Plastoquinone redox status influences carboxysome integrity via a RpaA- and ROS-dependent regulatory network

Plastoquinone redox status influences carboxysome integrity via a RpaA- and ROS-dependent regulatory network | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Plastoquinone redox status influences carboxysome integrity via a RpaA- and ROS-dependent regulatory network View ORCID Profile María Santos-Merino , Lauri Nikkanen , Emmanuel J. Kokarakis , Yagut Allahverdiyeva , Daniel C. Ducat doi: https://doi.org/10.1101/2025.01.24.634715 María Santos-Merino 1 MSU-DOE Plant Research Laboratory, Michigan State University , East Lansing, MI, United States, 48824 Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for María Santos-Merino Lauri Nikkanen 2 Laboratory of Molecular Plant Biology, Department of Life Technologies, University of Turku , Turku, Finland , 20014 Find this author on Google Scholar Find this author on PubMed Search for this author on this site Emmanuel J. Kokarakis 1 MSU-DOE Plant Research Laboratory, Michigan State University , East Lansing, MI, United States, 48824 3 Department of Microbiology and Molecular Genetics, Michigan State University , East Lansing, MI, United States, 48824 Find this author on Google Scholar Find this author on PubMed Search for this author on this site Yagut Allahverdiyeva 2 Laboratory of Molecular Plant Biology, Department of Life Technologies, University of Turku , Turku, Finland , 20014 Find this author on Google Scholar Find this author on PubMed Search for this author on this site Daniel C. Ducat 1 MSU-DOE Plant Research Laboratory, Michigan State University , East Lansing, MI, United States, 48824 4 Department of Biochemistry and Molecular Biology, Michigan State University , East Lansing, MI, United States, 48824 Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: ducatdan{at}msu.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract Carboxysomes are bacterial microcompartments that encapsulate Rubisco and are a core component of the cyanobacterial carbon concentration mechanism (CCM). While carboxysome number, size and spatial organization are observed to vary in different environmental conditions (CO 2 , light, temperature, light quality), molecular mechanisms underlying this potentially adaptive process remain elusive. Herein, we observed that mutants of the circadian rhythm/metabolism factor, Regulator of Phycobilisome Associated A (RpaA), exhibit a striking breakdown of carboxysomes under certain environmental conditions. We find that growth conditions leading to overreduction of the plastoquinone (PQ) pool (mixotrophic growth, high irradiance, or chemical inhibition of electron transfer from PQ to the cytochrome b 6 f complex) are accompanied by elevated generation of reactive oxygen species (ROS), and correlate with carboxysome breakdown. Carboxysome breakdown is reversed by environmental conditions or chemical inhibitors that prevent PQ overreduction and accompanying ROS generation. Taken together, our data supports a novel link between the redox status of the PQ pool and carboxysome status and/or integrity. Our results have implications for fundamental understanding of cyanobacterial energy balancing pathways and may indicate new research directions for understanding how the carboxysome is remodeled in response to changing environments. 1. Introduction Cyanobacteria and other photosynthetic organisms employ a variety of adaptive pathways to contend with variability in light, which fluctuates in predictable ( e.g. , diel cycles, seasonal variations) and stochastic ( e.g. , environmental shading, light flecks) patterns ( Labiosa et al., 2006 ). High irradiance or spikes in illumination can result in an over-reduced photosynthetic electron transport chain and lead to the generation of toxic reactive oxygen species (ROS) ( Foyer, 2018 ; Calzadilla and Kirilovsky, 2020 ). Photosynthetic organisms therefore possess a variety of regulatory mechanisms to coordinate the upstream processes of photochemical activity with downstream metabolic processes energetic demands to achieve a balance between light harvesting and energy utilization. In photosynthetic linear electron flow, electrons extracted from water are transferred from photosystem II (PSII) to photosystem I (PSI) via a series of electron carriers, including the membrane soluble plastoquinone (PQ) pool. While the ratio of proportional increase in ATP/NADPH requirements can create an energy imbalance, leading to the reduction or oxidation of the intersystem electron transport chain, and more specifically, the plastoquinone (PQ) pool ( Fujita et al., 1987 ; Sunil et al., 2013 ; Fan et al., 2021 ). State transitions ( Calzadilla and Kirilovsky, 2020 ) and photoprotective systems (such as orange carotenoid protein or flavodiiron proteins) ( Kirilovsky and Kerfeld, 2016 ; Nikkanen et al., 2021 ) are well-studied processes by which cyanobacteria attempt to rebalance the redox status of the pETC in the short term. While the formation of ROS is typically discussed as a dangerous byproduct of unbalanced photosynthetic activity, there are also well-established signaling roles for some of these molecules, especially hydrogen peroxide (H 2 O 2 ) ( Mironov et al., 2019 ). In addition to short-term changes in light availability, day/night cycles introduce irradiance changes that are anticipated by the well-characterized cyanobacterial circadian rhythm machinery ( Dong et al., 2010 ; Yang et al., 2010 ; Pattanayak et al., 2014 ; Cohen and Golden, 2015 ; Diamond et al., 2015 ; Martins et al., 2018 ; Taton et al., 2020 ). The cyanobacterial circadian clock consists of a core oscillator, KaiC, which undergoes cyclic rounds of phosphorylation and dephosphorylation that are programmed by upstream regulators connected to light reactions via the PQ pool and has output functions controlled by the two-component system proteins SasA and RpaA ( Nakajima et al., 2005 ; Ivleva et al., 2006 ; Takai et al., 2006 ; Kim et al., 2012 ; Kim et al., 2020 ). RpaA is a transcription factor that acts as a master regulator of the circadian clock outputs, by driving global rhythms of gene expression and gating of cell division ( Dong et al., 2010 ; Markson et al., 2013 ). RpaA was first discovered in Synechocystis sp. PCC 6803, as an OmpR-type response regulator that regulates the energy transfer from phycobilisomes to PSI and PSII ( Ashby and Mullineaux, 1999 ). Recently, other functions have been subscribed to RpaA beyond its roles in entraining the circadian clock and controlling clock output ( Diamond et al., 2015 ; Iijima et al., 2015 ; Diamond et al., 2017 ; Puszynska and O’Shea, 2017 ). For example, it has been proposed that RpaA may be more directly involved in regulating enzymes of core metabolism to influence carbon partitioning ( i.e. , towards glycogen storage vs. downstream metabolism) ( Puszynska and O’Shea, 2017 ). The carboxysome is an essential complex of the cyanobacterial carbon concentration mechanism (CCM) and is also dynamically regulated in response to changing light and other environmental conditions ( Burnap et al., 2013 ). Carboxysomes are bacterial microcompartments comprised of a proteinaceous shell that encapsulates Rubisco. The primary function of the carboxysome is to maximize the carbon-fixing capacity of Rubisco by defining a microenvironment that maximizes inorganic carbon availability and simultaneously minimizes photorespiratory flux ( Long et al., 2007 ). Carboxysomes respond to environmental changes by adjusting in number, size, and spatial organization based on CO 2 , light availability, redox state, temperature, and light quality ( Sun et al., 2016 ; Rohnke et al., 2018 ; Sun et al., 2019 ; Rillema et al., 2021 ; Lucius and Hagemann, 2024 ). Their positioning and carbon fixation capacity also vary with diurnal cycles and carbon demand ( Sun et al., 2020 ; Singh et al., 2022 ). For example, in the model cyanobacterium Synechococcous elongatus PCC 7942 ( S. elongatus ), an increase in irradiance is generally correlated with increased carboxysome number ( Sun et al., 2016 ), suggestive of a regulatory link between the availability of light energy and cellular investment in carbon fixation machinery. Similarly, we and others have recently shown that S. elongatus engineered to have a higher metabolic “demand” via the expression of heterologous metabolism also exhibit an increase in carboxysome number ( Santos-Merino et al., 2021a ; Singh et al., 2022 ). Therefore, the “upstream” processes of photosynthetic light reactions appear to be integrated with the “downstream” energy demands of the cell for carbon fixation and metabolism, although regulatory mechanisms that accomplish this are poorly understood ( Santos-Merino et al., 2021a ). We recently conducted a screen of all known two-component regulatory proteins S. elongatus with the goal of identifying potential cyanobacterial networks important for achieving energy balance between photosynthetic energy harvesting and integrated metabolic demand ( Santos-Merino et al., 2024 ). Briefly, our approach used a heterologous metabolic pathway (sucrose secretion) that can be experimentally activated to significantly draw upon primary products of photosynthesis. Activation of the sucrose secretion pathway has been previously shown to lead to a range of photosynthetic changes, including increased carboxysome number, increased CO 2 fixation rates, increased oxygen evolution, and reduced acceptor side limitation of PSI activity ( Abramson et al., 2016 ; Santos-Merino et al., 2021b ; Singh et al., 2022 ). Our screen for regulatory proteins involved in this process implicated RpaA, ManS, CikB, and NblS as leading factors important in coupling changes in metabolic demand in the cell to upstream enhancements in the flux photosynthetic processes. Herein, we report potential mechanisms that link RpaA function to the control of the organization of the carboxysome in response to ROS signals derived from an over-reduced pETC. Our results have implications for fundamental understanding of cyanobacterial energy balancing pathways and may provide insight into how the carboxysome is remodeled in response to changing environments and the utilization of central carbon metabolic intermediates. 2. Materials and methods 2.1. Strains and culture conditions S. elongatus cultures were grown in BG11 medium supplemented with 1 g L −1 HEPES to a final pH of 8.3 with NaOH. Flasks were cultured in a Multitron incubator (Infors HT) at 32 °C under ambient air CO 2 or supplemented with 2% CO 2 with ∼150 μmol photons m −2 s −1 of light provided by Sylvania 15 W Gro-Lux fluorescent bulbs and shaken at 150 rpm. Cultures were back-diluted daily to an OD 750 of 0.3 and acclimated to the medium/irradiance for at least 3 days prior to experiments or isopropyl-β-D-thiogalactoside (IPTG) induction. Where appropriate, 1 mM IPTG was added to induce cscB and sps gene expression. Erythromycin (Em; 100 μg mL −1 ), chloramphenicol (Cm; 25 μg mL −1 ), and spectinomycin (Sp; 100 μg mL −1 ) were used to maintain cscB-sps -, cscB -, and rbcS-mNG -containing cells, respectively. Kanamycin (Kn; 12.5 μg mL −1 ) was used to maintain the rpaA inactivation mutant Δ rpaA . In all cases, antibiotic selection was removed prior to conducting any of the reported experiments to minimize any unintended effects. When indicated, cultures were grown in the presence of photosynthesis inhibitors, including DCMU (20 µM) or DBMIB (10 µM). All strains used in this study are listed in Table 1 . View this table: View inline View popup Table 1. Cyanobacterial strains used in this study. 2.2. Strain construction S. elongatus with genomically integrated copies of cscB and sps under an IPTG-inducible promoter was previously obtained ( Santos-Merino et al., 2024 ), and S. elongatus with genomically integrated copies of cscB under an IPTG-inducible promoter was previously described ( Ducat et al., 2012 ). Genomic loci encoding rpaA gene was disrupted by inserting a kanamycin resistance cassette ( Qiao et al., 2019 ). To allow visualization of changes in carboxysome organization, we integrated a fluorescent reporter fused to the small subunit of Rubisco (RbcS-mNG) in NS1 by modifying a plasmid previously published ( Sakkos et al., 2021 ). Plasmid details are reported in Table 2 . View this table: View inline View popup Download powerpoint Table 2. Plasmids used in this study. 2.3. Sucrose quantification Secreted sucrose was quantified from supernatants using the Sucrose/D-Glucose Assay Kit (K-SUCGL; Megazyme). 2.4. Pigment determination Chl a was extracted from cell pellets by incubation in 100% methanol for 30 min at 4 °C. Chl a concentration was estimated by the spectrophotometric method described previously ( Porra et al., 1989 ). 2.5. Fluorescence measurements Apparent quantum yield of PSII (Φ II ) measurements were performed on a custom-built fluorimeter/spectrophotometer as described previously ( Santos-Merino et al., 2021b ). Briefly, samples containing cyanobacteria (2.5 μg mL −1 chlorophyll) resuspended in fresh medium sparged with 2% CO 2 in air were dark-adapted for 3 min before measuring. The apparent quantum yield of photosystem II (Φ II ), (F’ M - F S )/(F’ M ), and the coefficient of photochemical quenching (q p ), (F’ M - F S )/(F’ M - F’ 0 ), were measured using a 1.5 s saturating pulses of actinic light (∼5,000 μmol photons m −2 s −1 ). 2.6. PSI absorbance changes To evaluate P700 redox changes, samples were monitored semi-simultaneously with fluorescence measurements, using the instrument described in ( Hall et al., 2013 ) by measuring absorbance changes at about 703 nm. The measuring beam was generated by a pulsed LED (720 nm peak emission, Rebel LUXEON Far Red) filtered with a 5 nm band pass filter centered at 700 nm, resulting in a measured emission peak at approximately 703 nm ( Abramson et al., 2016 ). The signals were detected with a photodiode filtered with a Schott RG-695 filter to block actinic light ( Hall et al., 2013 ). Samples containing cyanobacteria cells were prepared by resuspended cells in fresh medium to a concentration of 5 μg mL −1 chlorophyll and sparged with 2% CO 2 in air. The samples were dark-adapted for 3 min before starting measurements. The percentage of oxidized P700 was calculated as [(P ox - P ss )/(P red - P ox )] · 100, where P ox was the maximum extent of P700 absorbance signal induced by a saturating pulse of light during ∼0.5 s (∼5,000 μmol photons m −2 s −1 ); P ss , taken to be the fraction of P700 + in under steady state illumination, estimated by the extent of P700 absorbance signal induced by a short dark interval, and assuming that P700 reaches full reduction; and P red is the level of P700 reduced in the dark. PSI traces were normalized to the last point of the steady-state level of oxidation (P ss ). 2.7. Dark-interval relaxation kinetics (DIRK) absorbance changes Steady-state levels of photooxidized P700 (P700 + ) were estimated by DIRK analysis ( Sacksteder and Kramer, 2000 ), after 21 s of actinic illumination at three different intensities (100, 275 and 500 μmol photons m −2 s −1 ). Samples containing cyanobacteria (2.5 μg mL −1 chlorophyll) resuspended in fresh medium containing 2% CO 2 and were dark-adapted for 3 min before measuring. The half-time of P700 + re-reduction (Tau), was measured from the absorbance change at 703 nm (ΔA 703 ) during a dark interval of 2100 ms. Tau was calculated by the monotonic decay kinetic of ΔA 703 produced after extinguishing the actinic light ( Baker et al., 2007 ). 2.8. Microscopy and Image Analysis All live-cell microscopy was performed on cells in exponential growth by centrifuging 2 mL of culture at 10,000 × g for 5 min, resuspending into 80 μL of BG11, and transferring a 2 μL aliquot to a 3% agarose pad. The cells were allowed to briefly equilibrate and be absorbed by the agarose (≥10 min) before the pad was placed onto a #1.5 glass coverslip for imaging. Images were captured using a Zeiss Axio Observer D1 inverted microscope equipped with an Axiocam 503 mono camera and a Zeiss Plan Apochromat 63X 1.4 NA oil-immersion lens. Image analysis was done in OMERO ( Allan et al., 2012 ) and Python 3. Cell segmentation was conducted using Cellpose ( Stringer et al., 2021 ). Mean fluorescence intensity plots were generated as previously described ( Sakkos et al., 2021 ). Briefly, cells were segmented using the chlorophyll autofluorescence channel, rotated such that the medial axis was horizontal, rescaled to ensure consistent boundaries, and the RbcS-mNG pixel intensity was averaged from each cell in the collection of images from its respective induction condition and time point. Foci locations were determined with a peak-finding algorithm using the Python package Photutils ( Bradley et al., 2016 ). 2.9. Cellular viability Cellular death was quantified using SYTOX Blue (1 mM in DMSO; Invitrogen, S34857) on a flow cytometer or SYTOX Orange (250 μM in DMSO; Invitrogen, S34861) and visualized by microscopy. SYTOX dyes are nucleic-acid-specific stains that are unable to reach the intracellular space due to the intact cell membrane of a non-damaged cell. At each time-point, 1 μL of the SYTOX blue working stock was added to 1 mL of culture (final concentration of 1 μM for SYTOX Blue or 250 nM for SYTOX Orange) and incubated in the dark at room temperature for 15 min. Heat-treated cyanobacterial cells were used as a positive control for dead cells. After incubation, 200 μL aliquots of the cultures were transferred to a 96-well plate to measure viability in the flow cytometer. Samples were collected on a 4-laser Attune CytPix with a CytKick Max Autosampler Software 6.2.0. The following optical configuration was used for each fluorophore (excitation|emission): SYTOX Blue [405nm|450/40] (BL1-A, 305V), Chlorophyll [405nm|660/20] (486V). Cyanobacterial samples were gated using FSC (400V) and SSC (250V) to distinguish the singlet population and with the chlorophyll to remove debris and noise, more than 10,000 cells were measured per sample type unless stated otherwise. Gating regions representing both intact, SYTOX-negative and membrane-damaged, SYTOX-positive cells were created in two-dimensional dot-plots (forward side scatter versus blue fluorescence). The raw data were analyzed using the FCS Express 7 software (De Novo Software, USA). For the microscope images, samples were processed following a protocol described in section 2.8 . 2.10. Membrane inlet mass spectrometry (MIMS) Online measurements of gas exchange were monitored using a mass spectrometer (model Prima PRO, Thermo Scientific). The membrane inlet system, consisting of a modified DW1 oxygen electrode chamber (Hansatech Instruments Ltd.) water-jacketed thermoregulated at 30 °C, was attached to the vacuum line of a mass spectrometer through a thin gas-permeable PTFE membrane (0.0125 mm) sealing the bottom of the chamber. 18 O 2 (isotope purity >98%; CK Gas Products Ltd.) tracing was used to discriminate O 2 uptake and O 2 production by PSII. 16 O 2 ( m/z 32), 18 O 2 ( m/z 36) and CO 2 ( m/z 44) were recorded with a time resolution of around 4 s. Samples were evenly mixed by constant stirring using a cross-shaped magnetic stirrer. A 2 mL aliquot of a cell suspension (10 μg mL −1 Chl a ) was placed in the measuring chamber and prior to the measurement, cells were supplemented with 18 O 2 at an equivalent concentration to 16 O 2 and with 1.5 mM NaHCO 3 . Then, samples were measured for 5 min in darkness to record oxygen consumption caused by respiration. Following this period, actinic light (500 μmol photons m −2 s −1 ) was applied via a 150-W, 21-V EKE quartz halogen-powered fiber optic illuminator (Fiber-Lite DC-950; Dolan-Jenner). Gas-exchange kinetics and rates were determined according to ( Beckmann et al., 2009 ). Final Chl a concentration, determined spectrophotometrically in 100% methanol according to Porra et al. ( Porra et al., 1989 ), was conducted at completion of each measurement for standardizing the calculated gas exchange rates. 2.11. Determination of intracellular glycogen content Glycogen content was determined as described previously with minor modifications ( Gründel et al., 2012 ). Cyanobacterial culture aliquots (2 mL) were pelleted down by centrifuging at 5,000 xg for 10 minutes. Pellets were flash-frozen in liquid nitrogen and were stored at -80 °C until extraction. For isolation of glycogen, the pellets were resuspended in 200 μL 30% (w/v) KOH and incubated in a heat block at 95 °C for 2 hours. Samples were cooled down on ice. Complete precipitation of glycogen was achieved by the addition of 600 µL of cold absolute ethanol and overnight incubation at −20 °C. The precipitated glycogen was recovered by centrifugation at 17,000 xg for 15 min at 4 °C. The supernatant was removed, and the glycogen pellets were dried for 40 min at 60°C using a SpeedVac. The precipitated glycogen was resuspended in 200 µL of miliQ H 2 O by vortexing. The homogeneous samples were quantified using the EnzyChrome glycogen assay kit (BioAssay Systems, E2GN-100) according to the manufacturer’s instructions. 2.12. Transmission Electron Microscopy At each time point, 2 ml samples were prepared by diluting cultures to a final OD 750 1.5. Cells were pelleted and fixed overnight at 4 °C with 2% glutaraldehyde/2% paraformaldehyde in phosphate buffer (pH 7.4), suspended into a 2% agarose bead and cut into ∼1 mm cubes. Following three washes with 0.1 M sodium cacodylate buffer, cells were suspended in 1% osmium tetroxide/1.5% potassium ferrocyanide and incubated overnight at 4 °C. After incubation, cells were washed with HPLC-quality H 2 O until they appear clear. Cells were then suspended in 1% uranyl acetate and microwaved for 2 min using a MS-9000 Laboratory Microwave Oven (Electron Microscopy Science), decanted, and washed until clear. Cells were dehydrated in increasing acetone series (microwave 2 min) and then embedded in Spurr’s resin (25% increments for 10 min each at 25 °C). A final overnight incubation at room temperature in Spurr’s resin was done, then cells were embedded in blocks which were polymerized by incubation at 60 °C for 3 days. Thin sections of approximately 50 nm were obtained using an MYX ultramicrotome (RMC Products), post-stained with 1% uranyl acetate and Reynolds lead citrate, and visualized on a JEM 100CX II transmission electron microscope (JEOL) equipped with an Orius SC200-830 CCD camera (Gatan). 2.13. Quantification of Reactive Oxygen Species (ROS) ROS were quantified by using the fluorescent marker H 2 DCFDA (2′,7′-dichlorodihydrofluorescein diacetate; Invitrogen, D399) as previously reported ( Diamond et al., 2017 ). Briefly, 2 mL of the cultures were collected and split into 1-mL aliquots. H 2 DCFDA was added to one sample at a final concentration of 5 μM. Tubes were protected from light and shaken at 30 °C for 30 min. After incubation, 200 μL of each tube was added to a separate well in a 96-well plate. The fluorescent product 2’,7’-dichlorofluorescein (DCF) was monitored via microplate reader (excitation 480 nm, emission 520 nm; SpectraMax M2 microplate reader by Molecular Devices). Untreated-sample background fluorescence was then subtracted from treated-sample fluorescence values and fluorescence data were normalized to OD 750 of each sample. 2.14. Determination of growth of cyanobacterial cultures in the presence of hydrogen peroxide To determine the growth of the different cyanobacterial strains in the presence of H 2 O 2 , cell cultures were adjusted to an OD 750 of 0.3. Hydrogen peroxide, at final concentrations ranging from 50 μM to 2 mM, was added to 1.5-mL aliquots of cultures in separate wells of a 24-well plate or to 50-mL flasks with cultures. Images of the plates, as well as fluorescence microscopy images, were taken after a 24-h incubation under standard growth conditions. 2.15. Statistical analysis Recorded measurements are represented as mean values, with error bars expressing the SD of n≥3 biological replicates experiments, as indicated. The significance of differences between groups was evaluated by one-way ANOVA followed by Tukey’s multiple comparison test or by an unpaired Student’s t-test. Statistical analyses were carried out using GraphPad Prism software (GraphPad Software Inc., San Diego, CA). Differences were considered statistically significant at P < 0.05. 3. Results 3.1. Remodeling carbon fixation machinery following sucrose secretion requires rpaA As discussed in the introduction, we recently implicated RpaA as a two-component protein that appears to be involved in the regulation of photosynthetic changes following activation of a heterologous sucrose secretion pathway ( Santos-Merino et al., 2024 ). To further investigate the potential roles of RpaA in adapting photosynthesis and the CCM we first validated the impact of loss-of-function mutants (Δ rpaA ) in a strain expressing a carboxysome reporter (RbcS-mNG) and bearing an inducible sucrose-export pathway (CscB-SPS export ). We confirmed that Δ rpaA strains lacked the characteristic rise in photosynthetic activity following activation of sucrose secretion, as measured by apparent quantum efficiency of photosystem II (Φ II ; Figure 1A ). Similarly, deletion of rpaA abrogated the increase in carboxysome number and size that follows sucrose secretion ( Singh et al., 2022 ), as monitored by the carboxysome reporter strain ( Figures 1B an 1C and Supplementary Figure S1). Importantly, the Δ rpaA background growth rate and other photosynthetic parameters appeared comparable to WT under the laboratory cultivation conditions with constant light, although Δ rpaA cell size was slightly diminished (Supplementary Figure S2). These results are consistent with other recent reports that have demonstrated that Δ rpaA strains perform similarly to wildtype under conditions of constant light and high CO 2 ; sometimes even exceeding wildtype growth ( Espinosa et al., 2015 ; Diamond et al., 2017 ; Puszynska and O’Shea, 2017 ). Therefore, deletion of rpaA eliminates the enhancements in photosynthesis and increased carboxysome number typically observed in cells induced to export sucrose ( Figures 1 and Supplementary Figure S1) but does not globally misregulate essential pathways required for growth and division under controlled laboratory conditions (Supplementary Figure S2). Download figure Open in new tab Figure 1. Acclimatory changes in the photosynthetic machinery following sucrose export are absent in Δ rpaA . (A) Apparent Φ II values measured at three different light intensities 24 hours after the induction of sucrose export in the strain CscB-SPS export in the presence/absence of RpaA. (B) Carboxysome number change (expressed as percentage) after induction of sucrose export in the strain CscB-SPS export in the presence/absence of RpaA. (C) Carboxysomes at 72 hours visualized via the RbcS-mNG reporter in the strain CscB-SPS export in the presence/absence of RpaA. Scale bar: 2 µm. (A, B) Averages of ≥3 independent biological replicates are shown + SD. Significance was calculated by one-way ANOVA followed by Tukey’s multiple comparison test. Data points labeled with different letters are significantly different (P < 0.05). 3.2. Δ rpaA mutants exhibit growth arrest and dramatic carboxysome disassembly under mixotrophic conditions As we and others have shown that carboxysome structure and composition is impacted by growth under mixotrophic conditions ( Muth-Pawlak et al., 2022 ; Singh et al., 2022 ), we next examined the impact of rpaA knockout on carboxysome restructuring under mixotrophy. We encoded the sucrose permease gene under the IPTG-inducible P trc promoter to allow genetic control over sucrose import (CscB import ) and supplemented the growth medium with sucrose. In a wildtype background, mixotrophic growth mildly suppressed carboxysome number and increased the incidence of carboxysome clustering ( Figure 2A ), consistent with prior studies ( Singh et al., 2022 ). By contrast, under conditions where CscB was activated, allowing uptake of extracellular sucrose, Δ rpaA strains exhibited an immediate growth arrest and carboxysome reorganization followed by a dramatic disassembly of carboxysomes within 1-3 days of the onset of mixotrophic growth ( Figures 2A-2C and Supplementary Figure S3). The loss of carboxysomal integrity appeared to be gradual, with some Δ rpaA cells displaying an increase in delocalized Rubisco signal throughout the cytosol in the population as early as 24 hours, but only a fraction of cells exhibiting a complete loss of carboxysomal puncta (Supplementary Figure S3A). The fraction of cells with diminished carboxysome puncta and increased cytosolic Rubisco localization increased over time, with nearly all cells lacking carboxysomes after 72 hours of sucrose feeding ( Figure 2A ). Finally, we observed that cells displaying increased autofluorescence of natural cyanobacterial pigments (possibly associated with impaired photosynthetic performance) were associated with a more rapid loss of carboxysome puncta over time ( Figure 2A ). Download figure Open in new tab Figure 2. Mixotrophic conditions induces carboxysome breakdown and growth arrest in Δ rpaA . (A) Time-course of carboxysome status under photoautotrophic and mixotrophic conditions in the strain CscB import in the presence/absence of RpaA. Scale bar: 2 µm. (B) Growth curves of the strain CscB import in the presence/absence of RpaA in photoautotrophic and mixotrophic conditions with zoom in of the growth curve of CscB/RbcS-mNG/Δ rpaA . Averages of ≥3 independent biological replicates are shown ± SD. (C) Appearance of the cultures of the strain CscB import in the presence/absence of RpaA under photoautotrophic and mixotrophic conditions. (D) Cartoon illustration of internal and external carboxysome components. (E) Carboxysome status under mixotrophic conditions in the strain CscB import in the presence/absence of RpaA by tracking mNG-McdB. Scale bar: 2 µm. The observed loss of puncta following mixotrophic growth conditions must be attributed to dissolution of existing carboxysomes rather than failure to build new microcompartments because of the complete growth arrest observed ( Figure 2B ). To better assess the dynamics of carboxysome breakdown, we monitored dynamics of an endogenously tagged outer carboxysome component maintenance of carboxysome distribution B (McdB)( Figure 2D ), a ParB-family protein involved in microcompartment positioning ( MacCready et al., 2018 ). McdB utilizes a conserved motif to bind to the cytosolic-facing side of bacterial microcompartment shell proteins, and it is therefore a relatively peripheral component of the carboxysome ( Basalla et al., 2024 ). Within 24 hours following onset of mixotrophic growth in Δ rpaA cells, we observed a near complete loss of McdB-mNG with the carboxysome shell in nearly all cells ( Figure 2E ). By comparison, the Rubisco core of carboxysomes remained present at this time point in most cells (Supplementary Figure S4), suggesting that carboxysome disassembly may initiate with components associated with the shell proteins and proceed inward. Consistent with the loss of the positional machinery, we observed that Rubisco puncta in mixotrophic Δ rpaA cells were frequently mispositioned relative to one another or localized to the cell pole (Supplementary Figure S5). The localization of Rubisco to cell poles has been proposed to be an initiating step of carboxysome disassembly in prior studies ( Hill et al., 2020 ). 3.3. Δ rpaA cells undergo a severe photosynthetic impairment under mixotrophy The physiological characteristics of rpaA mutants grown under mixotrophic conditions ( i.e. , cell growth arrest, bleaching of pigments, and carboxysome breakdown) ( Figure 2 and Supplementary Figures S3 and S4) suggest that photosynthetic processes may be misregulated ( Sunil et al., 2013 ; Calzadilla and Kirilovsky, 2020 ). We directly assessed photosynthetic performance of cultures grown under photoautotrophic and mixotrophic conditions via fluorimetry. In sucrose-fed Δ rpaA lines, the openness of PSII reaction centers (q p ) was dramatically decreased at all tested levels of illumination relative to modest changes in sucrose-fed control strain CscB/RbcS-mNG ( Figure 3A ). Similarly, the apparent Φ II was reduced almost to zero under mixotrophic growth in Δ rpaA cultures, although Φ II was also partially suppressed by sucrose feeding in the reference control ( Figure 3B ). Other measured photosynthetic values were not as strongly impacted by sucrose feeding in either line, including the oxidation state of PSI or the estimates of electron flux from cytochrome b 6 f (Supplementary Figures S6A and S6B), overall indicating that PSII and/or electron flux near PQ were the most strongly impacted. Download figure Open in new tab Figure 3. Mixotrophic growth conditions impair PSII activity and have different effects on the rate of O 2 and CO 2 fluxes. (A) q p values. (B) Apparent Φ II values. Quantification of the steady state fluxes rates shown in Supplementary Figures S6C-S6H for (C) CO 2 fixation, (D) gross O 2 evolution, (E) CO 2 dak respiration, and (F) light-dependent O 2 uptake for the strains CscB/RbcS-mNG and CscB/RbcS-mNG/Δ rpaA . CO 2 exchange rates in the strains CscB/RbcS-mNG and CscB/RbcS-mNG/Δ rpaA under photoautotrophic and mixotrophic conditions. Averages of ≥3 independent biological replicates are shown + SD. Significance was calculated by one-way ANOVA followed by Tukey’s multiple comparison test. Data points labeled with different letters are significantly different ( P < 0.05). To gain additional insight into photosynthetic fluxes in control and Δ rpaA lines, we turned to membrane inlet mass spectrometry (MIMS), which can disentangle CO 2 and O 2 gas exchange rates originating from photosynthetic and respiratory metabolism. Consistent with the sharp decline in q P and Φ II , we observed a rapid loss of photosynthetic CO 2 fixation under both photoautotrophic and mixotrophic growth conditions in Δ rpaA cells ( Figure 3C and Supplementary Figures S6C and S6D), accompanied by a loss of O 2 evolution only under mixotrophic conditions in the Δ rpaA strain ( Figure 3D and Supplementary Figures S6E and S6F). Continued CO 2 respiration was observed in the Δ rpaA cells under dark conditions ( Figure 3E and Supplementary Figures S6C and S6D), providing direct evidence that the cells remain metabolically active during growth arrest ( Figure 2 ). Almost a complete loss of O 2 uptake induced by light was observed under mixotrophic conditions ( Figure 3E and Supplementary Figures S6G and S6H), suggesting that reducing equivalents accumulated on the pETC are not efficiently consumed by alternative electron pathways that normally quench excess reductant by donating them to molecular oxygen ( i.e. , water-water cycles) in the Δ rpaA line ( Nikkanen et al., 2021 ). Given the severity of the photosynthetic impairment that mixotrophic conditions initiated in the Δ rpaA line, we used the vital dye SYTOX to confirm that cells exhibiting carboxysome breakdown remained viable. Fluorescence-activated cell sorting (FACS) showed that cell viability began to be impaired 48 hours under mixotrophic growth, with no viable cells after 72 hours ( Figure 4A and Supplementary Figure S7). This indicated that initiation of carboxysome disassembly ( i.e. , evident at 24 hours) precedes that of the loss of cell viability (between 48 and 72 hours under mixotrophic growth). Fluorescence microscopy with SYTOX confirmed this interpretation, as cells that show disruption of carboxysomes remain SYTOX negative 24 and 48 hours into the mixotrophic-induced growth arrest ( Figure 4B and Supplementary Figure S5).To further evaluate the interplay between carboxysome breakdown and cell viability in Δ rpaA cells, we fed sucrose for 60 hours (when nearly all Δ rpaA cells have no evident carboxysomes), then tracked recovery by returning cells to photoautotrophic growth (Supplementary Figure S8). Following sucrose removal, Δ rpaA cells exited the growth arrest (<24 hours), recovered pigmentation (24-48 hours), and reassembled carboxysomes (24-72 hours) (Supplementary Figures S8A-S8E). Whereas growth of Δ rpaA cells immediately resumed when sucrose was removed (Supplementary Figure S8D), recovery of cellular pigmentation and carboxysome puncta was delayed (Supplementary Figures S8A-S8C). A further indication that growth arrested cells remained metabolically active was the greatly-elevated ROS levels in sucrose-fed Δ rpaA cultures; ROS levels gradually returned to the baseline of paired controls by 72 hours following removal of sucrose (Supplementary Figure S8E). Download figure Open in new tab Figure 4. Carboxysome disassemble precedes cell death and is accompanied by glycogen accumulation. (A) Percentage of SYTOX Blue-positive cells (dead cells) measured by flow cytometry under photoautotrophic and mixotrophic conditions, with detail of CscB/RbcS-mNG and CscB/RbcS-mNG/Δ rpaA under photoautotrophic conditions. Averages of ≥3 independent biological replicates are shown ± SD. (B) Time-course of SYTOX Orange-positive cells and carboxysomes by tracking RbcS-mNG under mixotrophic conditions in the strain CscB import in the absence of RpaA. Scale bar: 2 µm. (C) Quantification of glycogen per OD 750 unit at different time points under photoautotrophic and mixotrophic conditions in the CscB strain in the presence and absence of RpaA. Averages of ≥3 independent biological replicates are shown + SD. Significance was calculated by one-way ANOVA followed by Tukey’s multiple comparison test. Data points labeled with different letters are significantly different ( P < 0.05). (D) Transmission electron microscopy of cells of the strain CscB import in the absence of RpaA grown photoautotrophically and mixotrophically; black arrows indicate glycogen granules. Scale bar: 500 nm. RpaA has recently been implicated in regulation of central carbon metabolism and glycogen mobilization ( Diamond et al., 2015 ; Diamond et al., 2017 ; Puszynska and O’Shea, 2017 ; Santos-Merino et al., 2024 ), so we examined the utilization of sucrose in Δ rpaA lines. We observed that sucrose feeding leads to glycogen accumulation in both wild-type (WT) and Δ rpaA backgrounds in the first 24 hours. However, while WT background restored normal glycogen levels by 48 hours, Δ rpaA lines exhibited a continual accumulation of glycogen over time ( Figure 4C ). Indeed, glycogen hyperaccumulation in sucrose-fed Δ rpaA cells was directly observable by electron microscopy, which revealed that glycogen bodies proliferated throughout the cytosol and within the thylakoid membranes ( Figure 4D ). Taken together, fluorescence kinetics, MIMS, vital dyes, and electron microscopy suggest that mixotrophic growth in Δ rpaA line induces severe impairment of photosynthetic reactions, arrests cell growth while maintaining metabolic activity, triggers abnormal levels of glycogen deposition, initiates carboxysome disassembly, and eventually can lead to cell death under prolonged exposure. 3.4. PQ overreduction leads to H 2 O 2 formation and carboxysome breakdown A variety of stressors can lead to imbalances within the pETC, often leading to activation of photoprotective mechanisms including alternative pathways to quench excess reductant to avoid formation of ROS and photodamage ( Pospisil, 2016 ). In the Δ rpaA mutant, we observe several signs that indicate an accumulation of electrons in the pETC, including a drop in the values of q p and in the apparent Φ II ( Figures 3A and 3B ), and the loss of gross O 2 evolution ( Figures 3D and 3F and Supplementary Figures S6E-S6H). Moreover, the suppression of light-induced O 2 uptake we observed in mixotrophic Δ rpaA strains ( Figure 3F ) suggests that alternative pathways for quenching electrons in the pETC are suppressed ( e.g. , flavodiiron proteins Flv1/3), which could further exacerbate the buildup of reductant ( Diamond et al., 2017 ). Because of the indicators of potential imbalances in the pETC, we directly monitored ROS production in Δ rpaA cells under mixotrophic conditions and observed a sharp increase in ROS production that peaked at 72 hours after initiation of sucrose feeding ( Figure 5A ). To further dissect the phenotype, we used chemical inhibitors and growth conditions to modulate redox status of the pETC. Treatment of Δ rpaA cells with 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), a specific inhibitor of PSII that blocks electron transport from Q A to Q B and thereby oxidizes the pETC at PQ and downstream electron carriers, decreased ROS production under mixotrophic conditions and prevented carboxysome breakdown ( Figures 5A and 5C and Supplementary Figure S10C). In parallel, sucrose feeding of Δ rpaA in the dark, where photosynthesis is inactive ( Khorobrykh et al., 2020 ), also prevented both ROS production and preserved carboxysome integrity. By contrast, treatment with dibromothymoquinone (DBMIB) blocks electron transfer from PQ to Cytochrome b 6 f, did not rescue sucrose-fed Δ rpaA cells, which continued to strongly produce ROS and exhibit carboxysome breakdown ( Figures 5A and 5C and Supplementary Figure S10C). Darkness and DCMU treatment also recovered the loss in cell viability and pigmentation usually observed in mixotrophic Δ rpaA cells (Supplementary Figure S10). Taken together, our results suggest overreduction of PQ in Δ rpaA cultures is associated with ROS formation and carboxysome breakdown. Download figure Open in new tab Figure 5. Reduction of the PQ pool leads to accumulation of ROS and carboxysome disassembly. Quantification of cellular ROS accumulation measured by H 2 DCFDA fluorescence at different time points (A) following activation of sucrose import, and (B) following the addition of photosynthesis inhibitors or growing the cells in darkness in sucrose feeding conditions in the CscB import strain in the absence of RpaA. (C) Carboxysome status at 72 hours in response to photosynthesis inhibitors or growing the cells in darkness in sucrose feeding conditions in the CscB import strain in the presence/absence of RpaA by tracking RbcS-mNG in the fluorescence microscope. Scale bar: 2 µm. (D) Carboxysome status after 24 hours exposure to different concentrations of H 2 O 2 in the CscB import strain in the presence/absence of RpaA by tracking RbcS-mNG. Scale bar: 2 µm. (E) Carboxysome status after exposure to 1 mM of H 2 O 2 for CscB import strain and 400 µM H 2 O 2 for CscB import strain in the absence of RpaA by tracking RbcS-mNG. Scale bar: 2 µm. (A, B) Averages of ≥3 independent biological replicates are shown + SD. Significance was calculated by one-way ANOVA followed by Tukey’s multiple comparison test. Data points labeled with different letters are significantly different ( P < 0.05). Hydrogen peroxide (H 2 O 2 ) is a dominant form of ROS that has established signaling roles in plants ( Mubarakshina and Ivanov, 2010 ), although any conserved regulatory function in cyanobacteria has not been clearly established ( Latifi et al., 2009 ). We therefore tested direct application of H 2 O 2 , observing that this treatment also impacted cellular pigmentation and carboxysome integrity in both WT and Δ rpaA strains ( Figure 5D ). Interestingly, the Δ rpaA mutant exhibited increased sensitivity to external addition of H 2 O 2 in terms of cell viability, pigmentation, and carboxysome integrity ( Figure 5D and Supplementary Figure S11). Acute treatment with H 2 O 2 (2 mM H 2 O 2 for WT or 750 µM H 2 O 2 for Δ rpaA ) led to a rapid bleaching and disassembly of most carboxysomes within the first 24 hours. After acute H 2 O 2 treatment, those cells that still contained visible RbcS-mNG puncta typically possessed only one or two carboxysomes which were typically located in the pole of the cells (Supplementary Figure S11D); this localization pattern has been previously reported as preceding carboxysome degradation ( Hill et al., 2020 ). Yet, cell cultures did not always recover growth following acute H 2 O 2 treatment, complicating interpretation of these results. We therefore conducted further analysis using H 2 O 2 treatments at concentrations where cell growth and pigmentation were unaffected (Supplementary Figure S11A). Cells treated with subacute H 2 O 2 (1 mM for WT or 400µM for Δ rpaA ) did not exhibit substantive changes in growth or pigmentation (Supplementary Figure 12). However, in the hours immediately following subacute H 2 O 2 treatment, subtle alterations in carboxysome number and RbcS-mNG puncta organization were observed ( Figure 5E ). Crucially, at later time points (36-72 hours post H 2 O 2 exposure), carboxysome phenotypes appeared exacerbated, with many cells in the population displaying heterogeneity in carboxysome brightness, mispositioned/polar carboxysome puncta, too few carboxysomes or no carboxysome puncta at all ( Figure 5E and Supplementary Figure S12). The carboxysome phenotypes at later time points (≥ 36 hours) following subacute H 2 O 2 treatment was less severe but were similar in character and timing to phenotypes observed under mixotrophic growth (>24 hours). 3.5. Physiological conditions that lead to ROS generation are associated with carboxysome rearrangement We next asked if other physiologically-relevant environmental conditions are associated with carboxysome disassembly beyond mixotrophic growth and we monitored any correlation with redox imbalance and ROS production. Light and inorganic carbon availability are two critical environmental factors strongly influencing cyanobacterial photosynthetic performance and ROS generation ( Khorobrykh et al., 2020 ; Krieger-Liszkay and Shimakawa, 2022 ), therefore we revisited WT and Δ rpaA cells under ambient CO 2 conditions while varying both light intensity (HL, 150 μmol photons m −2 s −1 ; LL, 17 μmol photons m −2 s −1 ) and sucrose feeding. At ambient CO 2 levels and HL, we found that Δ rpaA displayed substantial carboxysome breakdown even in the absence of sucrose feeding ( Figure 6A , HL). Ambient CO 2 alone was insufficient to induce carboxysome disassembly in Δ rpaA cells, as carboxysomes were well maintained in LL conditions both under photoautotrophic and mixotrophic conditions ( Figure 6A , LL). We also detected an increase in ROS accumulation that was more pronounced under photoautotrophic conditions at both HL and LL in comparison to mixotrophic conditions ( Figure 6B ), though the absolute values of detected ROS were substantially higher across all tested cultures under air relative to 3% CO 2 ( Figure 5A ). Intriguingly, mixotrophic growth partially rescued the loss of carboxysomes in Δ rpaA cells under ambient CO 2 and HL ( Figure 6A ). In each of these cases, elevated ROS was strongly correlated with carboxysome disassembly ( Figure 6B ), as well as chlorophyll content and cell viability, although growth arrest was not observed under these conditions (Supplementary Figures S13 and S14). Download figure Open in new tab Figure 6. Carboxysome breakdown is delayed under ambient air conditions and low light intensity. (A) Carboxysome status in response to ambient air conditions under mixotrophic conditions for the strains CscB/RbcS-mNG and CscB/RbcS-mNG/Δ rpaA by tracking RbcS-mNG. Scale bar: 2 µm. (B) Quantification of cellular ROS accumulation measured by H 2 DCFDA fluorescence at different time points following the transference to ambient air conditions during sucrose feeding conditions for the strains CscB/RbcS-mNG and CscB/RbcS-mNG/Δ rpaA . Averages of ≥3 independent biological replicates are shown + SD. Significance was calculated by one-way ANOVA followed by Tukey’s multiple comparison test. Data points labeled with different letters are significantly different ( P < 0.05). 4. Discussion Taken together, our data supports a novel connectivity between the redox status of the PQ pool and carboxysome status is linked to RpaA function. Recently, we reported that deletion of RpaA prevented a number of acclimatory responses in the light reactions of photosynthesis ( e.g. , increased O 2 evolution and apparent Φ II ) that are typically induced by activation of an engineered sucrose sink ( Santos-Merino et al., 2024 ). Another photosynthetic impact we and others have observed following activation of sucrose export pathways is an increase in CO 2 fixation rate that is associated with a change in carboxysome number and increased rubisco content ( Ducat et al., 2012 ; Singh et al., 2022 ; Wang et al., 2023 ). Here, we show that Δ rpaA mutants also fail to reorganize carboxysomes following sucrose export, but instead can display dramatic reorganization and disassembly of carboxysomes under environmental conditions that lead to overreduction of the pETC and formation of ROS ( Figures 2 , 5 and 6 and Supplementary Figures S3, S4 and S12). Under prolonged conditions of H 2 O 2 generation, carboxysome breakdown can be complete, although carboxysomes will reform in cells once the stress is removed ( Figure 2 and Supplementary Figures S3 and S8). Importantly, chemical inhibitors that block photosynthetic reduction of the PQ pool prevent the carboxysome breakdown phenotype ( Figure 5 and Supplementary Figure S10), suggesting that PQ redox status is especially important to levels of ROS and/or carboxysome integrity. Finally, we show that direct addition of H 2 O 2 can initiate a reorganization or breakdown of carboxysomes in vivo ( Figure 5D and Supplementary Figure S11). One possible interpretation of our data is that the reduction status of PQ and associated ROS formation is part of a normal mechanism that cyanobacteria use to remodel the carboxysome in response to different environmental changes. It has now been well documented that the size, number, and subunit composition of carboxysomes is correlated with light and CO 2 levels in a number of model cyanobacterial species ( Sun et al., 2016 ; Rohnke et al., 2018 ; Sun et al., 2019 ; Rillema et al., 2021 ; Lucius and Hagemann, 2024 ). In one particularly relevant example, high light was shown to significantly increase in carboxysome number in S. elongatus, but carboxysome remodeling was partially blocked via addition of DCMU ( Sun et al., 2016 ), although the mechanism for this block was unclear. It is also established that certain environmental conditions ( e.g. , high light, low CO 2 ), can lead to over-reduction of the PQ pool and resulting production of ROS ( Lea-Smith et al., 2013 ). Our data extends upon prior publications by suggesting PQ-dependent carboxysome remodeling may be a function of ROS (potentially H 2 O 2 ) derived from overreduction of this redox carrier. Cyanobacteria usually exhibit constitutive expression and assembly of carboxysomes and (to our knowledge) the complete breakdown of carboxysomes is nor reported under physiological conditions elsewhere, although other studies have examined conditions where carboxysome integrity is compromised. First, live-cell imaging studies of cyanobacteria engineered to be gradually depleted of key structural components of the carboxysome have demonstrated that degrading carboxysomes are mislocalized from the nucleoid to the cell pole prior to the dissolution of the Rubisco core ( Hill et al., 2020 ). This elegant imaging of the carboxysome lifecycle matches our observation of mispositioned and polar Rubisco puncta that precede carboxysome loss in Δ rpaA mutants ( Figures 2 , 5, and 6 and Supplementary Figures S3, S4, S5, S10, and S11). The cyanobacterial species, Microcystis aeruginosa , produces a toxin called microcystin that binds to Rubisco and interferes with its encapsulation within the carboxysome lumen, resulting in delocalized and thylakoid-associated Rubisco aggregates ( Barchewitz et al., 2019 ). It has also been shown that H 2 O 2 degrading activities of thioredoxin and peroxiredoxin are also inhibited by microcystin ( Alexova et al., 2016 ; Schuurmans et al., 2018 ), which could sensitize M. aeruginosa to ROS-related pathways, including the carboxysome breakdown phenotype we report here. Based on our data and prior literature Δ rpaA mutants appear to be particularly sensitized to redox imbalances and may thereby exhibit an amplified response to a redox related regulatory signal. RpaA is involved in controlling redox balance, as observed for the incapacity of Δ rpaA mutant to clear ROS out during the night, that have been excessively accumulated during the day ( Diamond et al., 2017 ). The inability to detoxify ROS of the Δ rpaA cells makes them unable to survive the dark period, leading to a conditional light–dark lethality. As deletion of RpaA has been associated with misregulation of the expression of hundreds of genes in cyanobacteria ( Markson et al., 2013 ), it is difficult to predict how directly RpaA is involved in mitigating or transducing ROS signals. However, Δ rpaA mutants are strongly downregulated in thioredoxin-dependent peroxidase (2-cys prx) ( Markson et al., 2013 ; Puszynska and O’Shea, 2017 ), and Δ rpaA mutants exhibit a deficit in NADPH that is required for the activity of ROS scavenger enzymes ( Diamond et al., 2017 ). Indeed, 2-cys prx is the main enzyme in S. elongatus responsible for detoxifying ROS and has been previously shown to be upregulated under photomixotrophic growth conditions ( Perelman et al., 2003 ; Tan et al., 2022 ). More directly, we have recently shown enrichment of thioredoxin peroxidase and catalase in proximity labeling studies with RpaA ( Santos-Merino et al., 2024 ), indicating a possible direct RpaA interaction that would require additional study. In this same interactome, we detected a putative RpaA-Flv3 interaction that may merit additional validation given that we observe a loss of light-induced O 2 uptake catalyzed by the flavodiiron proteins Flv1/Flv3. Curiously, recent reports connect RpaA with the redox state of the pETC through interaction with ferredoxin (Fd) and thioredoxin (Trx) ( Hanke et al., 2011 ; Kadowaki et al., 2015 ), and the activity of the latter act as a thiol redox switch to regulate the oligomeric state of the RpaA through redox-responsive cysteines ( Ibrahim et al., 2022 ). RpaA has also been implicated in central carbon metabolism and storage in cyanobacteria in ways that may indirectly contribute to the sensitivity of rpaA mutants to redox imbalances. Inefficient utilization or storage of primary products of photosynthesis is well documented to contribute to source/sink imbalances and photoinhibitory outcomes in a many green lineage species ( Adams et al., 2014 ; Santos-Merino et al., 2021a ). Conversely, remobilization of carbon from storage polymers ( i.e. , glycogen) is essential to rapidly replenish photosynthetic carbohydrate intermediates to efficiently ‘reboot’ the Calvin-Benson-Bassham (CBB) cycle during environmental fluctuations, such as dark to light transitions ( Makowka et al., 2020 ; Shinde et al., 2020 ). Previously, Δ rpaA mutants have been reported to inefficiently partition carbon towards glycogen under phototrophic conditions, despite the high levels of glycogen biosynthesis enzymes detected in this strain ( Puszynska and O’Shea, 2017 ), and this may contribute to redox stress under light/dark cycles ( Diamond et al., 2017 ). Here, we report that provided abundant organic carbon under mixotrophic growth, Δ rpaA mutants hyperaccumulate glycogen to levels above the typical physiological range ( Figure 4 ). Our results demonstrate that Δ rpaA mutants are not deficient in glycogen synthesis capacity, and therefore may fail to provision adequate glycogen under phototrophic conditions due to other limitations ( e.g. , misregulation of carbon partitioning and/or decreased CO 2 fixation rates). Multiple lines of evidence therefore suggest that Δ rpaA mutants are unable to flexibly adapt the flux of carbon to match metabolic needs with changing environmental conditions. As a whole, Δ rpaA mutants may be especially overreactive to ROS due to the roles for RpaA in regulating metabolic redox pools, detoxifying ROS, and/or controlling alternative electron transport pathways that can remove excess reductant on the pETC. Beyond their direct activity as potent oxidative compounds, ROS can also function as important signals to mediate a variety of cellular responses in photosynthetic organisms ( Mullineaux et al., 2018 ). For example, in plants ROS have been proposed to be a primary signal of overreduction of the PQ pool ( Mubarakshina and Ivanov, 2010 ), and H 2 O 2 specifically has important signaling roles in a variety of plant processes including senescence, flowering, and biotic stress ( Niu and Liao, 2016 ). In algae, H 2 O 2 -responsive pathways are specifically involved in the regulation of CCM, where it is proposed that ROS signals may serve as a proxy signal of imbalanced photosynthesis: activation of CCM processes may thereby help to rebalance the system and consequently suppresses further ROS formation ( Choi et al., 2022 ). Of particular relevance to this study, it was recently reported that H 2 O 2 (derived from photorespiratory processes or from direct chemical addition) can directly modulate the pyrenoid, inducing a more robust aggregation of rubisco into the pyrenoid core as well as a thicker and more defined pyrenoid starch sheath ( Neofotis et al., 2021 ). Despite the lack of conservation in many of the structural components between carboxysomes and the algal pyrenoid, our data suggests that H 2 O 2 derived from an overreduced pETC as a functionally conserved signal to coordinate downstream CCM (re)organization in cyanobacteria as well. In natural environments, cyanobacteria experience changing environmental conditions ( e.g. , high light, nutrient limitations) and must adaptively modulate core machinery to maintain robust photosynthesis ( Stockenreiter et al., 2021 ). Cyanobacteria have evolved the ability to homeostatically regulate the redox status of its PQ pool, instead of functioning as a source of a regulatory signal ( Schuurmans et al., 2014 ). Due to its central location between the two photosystems, the redox status of the PQ pool plays a pivotal role in sensing cellular status and in regulating photosynthetic capacity ( Mullineaux and Allen, 1990 ). Particularly, the redox status of the PQ pool have regulatory roles related to photosystem composition ( Fujita et al., 1987 ), state transitions ( Mullineaux and Allen, 1990 ), and redistribution of respiratory complexes ( Liu et al., 2012 ). Collectively, our data points to an additional regulatory role of the PQ pool as connected to the integrity and/or remodeling of the carbon fixation machinery under transitional periods where there may be a mismatch between the light and dark reactions. This control appears to involve RpaA, revealing a previously underappreciated function for the multi-layered regulation of both PQ and RpaA towards achieving energy balance in cyanobacteria. 6. Funding This work was primarily supported by the Department of Energy and Basic Energy Sciences Division (Grant: DE-FG02-91ER20021). Acknowledgments The Attune Cytpix, located in the MSU Flow Cytometry Core Facility, is supported by the Equipment Grants Program, award no. 2022-70410-38419, from the U.S. Department of Agriculture (USDA) National Institute of Food and Agriculture (NIFA). The authors thank Dr. Alicia Withrow and the Michigan State University Center for Advanced Microscopy for assistance with electron microscopy. References ↵ Abramson , B.W. , Kachel , B. , Kramer , D.M. , and Ducat , D.C . ( 2016 ). Increased photochemical efficiency in cyanobacteria via an engineered sucrose sink . Plant & cell physiology 57 , 2451 – 2460 . OpenUrl CrossRef PubMed ↵ Adams , W.W. , Muller , O. , Cohu , C.M. , and Demmig-Adams , B. ( 2014 ). Photosystem II efficiency and non-photochemical fluorescence quenching in the context of source-sink balance . In Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria , B. Demmig-Adams , G. Garab , W. 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