechocystis demonstrated a growth advantage overthe wild type under dark- and light-activated heterotro-phic conditions, and under photomixotrophic conditions(Hiraide et al., 2015). Under dark heterotrophic condi-tions, DCytM had higher dark respiration and net pho-tosynthesis. However, the physiological mechanism andthe functional role of CytM remain entirely unknown.In this study, we sought to uncover the bioenergetics

of photomixotrophically grown Synechocystis and thephysiological background behind the growth advan-tage of DCytM by characterizing their photosyntheticmachinery and the proteomic landscape. We demon-strate gradual inhibition of QA

2 reoxidation, resultingin repression of linear electron transport and CO2fixation in Synechocystis during photomixotrophicgrowth. Amutant lacking CytM circumvents inhibition

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of QA2 reoxidation during photomixotrophic growth,

enabling higher rates of net photosynthesis. In order tomeet the substrate demand for enhanced growth, themutant retains transporter proteins, cofactor biosyntheticenzymes, and slightly adjusts central carbon metabolismcompared to the photomixotrophic wild type. Althoughthe function of CytMwas previously associatedwithCox,both thylakoid respiratory terminal oxidases, Cox andCyd, were found to be dispensable for the metabolic ad-vantage conferred by deletion of CytM in photomixo-trophy. We conclude that when cells are exposed to highGlc conditions, CytM reduces the photosynthetic capacityand contributes to regulation of the redox state of theintertwined photosynthetic and respiratory electron trans-port chain in order to accommodate this newenergy source.

RESULTS

Deletion of CytM Confers a Growth Advantage on DCytMand DCox/Cyd/CytM in Photomixotrophy

In order to elucidate the physiological role ofCytM and its possible functional association with

thylakoid-localized RTOs, we studied the DCytM,DCox/Cyd, and DCox/Cyd/CytM mutants. Unmarkedmutants of Synechocystis lacking CytM were con-structed by disrupting the cytM gene (sll1245) in thewild type (Supplemental Fig. S2) and the DCox/Cydmutant (Lea-Smith et al., 2013). Strains were then pre-cultured under photoautotrophic conditions at 3% CO2and examined under a range of different growth con-ditions at air-level CO2.

First, we determined whether deletion of cytM af-fected photoautotrophic growth by culturing cells un-der moderate constant 50 mmol photons m22 s21 light.In line with previous studies (Malakhov et al., 1994;Hiraide et al., 2015), no growth difference was observedbetween DCytM and the wild type under photoauto-trophic conditions (Fig. 1A).

Next, we characterized growth under photo-mixotrophic conditions. To determine how differentstarting Glc concentrations affected photomixotrophicgrowth (Fig. 1, A and B), we supplemented themediumwith 5 mM and 10 mM Glc and cultivated the strainsunder constant 50 mmol photons m22 s21 light. Basedon optical density measurements (OD750), all cultureswith added Glc grew substantially faster than those

Figure 1. Impact of different Glc con-centrations and light regimes on thegrowth of wild type (WT), DCytM,DCox/Cyd, and DCox/Cyd/CytM. Cul-tureswere exposed to 50mmol photonsm22 s21 light (A and B) andwere grownunder photoautotrophic conditionswithout Glc (dash-dotted line) or underphotomixotrophic conditions with 5 mM

Glc (solid line) or 10 mM Glc (dashedline). Growth was then assessed undervarious light regimes in cultures con-taining 10 mM Glc (C and D), underconstant 50 mmol photons m22 s21 light(dashed line), constant 10 mmol photonsm22 s21 light (solid line), and LAHG,which included 15 min of 50 mmolphotons m22 s21 light exposure every24 h (dash-dotted line). Values aremeans 6 SD, n 5 3–7 biological repli-cates.OD750,Optical density at 750 nm.

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cultured photoautotrophically (Fig. 1, A and B). Dele-tion of cytM had no effect on cells grown at 5 mM Glc.However, after 3 d of culture with 10 mM Glc, DCytMhad an OD750 1.9 6 0.4 (P 5 6E26) times higher thanthat of the wild type and DCox/Cyd/CytM had anOD750 1.9 6 0.6 (P 5 0.002) times higher than that ofDCox/Cyd. In line with this, DCytM consumed moreGlc than did the wild type (Fig. 2A), as quantified bymeasuring the Glc concentration of the cell-free spentmedia on day 3 of photomixotrophic growth.We next characterized growth under photo-

mixotrophic conditions but with different light regimes(Fig. 1, C and D), either constant light at 10 mmol pho-tons m22 s21 (low-light photomixotrophy) or 50 mmolphotons m22 s21 light for 15 min every 24 h (light-ac-tivated heterotrophic growth [LAHG]). These cultureswere supplemented with 10 mM starting Glc. Interest-ingly, under low-light photomixotrophy, neitherDCytM nor DCox/Cyd/CytM demonstrated a growthadvantage compared to the wild type and DCox/Cyd,respectively. Under the LAHG condition, DCytM grewfaster than the wild type, as previously reported(Hiraide et al., 2015). The DCox/Cyd and DCox/Cyd/CytM mutants were unable to grow under LAHG,supporting the findings of Pils (1997), who reportedthat Cox is indispensable under this condition.We next examined the morphology of DCytM and

wild-type cells on day 3 of photomixotrophic growth(10 mM Glc, 50 mmol photons m22 s21 constant light),when the highest difference in OD750 was observed.Cell size, cell number per OD750, and chlorophyll (chl)concentration per cell were determined. No differencewas observed in cell size between DCytM and the wildtype (Supplemental Fig. S3), and the cell number perOD750 was similar in both strains (Fig. 2B), confirmingthat the difference in OD750 reflects higher growth.However, the chl a content per cell increased in DCytM(Fig. 2C), suggesting that the photosystem content, orPSII/PSI ratio, has been altered in this strain.Overall, the most pronounced growth advantage of

DCytM over the wild type was observed when cellswere exposed to a light intensity of 50 mmol photonsm22 s21 and Glc concentration of 10 mM. Therefore,these conditions were used for all subsequent pheno-typing experiments in which cells cultured photo-mixotrophically were examined. The same phenotypemanifested in the triple DCox/Cyd/CytM mutant,showing that Cox and Cyd are not required for thegrowth advantage. Moreover, we demonstrate thatdeletion of cytM leads to a higher cellular chl a content,which implies an altered photosynthetic machinerywhen cells are cultured photomixotrophically.

Deletion of CytM Circumvents Inhibition of QA2

Reoxidation under Photomixotrophy

To determine how long-term exposure to photo-mixotrophy affects the photosynthetic machinery ofSynechocystis wild type and how deletion of CytM

rescues this phenotype, we first analyzed net photo-synthesis by probing the O2 evolution capacity of cells(Fig. 3E). When wild-type cells were grown photo-mixotrophically, only marginal net photosynthetic O2evolution was observed on day 3. Strikingly, in thepresence of the artificial electron acceptor 2,6-dichloro-p-benzoquinone (DCBQ), the O2 evolving capacity ofthe photomixotrophically grown wild type increased,although not to the level of the photoautotrophicallycultured wild type. DCBQ accepts electrons from QAand/or QB, disconnecting PSII from the downstreamelectron transfer chain (Srivastava et al., 1995). Thissuggests that a high proportion of PSII complexes arefunctional in the photomixotrophically cultured wildtype and that inhibition of net photosynthesis is in-duced by a blockage downstream of PSII. Photo-mixotrophically grown DCytM demonstrated netphotosynthetic O2 production and PSII activity similarto those of the photoautotrophically culturedwild type,

Figure 2. Glc consumption, cellular chl content, and cell number ofwild-type (WT) and DCytM cultures on day 3 of photomixotrophicgrowth. A, Amount of Glc consumed by the cells was deduced from theremaining Glc in spent media on day 3. This number reflects the con-sumption of the whole culture rather than the Glc uptake rate of a givennumber of cells. B and C, Optical density per cell number (B)and cellular chl content (C) were determined. Values aremeans6 SD forn5 3 biological replicates. Cultures were grown photomixotrophicallyunder constant 50 mmol photons m22 s21 illumination supplementedwith 10 mM Glc. Samples were taken on day 3. OD750, Optical densityat 750 nm.

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Figure 3. Fluorescence yield in wild-type (WT) and DCytM cells and quantification of O2 production capacity during photo-mixotrophic growth. A to D, Chl fluorescence of photoautotrophically (A and B) and photomixotrophically (C andD) grownwild-type andDCytMwhole cells. Photoautotrophic and photomixotrophic cultureswere grown under constant 50mmol photonsm22

s21 illumination for 3 d, with or without 10mMGlc, respectively. Prior tomeasurements, cells were resuspended in BG-11with (Cand D) andwithout (A and B) 10 mM Glc and dark adapted for 15 min. Maximum fluorescence was determined by applying a MTsaturating pulse (500ms at 5,000mmol photonsm22 s21) in darkness (black bars), under 40Wm22 far-red light (brown bars), andunder 50 mmol photons m22 s21 actinic red light (red bars). F0, initial fluorescence; Fm

D, maximum fluorescence in dark; FmFR,

maximum fluorescence in far-red light; Fm9, maximum fluorescence in actinic red light; FsFR, steady-state fluorescence in far-red

light; Fs, steady-state fluorescence in actinic red light. E, Rates of net oxygen production of photoautotrophically (PA WT) andphotomixotrophically grown wild type and DCytM were determined in cells taken on day 3 of growth. O2 production was ini-tiated with white light (1,000 mmol photons m22 s21) in the absence (control) and presence of 0.5 mM DCBQ. Rates areexpressed as micromoles O2 per milligram chlorophyll per hour, with the DCBQ-treated wild type considered as 100%. Valuesare means6 SD for n5 4 biological replicates. Asterisks indicate statistically significant differences (*P, 0.05 and **P, 0.001).F, Immunoblot analysis with D1-N antibody was performed on samples taken on day 3. Fifteen micrograms total protein extractwas loaded per 100% lane; 50% and 200% correspond to 7.5 and 30 mg, respectively. a.u., Arbitrary units.

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implying that deletion of CytM preserves photosyn-thetic activity under photomixotrophy. Immunoblot-ting performed on total protein extracts from thephotomixotrophically grown wild type and DCytMdemonstrated a higher accumulation of PSII reactioncenter protein D1 in CytM compared to the former(Fig. 3F), suggesting that PSII levels are maintained inthe mutant throughout photomixotrophic growth. Theincreased amount of D1 in DCytM likely contributes tothe higher O2 production in this mutant compared tothe wild type, although it is unlikely to entirely accountfor the difference.Next, we assessed photosynthetic activity by probing

chl fluorescence in wild-type and DCytM whole cellswith multiple-turnover (MT) saturating pulses in thedark and under far-red and actinic red light (Fig. 3,A–D). Compared to DCytM cells cultured photoauto-trophically (Supplemental Fig. S4A), wild-type cellsgrown photomixotrophically demonstrated substan-tially higher initial fluorescence (F0) and slower relax-ation of pulse-induced fluorescence in the dark (see FmD

relaxation in Fig. 3C), which suggests that the PQ poolis highly reduced. To verify this, cells were exposed tofar-red light, which preferentially excites PSI, resultingin oxidation of the PQ pool. If the PQ pool is highlyreduced, one would expect a lower steady-state fluo-rescence level (Fs) upon illumination of the cells withfar-red light, similar to that observed in the DCox/Cydmutant (Ermakova et al., 2016). Interestingly, the op-posite effect, a considerable increase in FsFR, was ob-served (Fig. 3C). This increase suggests that inhibitionof electron transport occurs at QB, since the negligibleactinic effect of far-red light is sufficient to reduce QA,resulting in increased fluorescence. Indeed, a similarrise in fluorescence was observed in photoautotrophi-cally cultured wild-type cells measured in the presenceof 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU;Supplemental Fig. S4C), a chemical occupying the QBsite, thus blocking QA-to-QB forward electron transferin PSII.Moreover, the Fs level under steady-state actinic light

was considerably higher compared to cells grownphotoautotrophically, and firing saturating pulsesbarely increased fluorescence (see Fm9 in Fig. 3C), im-plying a highly reduced QA and negligible effective PSIIyield [Y(II); Supplemental Fig. S5A]. Similar results wereobserved in a different wild-type Synechocystis substraincommonly used in our laboratory (Supplemental Fig. S6)and in cells exposed to longer periods of illumination(Supplemental Fig. S7A). Taken together, these resultssuggest a limited capacity to oxidize the PSII acceptorsite, i.e. QA, in photomixotrophically cultured wild-typecells under illumination.

Figure 4. Relaxation of flash-induced fluorescence yield in cells ex-posed to darkness. Subsequent relaxation of fluorescence yields in thedark was measured after a single-turnover saturating pulse in photo-mixotrophically cultured cells taken on days 1 (A), 2 (B), and 3 (C) of

cultivation. Growth conditions are described in Figure 3. Prior tomeasurements, the cell suspension was adjusted to 5 mg chl mL21,resuspended in BG-11 supplemented with 10 mM Glc, and darkadapted for 5 min. WT, Wild type.

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Compared to the photomixotrophically grown wildtype, DCytM cultured under the same conditionsdemonstrated 24.8% 6 8.3% lower F0 and the pulse-induced fluorescence relaxation in darkness wasmarkedly faster (see FmD in Fig. 3D). Far-red illumina-tion did not increase fluorescence, while saturatingpulses greatly increased it (see Fm9 in Fig. 3D), sug-gesting that the PSII effective yield, Y(II), remainedsignificantly higher, unlike in photomixotrophicallygrown wild-type cells (Supplemental Fig. S5A). Thus,in sharp contrast to the wild type, DCytM preserved awell-oxidized electron transport chain under photo-mixotrophy. Similarly, the triple mutant DCox/Cyd/CytM demonstrated high Y(II) compared to DCox/Cydunder photomixotrophy (Supplemental Figs. S7, C andD, and S8, C and D).

To determine how the wild type builds up a highlyreduced QA over 3 d of photomixotrophic growth, wemonitored the redox kinetics of the PSII primary elec-tron acceptor QA (Fig. 4) by firing a single-turnoversaturating flash on dark-adapted cells. Relaxation ofthe chl fluorescence yield was then recorded in the pe-riod of subsequent darkness. No difference was ob-served between wild-type and DCytM cells culturedphotoautotrophically (Supplemental Fig. S9A), and onthe first day of photomixotrophy, both wild-type andDCytM cells demonstrated typical flash-fluorescencerelaxation in the darkness. On day 2, wild-type cellsdemonstrated a substantial slow-down in QA

2 reox-idation, reflected by slow decay kinetics (Fig. 4B), whileon day 3, there was a nearly complete loss of QA-to-QBelectron transfer (Fig. 4C).

Interestingly, the kinetics from day 3 resembled acurve recorded in the photoautotrophically cul-tured wild type supplemented with DCMU prior tomeasurement (Supplemental Fig. S9A). This sup-ports the conclusion that QA-to-QB electron transferwas strongly inhibited in the majority of PSIIcenters in the wild type on the third day of photo-mixotrophy. Preillumination of the cells with far-red light did not accelerate QA

2 reoxidation(Supplemental Fig. S9B), thus supporting the ideathat the inhibition is not simply due to a highly re-duced PQ pool, although overreduction of the PQpool cannot be excluded.

DCox/Cyd and DCox/Cyd/CytM displayed pro-nounced waving in the fluorescence yield relaxationkinetics (Fig. 4). The wave phenomenon is an unusualpattern in the decay of flash-induced chl fluorescenceyield in the dark. The feature is characterized by a dip,corresponding to transient oxidation of QA

2, and asubsequent rise, reflecting rereduction of the PQ poolbyNDH-1 (Deák et al., 2014). During growth over the 3-d period, the wave phenomenon in DCox/Cyd becameless evident due to gradual inhibition of QA-to-QB elec-tron transfer. In contrast, DCox/Cyd/CytM displayedprominent waving during all 3 d of photomixotrophicgrowth, demonstrating that QA

2 reoxidation was sus-tained. Slight waving in DCox/Cyd under photoauto-trophic conditions was reported previously (Ermakova

et al., 2016), and here we demonstrate that Glc induces astrong wave phenomenon.

In order to evaluate electron transfer through Cyt b6f,the redox kinetics of Cyt f were examined (Fig. 5). Boththe photoautotrophically grown wild type (Fig. 5A)and DCytM (Supplemental Fig. S10) demonstrated fastoxidation of Cyt f followed by its reduction and reox-idation, exhibiting wavelike kinetics upon dark-to-lighttransition. In the subsequent dark, rapid reduction ofCyt f was observed. When DCMU was added to thewild type prior to the measurement (Fig. 5A), illumi-nation initiated steady oxidation, but the transientrereduction was eliminated and the subsequent re-duction in dark was slower. The photomixotrophicallygrown wild type (Fig. 5B) demonstrated trends simi-lar to DCMU-treated wild-type cells grown underphotoautotrophic conditions, confirming that electrontransfer from PSII to Cyt b6f is inhibited. In contrast,DCytM grown photomixotrophically (Fig. 5B) resembled

Figure 5. Redox kinetics of Cyt f in wild-type (WT) and DCytM cells.Cells were grown for 3 d under photoautotrophic (A) and photo-mixotrophic (B) conditions, as described in Figure 3. Oxidation of Cyt fwas induced by 500 mmol photons m22 s21 green light. When indi-cated, 20 mM DCMU was added prior to the measurement. The curveswere normalized to their respective maximal oxidation. The kinetics arerepresentative of three biological replicates.

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untreatedwild-type cells subjected to photoautotrophicconditions.These results demonstrate that during photo-

mixotrophic growth, the electron flow at the PSII ac-ceptor site gradually becomes inhibited in the wildtype, leading to drastically slower electron transferfrom PSII to Cyt b6f on day 3. Deletion of CytM cir-cumvents this inhibition and maintains PSII reactioncenter protein D1 amounts and a steady electron fluxfrom PSII to Cyt f.

DCytM Has a Larger Pool of Oxidizable PSI Than the WildType under Photomixotrophy

Next, we determined activity of PSI by monitoringthe redox kinetics of P700, the primary electron donorof PSI (Fig. 6), which was performed simultaneouslywith chl fluorescence measurements (Fig. 3). First, themaximal amount of oxidizable P700, Pm, was deter-mined (Fig. 6A). Compared to cells cultured underphotoautotrophic conditions, wild-type cells grownphotomixotrophically had 45.2% 6 0.03% lower Pm.However, the difference between DCytM cultured

under photomixotrophic and photoautotrophic condi-tions was negligible (17.2% 6 19.3%). Thus, underphotomixotrophic conditions, the maximum amountsof oxidizable P700were 132%6 18.7% higher inDCytMthan in the wild type (Fig. 6A). In line with this, im-munoblotting revealed higher levels of PSI reactioncenter subunit PsaB in DCytM compared to the wildtype under photomixotrophic growth (Fig. 6B). To de-termine the PSI:PSII ratio, samples were analyzed at77K by measuring chl fluorescence emission. No sta-tistical difference was observed between the wild typeand DCytM (Supplemental Fig. S11), demonstratingthat the PSII:PSI ratio was similar in both strains.The PSI effective yield, Y(I), was also quantified,

and was three times lower in photomixotrophicallycultured wild-type cells compared to those grownphotoautotrophically (Supplemental Fig. S5B). Thisis due to a strong donor side limitation of PSI [Y(ND);Supplemental Fig. S5C], which demonstrates an electronshortage to P7001. In contrast, photomixotrophicallycultured DCytM demonstrated similar Y(I) and onlyslightly increased Y(ND) compared to photoautotro-phically cultured wild type and DCytM (SupplementalFig. S5, B and C). As a result, Y(I) was more than three

Figure 6. Characterization of PSI in cellscultured photomixotrophically. The max-imal amount of oxidizable P700, Pm (A),and immunoblotting of the PSI reactioncenter protein PsaB (B) were determinedin cells cultured photomixotrophically.Values aremeans6 SD for n5 3 biologicalreplicates. C to F, Slow (C and D) and fast(E and F) kinetics were measured in par-allel with fluorescence (Fig. 3). Fast ki-netics curves are normalized to Pm andreferenced against their respective mini-mum P700 signal detected after the pulse.Cultivation, sample preparation, and ex-perimental parameters are similar to thosedetailed in Figure 3. P0, Initial P700; Pm

D,maximum P700 in darkness; Pm, maxi-mum P700 under far-red light; Pm9, maxi-mum P700 under red actinic light; WT,wild type.

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times higher in DCytM than in the wild type underphotomixotrophy (Supplemental Fig. S5B).

Next, pulse-induced P700 fast kinetics were com-pared between the photoautotrophically and photo-mixotrophically cultured wild type (Fig. 6E) andDCytM (Fig. 6F). These fast kinetics reveal the dynamicsof P700 oxidoreduction during saturating pulses on themillisecond scale. Saturating pulses are flashed indarkness (Pm

D) under far-red light (Pm) and actinic redlight (Pm9). Typically, the photoautotrophically cul-turedwild type (Supplemental Fig. S12A) demonstratestransient P7001 rereduction during light pulses. How-ever, the photomixotrophically grown wild type didnot exhibit the typical transient rereduction (Fig. 6E).Importantly, P7001 relaxation after the pulse (Fig. 6E)was markedly slower compared to that observed inphotoautotrophically cultured cells (Supplemental Fig.S12A). Collectively, these results confirm that fewer elec-tronswere transferred toP7001, leading to a higherY(ND)in the photomixotrophically grown wild type. Photo-mixotrophically cultured DCytM (Fig. 6F) displayedtransient rereduction during the pulses (see PmD, PmFR andPm9 in Fig. 6F) and rapid relaxation after the pulse(Fig. 6F), resembling the photoautotrophically culturedDCytM and wild type (Supplemental Fig. S12, A and B).

Here, we have shown that the effective yield of PSI inphotomixotrophically cultured wild-type cells wasconsiderably lower compared to photoautotrophicallycultured cells due to an electron shortage at P7001. Thisphenotype is eliminated by deleting cytM, as increasedY(I) and higher amounts of oxidizable P700 (Pm) andPsaB were observed in DCytM compared to the wildtype on day 3 of photomixotrophy.

DCytM and DCox/Cyd/CytM Sustain Efficient NetPhotosynthesis and CO2 Fixation under Photomixotrophy

To analyze real-time gas exchange in the photo-mixotrophically grown wild type, DCytM, DCox/Cyd,and DCox/Cyd/CytM (Fig. 7), whole-cell fluxes of O2and CO2 were simultaneously monitored using mem-brane inlet mass spectrometry (MIMS). In contrast to aclassical oxygen electrode, which only determines netO2 changes, MIMS, via enrichment of the samples withthe stable 18O2 isotopologue, makes it possible to simul-taneously measure the rates of gross 16O2 production byPSII and 18O2 consumption mediated by FDPs (Flv1–Flv4) and RTOs (Ermakova et al., 2016; Santana-Sanchezet al., 2019). Net O2 fluxes were calculated by finding thedifference between gross rates of 16O2 production and18O2 consumption. Furthermore, light-induced O2 con-sumption was calculated by subtracting the rate of 18O2consumption in the dark from that in the light.

Although Rubisco fixes CO2, and the instrumentcan only measure the concentration of CO2 in a sample,cells consume both CO2 and HCO3

2 from the medium.The pH-dependent equilibriumbetweenCO2 andHCO3

2

makes it possible to calibrate the CO2 concentrationmeasured with MIMS to the total inorganic carbon (Ci)

concentration in the sample. Based on the assumptionthat during steady-state photosynthesis the consumptionof total Ci is a function of Rubisco activity (Badger et al.,1994; Sültemeyer et al., 1995), the total Ci flux representsCO2 consumption rates.

In the wild type under 200 mmol photons m22 s21-white light, O2 consumption and gross production rateswere similar, resulting in nearly zero net photosyn-thetic O2 production. This is in line with the dataobtained by the O2 electrode (Fig. 3E). Correspondingto the minor net photosynthetic O2 production ob-served, the rate of CO2 consumption was negligible(Fig. 7A; Supplemental Fig. S13A). Importantly, nolight-induced O2 consumption was observed in thewild type (Fig. 7, A and C), although a substantialamount of Flv3 was detected by immunoblotting(Fig. 7B). While the thylakoid-localized RTOs, Cox andCyd, were shown to be active in light (Ermakova et al.,2016), a slight inhibition of respiratory O2 consumptionoccurred in the wild type under 200 mmol photons m22

s21 illumination. In contrast, DCytM exhibited a posi-tive net O2 production rate and active CO2 consump-tion (Fig. 7, A and E; Supplemental Fig. S13C).Strikingly, gross O2 production was ;10 times higherthan in the wild type, and 18O2 consumption in lightfollowed a triphasic pattern, a characteristic trendreflecting the contribution of Flv1/Flv3 and Flv2/Flv4 toO2 consumption in light (Santana-Sanchez et al., 2019).The triphasic pattern in DCytMwas observed as an initialburst of O2 consumption following the dark-to-lighttransition, which faded after 1 to 1.5min and continued ata relatively constant rate (Fig. 7E). Accordingly, immu-noblotting confirmed higher accumulation of the Flv3proteins in DCytM. The rate of light-induced O2 con-sumption in DCytM is comparable to the reported valuesof the photoautotrophically grown wild type (Huokkoet al., 2017; Santana-Sanchez et al., 2019). The dark res-piration rate was slightly higher in DCytM compared tothe wild type, as previously observed in DCytM culturedunder dark, heterotrophic conditions (Hiraide et al., 2015).

Similar to the wild type, DCox/Cyd (Fig. 7, A and D)showed minimal photosynthetic activity on day 3 ofphotomixotrophic growth. During illumination, net O2production remained negative, and CO2 consumptionwas found to be negligible (Fig. 7A; Supplemental Fig.S13B). Only residual gross O2 production was ob-served, and O2 consumption was not stimulated bylight (Fig. 7, A and D). Flv3 protein abundance inDCox/Cyd was comparable to that in the wild type(Fig. 7B). In sharp contrast to DCox/Cyd, DCox/Cyd/CytM demonstrated high PSII activity and a net O2production rate similar to that of DCytM (Fig. 7, A andF). DCox/Cyd/CytM displayed a triphasic O2 con-sumption pattern under illumination (Fig. 7F), and thelight-induced O2 consumption was comparable to thatof DCytM in steady state (Fig. 7E). Compared to DCox/Cyd, DCox/Cyd/CytM had higher levels of Flv3(Fig. 7B). Notably, deleting cytM in the DCox/Cydmutant did not enhance dark respiration, whereasDCytM had higher rates compared to the wild type.

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To conclude, mutants lacking CytM sustained asteady electron flux toward O2 and CO2 under photo-mixotrophy, reflected by substantial net O2 productionand active CO2 consumption during illumination.

Photomixotrophically Cultured DCytM Cells AccumulateTransport Proteins and Cofactor Biosynthetic Enzymes

In order to understand the metabolism of photo-mixotrophically grown wild type and DCytM, weanalyzed the total proteome by nanoscale liquid

chromatography electrospray ionization tandem MSvia the data-dependent acquisition method. Samples foranalysis were collected on day 2, when both wild-typeandDCytMcellswere in the late exponential phase and asubstantial growth difference was observed between thestrains (Fig. 8A).In total, 2,415 proteins were identified (Supplemental

Dataset S1), despite the fact that the dataset was slightlybiased against basic (Fig. 8D) and hydrophobic proteins(Fig. 8E), which is a known issue with this technique(Chandramouli and Qian, 2009). Out of 2,415 proteins,

Figure 7. O2 and CO2 fluxes in photo-mixotrophically cultured wild-type (WT),DCytM, DCox/Cyd, and DCox/Cyd/CytMcells. A, Rates of O2 and CO2 fluxes atsteady state. Values are means6 SD for n53 to 5 biological replicates. B, Total proteinextracts were analyzed by immunoblottingwith a-Flv3-specific antibody: 15 mg totalprotein was loaded per 100% lane; 50%and 200% correspond to 7.5mg and 30mg,respectively. C to F, Kinetics of O2 flux ratesin whole cells. Cultivation, sample prepa-ration, and experimental conditions aredetailed in Figure 3. In the light phase, 200mmol photonsm22 s21 constant white lightwas applied. Samples were supplementedby 1.5 mM NaHCO3. Kinetics are repre-sentatives of three to six biological repli-cates. The source data for Figure 7A can befound in Supplemental Table S2.

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634 were quantified, with 162 displaying a statisticallydifferent abundance in DCytM compared to the wildtype (fold change [FC]. 1.5 and FC,21.5 [P, 0.05];Supplemental Dataset S2). The functional classificationof differentially regulated proteins (Fig. 8C) revealedthat apart from unknown or hypothetical proteins,mainly transporters and biosynthetic enzymes werealtered in photomixotrophically cultured DCytM cells.

Supplemental Dataset S3 shows a selection of pro-teins whose abundance was different in DCytM com-pared to the wild type. The highest fold change wasobserved in transport proteins. Among these, the con-stitutive low-affinity ABC-type phosphate transporters(PstA1, PstB1, PstB19, and PstC), periplasmicphosphate-binding proteins (SphX and PstS1), and ex-tracellular lytic enzymes (PhoA and NucH) were moreabundant in DCytM. Among proteins related to Ci

uptake, a thylakoid b‐type carbonic anhydrase, EcaB,was 2.32 times (P 5 7.50E203) more abundant inDCytM. EcaB is a CupA/B-associated protein proposedto regulate the activity of NDH-13 (NDH-1 MS) andNDH-14 (NDH-1 MS9; Sun et al., 2018). NDH-13 facili-tates inducible CO2 uptake, whereas NDH-14 drivesconstitutive CO2 uptake (Ogawa, 1991). CupB is ex-clusively found in the NDH-14 complex and convertsCO2 into HCO3

2. Interestingly, no significant changewas observed in the level of the Glc transporter GlcP,although the growth advantage of DCytM was ob-served upon exposure to Glc.

Chl a biosynthetic enzymes were found to accumu-late in the mutant (Supplemental Dataset 3). ChlL, asubunit of the light-independent protochlorophyllidereductase (Wu and Vermaas 1995), and ChlP (4.61E203), a geranylgeranyl reductase (Shpilyov et al., 2005),

Figure 8. Characteristics at the sampling stage and functional classification of differentially regulated proteins in DCytM. A,Growth of the analyzed cultures, with the ellipsis marking the sampling day. Cells were cultured similarly to those used in thebiophysics analysis, except that the cells for proteomicswere precultivated under atmospheric CO2 in order to fully adapt the cellsto these conditions. Importantly, the extra preculturing step did not affect the growth of the experimental cultures. Values aremeans6 SD for n5 3 biological replicates. B, Relaxation of the flash-induced fluorescence yield in the dark was measured in theabsence (solid symbols) and presence (open symbols) of 20 mM DCMU. C, Differentially regulated proteins in DCytMwere grouped according to their function. In total, 2,415 proteins were identified, of which 634 proteins were quantified and 162were differentially regulated. The practical significance of differentially regulated proteins was set to FC . 1.5 and FC , 21.5(P , 0.05). D and E, Effects of pI (D) and hydrophobicity (GRAVY; E) of the proteins on the identification rate were determined.Black squares mark all of the 3,507 predicted proteins in Synechocystis; lilac circles mark each protein identified in the wild type(WT) and DCytM.

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were upregulated 9.28-fold (P 5 5.32E203) and 1.52-fold (P 5 4.61E203), respectively, in DCytM. Incorpo-ration of chl into photosystems is likely increased due tothe elevated level of Pitt, a protein contributing to theformation of photosynthetic pigments/proteins at theearly stages of biogenesis (Schottkowski et al., 2009).The ligand of the tetrapyrrole ring of chl is Mg21, andaccordingly, the magnesium uptake protein MgtE ac-cumulated in DCytM along with a periplasmic iron-binding protein, FutA2, part of the complementaryuptake system of iron, a vital element of the photo-synthetic machinery (Kranzler et al., 2014). Amongpigment biosynthetic enzymes, DCytM showed in-creased levels of the heme oxygenase Ho1, catalyzingthe final step in the production of biliverdin (Willowset al., 2000). Biliverdin is the precursor of phycocya-nobilin, which is incorporated into phycobilisomes, thelight-harvesting complexes of Synechocystis.Among the photosynthetic proteins, the PSI reaction

center subunit PsaB was found in equal amounts in thewild type and DCytM. However, immunoblotting withan anti-PsaB antibody demonstrated that DCytM con-tained higher amounts of PsaB than the wild type(Fig. 6B). This discrepancy may be due to the fact thatdespite the robustness of the MS-based data-dependentacquisition method, hydrophobic membrane proteinsare prone to misquantification. Via MS analysis, quan-tification of psbA-encoded D1 was not successful.Therefore, its abundance was only determined by im-munoblotting (Fig. 3F), which revealed higher levels ofD1 proteins in DCytM compared to the wild type. In-terestingly, and somewhat contradictorily, the numberof PSII assembly proteins encoded by the PSII assemblyproteins operon (Wegener et al., 2008) decreased in themutant. We also note that lower levels of NorB, aquinol-oxidizing nitric oxide reductase (Büsch et al.,2002), were observed in DCytM.Since the growth advantage of DCytM was observed

in the presence of Glc, alterations are expected in theabundance of the intermediary carbon metabolic en-zymes. In Synechocystis, ;100 enzymes participate inthis metabolic network. In our study, 40 were quanti-fied, and surprisingly, only a few proteins were differ-entially regulated in DCytM. One notable example isphosphofructokinase PfkA, the key regulatory enzymeof the glycolytic Embden-Meyerhof-Parnas pathway,which was 1.86 times (P 5 1.96E205) less abundant inDCytM, suggesting that carbon fluxmight be redirectedinto the Entner-Doudoroff or oxidative pentose phos-phate pathways. Phosphoglycerate kinase Pgk, whichis involved in each glycolytic pathway, was 2.06 times(P 5 1.27E205) as abundant in DCytM. Phosphoenolpyr-uvate synthetase PpsA, a protein that catalyzes the firststep of gluconeogenesis, was 2.21 times (P 5 3.10E204)less abundant in DCytM.To conclude, global proteomic analysis revealed that

photomixotrophically cultured DCytM accumulatestransporter and chl biosynthetic proteins, while slightchanges in the amount of certain glycolytic and pho-tosynthetic proteins were also observed.

DISCUSSION

The effect of importing and metabolizing organiccarbon on the bioenergetics properties of cyanobacteriaover a long-term period is not fully understood. Pre-vious studies have focused on the cellular changes fol-lowing relatively short-term (from 10 min to 24 h)exposure to organic carbon (Lee et al., 2007; Takahashiet al.. 2008; Haimovich-Dayan et al., 2011; Zilliges andDau, 2016). Themajority of these reports suggest partialinhibition of photosynthetic activity, whereas somestudies demonstrated increased net photosynthesisunder air-level CO2 after 2 h exposure to 10 mM Glc(Haimovich-Dayan et al., 2011). However, long-termchanges to bioenergetics processes, particularly photo-synthesis, remain to be elucidated. In this study, weinvestigated the effect of long-term photomixotrophicgrowth on wild-type and DCytM cells, most notably onthe photosynthetic machinery, by analyzing chl fluo-rescence, the redox kinetics of P700, real time O2 andCO2 fluxes, and changes within the proteome.

Gradually Disconnecting PSII from Cytb6f LimitsPhotosynthesis in the Photomixotrophically CulturedWild Type

By characterizing wild-type cells shifted from pho-toautotrophic to photomixotrophic conditions, weshow that photosynthesis was markedly decreasedover 3 d of cultivation. This is deduced from the lowPSII (Fig. 3C; Supplemental Fig. S5A) and PSI yields(Fig. 6C; Supplemental Fig. S5B) and,most importantly,the negligible net O2 production (Fig. 3E) and CO2 fix-ation rates (Fig. 7A; Supplemental Fig. S13A) on day 3.A residual PSII activity is ensured by circulating elec-trons in a water-water cycle. This was demonstrated byreduced PSII gross O2 production (Fig. 7, A and C),which nearly equalled O2 consumption in the light,resulting in practically zero net O2 production. Sinceaddition of an artificial PSII electron acceptor, DCBQ,largely restores O2 evolving activity (Fig. 3E), a sub-stantial portion of PSII centers are functional, butdownstream electron flux is restricted. This could bedue to a highly reduced PQ pool, which in turn affectsthe redox potential of QB and thus QA

2 reoxidation(Haimovich-Dayan et al., 2011). However, far-red light,which specifically excites PSI and drains electrons fromthe PQ pool, did not accelerate QA

2 reoxidation in pho-tomixotrophically culturedwild type (Supplemental Fig.S9B). Thus, overreduction of the PQ pool cannot be thesole reason for the restricted downstream electron flux.Interestingly, the photomixotrophically cultured wildtype resembles DCMU-treated cells in many ways: (1)far-red light illumination increases steady-state fluores-cence (Fig. 3C; Supplemental Fig. S4); (2) transient rere-duction and subsequent reoxidation of Cyt f underillumination is nearly absent, andCyt fdecay in darknessis slow (Fig. 5); (3) flash-induced decay of QA

2 after 3 dexposure to Glc highly resembles the kinetics of DCMU-treated cells (Fig. 8B) and differs from kinetics observed

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in the DBMIB-treated wild type (Supplemental Fig.S9B; Deák et al., 2014). These results suggest that pho-tosynthetic electron flow from PSII to the PQ pool andCyt b6f is hindered. However, this is not simply due to ahighly reduced PQ pool.

The gradual disconnection between PSII and Cyt b6fand the resulting decrease in photosynthesis could bedue to a spatial isolation of PSII, via rearrangement inthe thylakoid, to another location. Rearrangement ofthylakoid-localized complexes, specifically NDH-1 andSDH, has been observed in response to redox-regulatedchanges in the electron transport chain (Liu et al., 2012).Applying the same analogy to PSII, the highly reducedstate of the PQ pool might trigger the complexes toarrange into a more sparse distribution during photo-mixotrophic growth. Although cyanobacterial thyla-koids are densely packed membranes (Kaňa, 2013),lateral heterogeneity (Agarwal et al., 2010) andmobilityof PSII (Casella et al., 2017) have been demonstrated.

Photomixotrophy Does Not Alter Photosynthetic ElectronTransport in DCytM

Surprisingly, deletion of CytM reverses the down-regulation of photosynthesis in photomixotrophy,resulting in a profile similar to that of wild-type andDCytM cells grown under photoautotrophic condi-tions. Importantly, DCytM demonstrated unrestrictedelectron flow between PSII, Cyt b6f, and PSI. The rate ofgross O2 production (Fig. 7, A and E) was 10 timeshigher in DCytM cells compared to wild-type cellscultured under photomixotrophic conditions. In con-trast to the photomixotrophically cultured wild type,DCytM showed a clear wave pattern in Cyt f kineticsupon dark-to-light transition and did not demonstrateslow rereduction of Cyt f in the dark (Fig. 5B) or PSIdonor-side limitation (Supplemental Fig. S5C). Finally,the abundance of D1 (Fig. 3F), PsaB (Fig. 6B), and PetA

and PetB (Supplemental Dataset S3), the core subunitsof PSII, PSI, and Cyt b6f, respectively, was higher inDCytM than in the wild type, although the PSI:PSIIratio was unaltered (Supplemental Fig. S11). As a con-sequence, the rate of net O2 production and CO2 con-sumption (Fig. 7A) was substantially higher in DCytM,demonstrating that deletion of CytM conserves photo-synthetic activity and circumvents the inhibition of QA

2

reoxidation in photomixotrophy.The exact mechanism by which DCytM alleviates

blockage of the electron transport pathway was notelucidated in this work, nor has an exact role for thisprotein been determined in previous studies. CytM hasbeen suggested to play a role in transferring electronsfrom Cyt b6f to Flv1/Flv3, limiting productivity butproviding a possible alternative route for safely trans-ferring electrons to O2 (Hiraide et al., 2015). However,given the low midpoint potential of CytM, a large en-ergy barrier would have to be overcome in order forelectron transfer downstream of Cyt b6f to occur (Choet al., 2000). Moreover, we demonstrated that the ab-sence of CytM does not decrease O2 photoreductiondriven by FDPs in DCox/Cyd/CytM (Fig. 7, A and F),thus excluding this possibility. Recently, a cyano-bacterial ferredoxin, Fed2, was shown to play a role iniron sensing and regulation of the IsiA antenna protein,a protein that is typically expressed when cells are ex-posed to low-iron conditions (Schorsch et al., 2018).Similar to Fed2, it is possible that CytM plays a regu-latory role in the cell, rather than being directly in-volved in electron transport under photomixotrophy.

Under conditions in which cells are exposed to Glc orother sugars, CytM may regulate carbon assimilation.DCytM demonstrates substantial growth under darkheterotrophic conditions (Hiraide et al., 2015). How-ever, the majority of the known cyanobacteria cannotgrow heterotrophically, indicating that the function ofCytM extends beyond the modulation of heterotrophicgrowth (Bialek et al., 2016). Under photomixotrophic

Figure 9. Schematic showing changes in themetabolism of photomixotrophically grownDCytM cells compared to the wild type. Proteins,compounds, and metabolic routes with increasedabundance or activity in DCytM relative to thewild type are represented in green. Blue indicateslower abundance in DCytM, gray indicates nochange, and white indicates undetermined a-bundance or activity. CB, Calvin-Benson cycle; Ci

uptake, inorganic carbon uptake; Co uptake, or-ganic carbon uptake; ED, Entner-Doudoroffpathway; EMP, Embden-Meyerhof-Parnas path-way; OPP, oxidative pentose phosphate pathway;Pi uptake, inorganic phosphate uptake; PM,plasma membrane; TCA, tricarboxylic acid cycle;TM, thylakoid membrane.

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conditions, CytM likely is involved in regulation ofthylakoid rearrangements or photosynthetic electrontransport and carbon fixation, limiting CO2 uptake anddecreasing the total amount of photosynthetic proteins,which in turn reduces photosynthesis. In line with this,we observed accumulation of EcaB in DCytM (Fig. 9;Supplemental Dataset S3). Enhanced EcaB levelslikely result in greater inorganic carbon assimilation,higher carbon fixation, and increased turnover ofNADPH, the terminal electron acceptor in linearphotosynthetic electron transport. This in turn likelylimits overreduction of the photosynthetic electrontransport chain.Regardless of the exact role of CytM, it is clear that

deletion of this protein substantially increases growthof Synechocystis in photomixotrophy (Fig. 1), in linewith previous studies (Hiraide et al., 2015). This ispossibly due to an increase in photosynthetic capacitycombined with efficient assimilation of Glc into centralmetabolism, resulting in greater biomass accumulation.This resulted in increased production of proteins re-quired for enhanced growth, including those involvedin phosphate uptake (PstA1, PstB1, PstB19, and PstC;Supplemental Dataset S3), import of Mg21 (MgtE),Zn21 (ZiaA), and Fe21 (FutA2), and production of chl(ChlP and ChlL; Fig. 9; Supplemental Dataset S3).In conclusion, under long-term photomixotrophy

Synechocystis cells gradually decrease photosyntheticelectron transport by disconnecting PSII from Cyt b6f.Deletion of CytM allows Synechocystis to maintain ef-ficient photosynthesis and enhanced growth underlong-term photomixotrophy. While we have not de-termined the exact function of CytM, we propose that itplays a role in reducing photosynthesis under condi-tions when both light intensity and Glc concentrationfluctuate (Ittekkot et al., 1981; Hieronymi and Macke,2010), and the redox state of the intertwined photo-synthetic and respiratory electron transfer rapidlychanges.

MATERIALS AND METHODS

Plasmid Construction

The genome sequence of Synechocystis sp. PCC6803 released onNovember 5,2004 was consulted via Cyanobase (http://genome.kazusa.or.jp/cyanobase)for primer design. Primers are listed in Supplemental Table S1. The cytM(sll1245) gene was deleted by amplifying a 906-bp fragment upstream of cytMusing primers CytMleftfor and CytMleftrev and a 932-bp fragment down-stream of cytM using primers CytMrightfor and CytMrightrev, followed byinsertion of the respective fragments into the SacI/EcoR1 andXbaI/BamH1 sitesof pUC19 to generate pCytM-1. The BamH1-digested npt1/sacRB cassette frompUM24Cm (Ried and Collmer, 1987) was inserted into the BamH1 site betweenthe upstream and downstream fragments in pCytM-1 to generate pCytM-2.

Construction of cytM Deletion Mutants

Unmarked mutants of Synechocystis lacking cytM were constructed via atwo-step homologous recombination protocol according to Lea-Smith et al.(2016b). To generate marked mutants, ;1 mg of plasmid pCytM-2 was mixedwith Synechocystis cells for 6 h in liquidmedia, followed by incubation on BG-11agar plates for ;24 h. An additional 3 mL of agar containing kanamycin was

added to the surface of the plate, followed by further incubation for ;1 to 2weeks. Transformants were subcultured to allow segregation of mutant alleles.Segregation was confirmed by PCR using primers CytMf and CytMr, whichflank the deleted region. To remove the npt1/sacRB cassette to generate un-marked mutants, mutant lines were transformed with 1 mg of the markerlessCytM-1 construct. Following incubation in BG-11 liquid media for 4 d and agarplates containing Suc for a further 1 to 2 weeks, transformants were patched onkanamycin and Suc plates. Suc resistant, kanamycin sensitive strains containingthe unmarked deletion were confirmed by PCR using primers flanking thedeleted region (Supplemental Fig. S2B). The ΔCox/Cyd/CytM unmarkedstrain was generated via the same method in the background of the unmarkedΔCox/Cyd strain (Lea-Smith et al., 2013).

Cultivation

Cells kept in cryogenic storagewere revived onBG-11 agar plates at 3% (v/v)CO2. Pre-experimental cultures were inoculated at 0.1 OD750 by transferring apatch of cells from plates into 30 mL BG-11 medium buffered with 10 mM TES-KOH (pH 8.2) in 100-mL Erlenmeyer flasks. Cultures were shaken at 120 rpm at30°C and exposed to constant white fluorescent light of 50 mmol photons m22

s21 intensity in a Sanyo Environmental Test Chamber supplemented with 3%(v/v) CO2. Pre-experimental cultures were cultivated for 3 d, with densitytypically reaching 2.5 6 0.5 OD750.

Experimental cultures for growth and photophysiological experiments wereinoculated in 30 mL fresh BG-11 media at 0.1 OD750 from harvested pre-experimental cultures. The media was buffered with 10 mM TES-KOH (pH8.2), the CO2 concentration was atmospheric, and cultures were agitated in 100-mL Erlenmeyer flasks at 120 rpm in AlgaeTRON AG130 cool-white LEDchambers (PSI Instruments). Growthwas tested under constant light of 50mmolphotons m22 with different initial Glc concentrations: (1) no Glc; (2) 5 mM Glc;and (3) 10 mM Glc. At 10 mM Glc, additional light regimes were tested: (4)10 mmol photons m22 s21 light and (5) 15 min 50 mmol photons m22 s21 every24 h. For photophysiological studies, cells were cultivated under condition 3 for3 d. For proteomics analysis, cells were cultivated similarly to those in condition3, with the exception of an extra 3-d-long precultivation step at atmosphericCO2 without Glc.

Cell Counting, Cell Size Determination

Cell number was determined with a Nexcelom Cellometer X2 via the fol-lowing method. Sample OD750 was adjusted to 1, brightfield images werecaptured, and the cell number was determined by the Nexcelom software. Inorder to exclude the visual glitches falsely recognized as cells by the software,only the four most populous cell-size groups were averaged. Typically, 3,000cells were counted per plate.

Glc Determination

Glc concentrationof the spentmediawasdetermined spectrophotometricallywith the commercial High Sensitivity Glc Assay Kit (Sigma-Aldrich). Prior tomeasurements, the cell suspension was centrifuged at 5,000g for 10 min and thesupernatant was filtered through a 0.2-mm filter.

MIMS Measurements

Gas fluxes of intact cells were measured using MIMS. The in-house-builtsystem consists of a DW-1 oxygen electrode chamber (Hansatech) connected tothe vacuum line of a MS (Prima PRO model, Thermo Scientific). The samplecuvette was separated from the vacuum line by a Hansatech S4 polytetra-fluoroethylenemembrane. Sampleswere pelleted and resuspended in fresh BG-11 supplemented by 10 mM Glc and buffered to pH 8.2 with 10 mM TES-KOH.Chl a concentration was adjusted to 10 mg mL21. Prior to measurements, thesample was enriched with 98% (v/v) 18O2 heavy isotope (CK Isotopes), thedissolved total inorganic carbon concentration was adjusted to 1.5 mM byadding NaHCO3, and 10 to 15 min dark adaptation was applied. The mea-surement was performed in a semiclosed cuvette at 30°C with constant stirring.The light source was a 150Watt, 21 V, EKE quartz halogen-powered fiber-opticilluminator (Fiber-Lite DC-950, Dolan-Jenner). A two-point calibration wasused to calibrate the O2 signal in milli-Q water. Total inorganic carbon wascalibrated by injecting known HCO3

2 samples into a known volume of growthmedia buffered to pH 8.2 with 10 mM TES-KOH. A mathematical offset

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accounted for the changing concentration of the 18O2 and 16O2 isotopologuesover the course of an experiment to enable the accurate determination of rates(Hoch and Kok, 1963). Rates were calculated as described previously(Beckmann et al., 2009).

Clark-Type Electrode Measurements

Net O2 production of intact cells was tested in the presence of 0.5 mM DCBQat 30°C with a Clark-type oxygen electrode and chamber (Hansatech). Prior tothe measurements, cells were resuspended in BG-11 (pH 8.2) supplementedwith 10 mM Glc, the chl a concentration was adjusted to 7.5 mg mL21, then thesamples were dark adapted for 1 to 2 min. O2 production was initiated by 1,000mmol photons m22 s21 white light using a Fiber-Lite DC-950 light source. Ratesof oxygen production were calculated using the Hansatech software.

Chl Fluorescence and P700 Oxidoreduction Measurements

Whole-cell chl fluorescence was measured simultaneously with P700 with apulse amplitude-modulated fluorometer (Dual-PAM-100). Prior to measure-ments, cells were resuspended in BG-11 (pH 8.2) supplementedwith 10mMGlc,and the chl a concentration was adjusted to 15 mg mL21. Measurements wereperformed at 30°C, and samples were initially incubated in darkness for 15 minwith stirring. To determine Pm, 30 s strong far-red light (720 nm, 40Wm22) andred MT saturating pulses were applied. MT pulses were set to an intensity of5,000 mmol photons m22 s21 (width 500 ms). Red (635 nm) actinic light of50 mmol photons m22 s21 intensity was used as background illumination.Photosynthetic parameters were calculated as described previously(Klughammer et al., 2008a, 2008b).

Relaxation of flash-induced fluorescence yield was monitored using a flu-orometer (FL3500, PSI Instruments), as outlined previously (Allahverdiyevaet al., 2013). Prior to the measurement, cells were resuspended in BG-11 (pH8.2) supplemented with 10 mM Glc, adjusted to 5 mg chl a mL21 and darkadapted for 5 min. Curves were referenced against F0 and normalized to Fm.

Measurement of Cytochrome f Redox Kinetics

Cyt f redox kinetics were determined in intact cells by deconvoluting ab-sorbance changes at 546, 554, 563, and 573 nm that were measured using a JTS-10 pump probe spectrophotometer (BioLogic) and appropriate 10 nm FWHMinterference filters. BG39 filters (Schott) were used to shield the light detectorsfrom scattered light. Deconvolution was performed with the JTS-10 software.Prior to the experiments, cells were harvested and chl a concentration wasadjusted to 5 mg mL21 by resuspension in fresh BG-11 with or without 10 mM

Glc. Cells were dark adapted for 2 min prior to measurements with each in-terference filter, and then illuminatedwith green light of 500mmol photonsm22

s21 intensity for 5 s. Flashes of white detection light were administered during200-ms dark intervals in actinic illumination. When appropriate, 20 mM DCMUwas added to the samples before dark adaptation.

Western Blotting

Total protein extraction, electrophoresis, and immunoblotting were per-formed as described previously (Huokko et al., 2019). Antibodies raised againstPsaB (AS10 695, Agrisera), D1 (AS11 1786, Agrisera), and Flv3 (Antiprot) wereused in this study.

MS Analysis: Sample Preparation, Data-DependentAnalysis, Protein Identification and Quantitation

For data analysis, we used the proteome of Synechocystis sp. 6803 substrainKazusa sequenced in 2004. Protein annotation was downloaded from Uniprotand Cyanobase. Hydrophobicity was determined via the grand average ofhydropathy (GRAVY) index at www.gravy-calculator.de, and pI was calcu-lated via https://web.expasy.org/compute_pi/.

Sample preparation for MS, data-dependent analysis, and protein identifi-cation was performed as detailed previously (Huokko et al., 2019). The MSproteomics data were deposited to the ProteomeXchange Consortium via thePRIDE partner repository with the dataset identifier PXD015246 and 10.6019/PXD015246 (Perez-Riverol et al., 2019).

Statistical Analysis

P-values were calculated by one-way ANOVA and differences in the datawere considered statistically significant at P , 0.05.

Accession Numbers

Gene/protein names and accession numbers of all genes/proteins identifiedin this study are listed in Supplemental Dataset S1. The MS proteomics datawere deposited to the ProteomeXchange Consortium via the PRIDE partnerrepository with the dataset identifier PXD015246 and 10.6019/PXD015246(Perez-Riverol et al., 2019).

Supplemental Data

The following supplemental materials are available.

Supplemental Figure S1. Alignment of CytM from sequenced cyanobac-terial species.

Supplemental Figure S2. Generation of cytM deletion mutants in Synechocystis.

Supplemental Figure S3. Cell size of wild type and DCytM grown photo-mixotrophically and photoautotrophically.

Supplemental Figure S4. Fluorescence transients of photoautotrophicallycultivated wild type and DCytM determined in the presence of2 mM DCMU.

Supplemental Figure S5. Photosynthetic parameters of wild type andDCytM grown photomixotrophically and photoautotrophically.

Supplemental Figure S6. Fluorescence transients and P700 oxidoreductionon day 3 of photomixotrophic growth of the wild-type Synechocystissubstrain.

Supplemental Figure S7. Fluorescence transients and P700 oxidoreductionkinetics of photomixotrophically grown wild type, DCytM, DCox/Cyd,and DCox/Cyd/CytM.

Supplemental Figure S8. Fluorescence transients and P700 oxidoreduxtionkinetics of photoautotrophically grown wild type, DCytM, DCox/Cyd,and DCox/Cyd/CytM.

Supplemental Figure S9. Flash-induced increase of fluorescence yield andits relaxation in dark in photoautotrophically grown wild type andDCytM.

Supplemental Figure S10. Redox kinetics of Cyt f in photoautotrophicallygrown DCytM cells.

Supplemental Figure S11. 77K steady-state fluorescence emission spectraof wild type and DCytM grown photomixotrophically.

Supplemental Figure S12. Fast kinetics of P700 oxidoreduction of wildtype and DCytM grown under photoautotrophic conditions.

Supplemental Figure S13. The rate of CO2 fluxes in photomixotrophicallygrown wild type, DCytM, DCox/Cyd, and DCox/Cyd/CytM.

Supplemental Table S1. List of oligonucleotides used in this study.

Supplemental Table S2. Rates of O2 and CO2 fluxes in photomixotroph-ically grown wild type, DCytM, DCox/Cyd, and DCox/Cyd/CytM.

Supplemental Dataset S1. Proteins identified by data-dependent analysisin photomixotrophically grown wild type and DCytM.

Supplemental Dataset S2. Differentially expressed proteins in photomix-otrophically grown DCytM versus the wild type.

Supplemental Dataset S3. A selection of differentially expressed pro-teins in photomixotrophically cultured DCytM compared to thewild type.

ACKNOWLEDGMENTS

We thank Steffen Grebe for helping with Dual-PAM measurements andVille Käpylä for laboratory assistance. MS analysis was performed at the Turku

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Proteomics Facility hosted by the University of Turku and Åbo Akademi Uni-versity, supported by Biocenter Finland.

Received March 6, 2020; accepted March 23, 2020; published April 21, 2020.

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