A functional cyanophage thioredoxin increases competitive phage fitness

A functional cyanophage thioredoxin increases competitive phage fitness | 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 A functional cyanophage thioredoxin increases competitive phage fitness View ORCID Profile Ran Tahan , Dror Shitrit , View ORCID Profile Debbie Lindell doi: https://doi.org/10.1101/2025.10.05.680603 Ran Tahan Faculty of Biology, Technion – Israel Institute of Technology , Haifa 320003, Israel Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Ran Tahan Dror Shitrit Faculty of Biology, Technion – Israel Institute of Technology , Haifa 320003, Israel Find this author on Google Scholar Find this author on PubMed Search for this author on this site Debbie Lindell Faculty of Biology, Technion – Israel Institute of Technology , Haifa 320003, Israel Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Debbie Lindell For correspondence: dlindell{at}technion.ac.il Abstract Full Text Info/History Metrics Preview PDF Abstract Thioredoxins are ubiquitous redox proteins that are found in all domains of life. These conserved proteins are also found in many phages, including marine cyanophages that infect the ecologically important marine cyanobacteria. However, their role in phage infection is not known. Cyanophages also carry many small genes lacking homology to known functional domains. Whether these have a functional role or not remains unknown. Here, we explore the distribution and role of a cyanophage thioredoxin ( trxA ), and that of a small gene directly downstream of it ( g26 ), in phage infection. For this we used the T7-like cyanophage, Syn5, which infects an open-ocean marine Synechococcus strain, WH8109. We found that thioredoxin genes are common in phage genomes, including in cyanophages. The g26 gene, however, is restricted in it distribution to the cyanophages. The cyanophage thioredoxin is catalytically active and it increases phage DNA replication, progeny production and competitive fitness. It also negatively impacts host growth. The g26 gene product is translationally coupled to, and thus dependent on, translation of the thioredoxin gene. This gene itself significantly increases phage virulence and fitness, yet reduces burst size. Our findings demonstrate that cyanophage thioredoxins impact phage fitness and infection physiology and that small viral genes with no homology to known genes can play an important role in the infection process. These findings provide insights into the importance of unusual genes in phage genomes and show that they are likely to play an important role in the interactions between abundant cyanobacteria and cyanophages in ocean ecosystems. Introduction Marine picocyanobacteria of the genera Synechococcus and Prochlorococcus are the numerically dominant phytoplankton in the ocean [ 1 , 2 ]. They are estimated to be responsible for approximately 25% of ocean primary production [ 3 ]. Cyanobacteria coexist with a variety of phages that can infect them (cyanophages). Marine cyanophages, are dsDNA lytic phages belonging to the class of Caudoviricetes [ 4 – 6 ]. The most abundant cyanophage groups are the T4-like and the T7-like cyanophages of the Kyanoviridae family and Autographivirales order respectively [ 7 – 10 ], although other cyanophages are also known, such as a variety of siphoviruses, TIM5-likes, and others. The T7-like cyanophages can be divided into three major clades - A, B and C which differ in their abundance, infection physiology and gene content [ 11 – 14 ]. A common feature of cyanophage genomes are host-like auxiliary metabolic genes (AMG) [ 15 , 16 , 4 , 13 , 17 – 19 ]. These genes, often acquired from their hosts in the evolutionary past, are thought to reshape host cell metabolism for the benefit of the phage [ 15 , 16 , 20 ]. Among others, these genes include photosynthesis genes, genes involved in carbon metabolism, and nutrient utilization genes. One of the AMGs found in cyanophages is thioredoxin [ 16 , 13 , 21 , 22 ]. Thioredoxins are ubiquitous redox enzymes, found in all domains of life. [ 23 , 24 ]. In bacteria, thioredoxins function as part of the cellular oxidative stress response by reducing oxidized proteins and directly scavenging reactive oxygen species (ROS) [ 24 ]. They also regulate the activities of enzymes and transcription factors by thiol switches which link the activity of a protein to the redox state of the cell [ 24 ]. In cyanobacteria, thioredoxin is essential for growth [ 25 ], linking the redox state of the photosynthetic electron transfer chain to transcription factors [ 26 , 27 ] and metabolic enzymes [ 28 ]. Whether cyanophage thioredoxins function in a similar manner to that of cells is not known, nor is it known whether they are important for cyanophage infection. The genomes of phages from the marine environment contain a fair number of small genes with no recognizable functional domains. These can make up between 40-90% of their genomes and are particularly high for environmental phages that are not model research systems [ 29 , 30 ]. However, it is unclear whether such genes are functional and their role in phage infection has not been tested. In this study, we explore the role of a cyanophage thioredoxin gene ( trxA ) and a small gene directly downstream of it ( g26 ) which overlaps with the trxA gene. Our model system is the clade A T7-like cyanophage, Syn5, that infects Synechococcus WH8109, belonging to the common open-ocean clade II of Synechococcus subcluser 5.1. We found that the gene is a functional thioredoxin and that it is translationally coupled to gp26. Together these two genes serve to increase the fitness of the Syn5 cyanophage, with each gene contributing to this fitness improvement differently. The thioredoxin gene increased phage DNA replication and progeny production while g26 increased the virulence of the cyanophage. These findings demonstrate that cyanophages carry functional thioredoxin genes that impact the infection process. They further show that small genes of unknown function can have a significant impact on phage infection and phage fitness. Results and Discussion In this study we set out to study the role of a putative thioredoxin gene in Syn5, a T7-like cyanophage that infects Synechococcus sp. strain WH8109 ( Synechococcus WH8109 from herein). This gene is located amongst DNA replication genes in the Syn5 genome[ 21 ]. Directly downstream of the trxA gene is a small gene with no recognizable functional domains. Interestingly, the stop codon of the trxA gene overlaps the start codon of the g26 gene and may thus be functionally coupled. We began by first assessing the prevalence of both the putative thioredoxin gene ( trxA ) and the g26 in phage genomes. We then assessed whether the putative cyanophage thioredoxin has redox activity and whether it impacts host growth. Using a genetic manipulation system we deleted each of the trxA and g26 genes from the Syn5 genome and explored their effect on cyanophage fitness and the infection cycle. Thioredoxin is widespread in phages We began by assessing how widespread thioredoxin genes are in phages. A gene was considered to be a putative thioredoxin based on sequence homology and the presence of a thioredoxin catalytic site, which is characterized by a conserved sequence consisting of two cysteine amino acids (CGPC). We found that many different phages, infecting diverse hosts, carry putative thioredoxin genes ( Fig. 1 ). These include phages that infect strains of Synechococcus , Escherichia , Bacillus , Acinetobacter and more. The phages, and cyanophages in particular, carrying the genes belong to a variety of phage families. Interestingly, the putative thioredoxin is present only in clade A T7-like cyanophages. Download figure Open in new tab Figure 1: Gene tree of phage and bacterial thioredoxin. Maximum-likelihood tree for TrxA was inferred from 100 amino acid positions. Bootstrap values equal or greater than 0.5 are shown at branch nodes. Genes from T7-like cyanophages are shown in blue, Synechococcus WH8109 genes are shown in pink, archaea and archaea virus are shown in orange and phage genes within the bacteria clade are shown in purple. We constructed a thioredoxin gene tree to study the diversity and phylogenetic relationships between bacterial and phage thioredoxins. We found that nearly all putative phage thioredoxins cluster separately from bacterial thioredoxins, including all of the cyanophage thioredoxin genes ( Fig. 1 ). This suggests that phages acquired the gene from bacteria in the distant past and that the phage and bacterial thioredoxins have evolved separately over a long period of time. There are, however, four phages with genes that cluster within bacterial clades in close proximity to their respective hosts. This suggests more recent and independent acquisitions of these thioredoxin genes from their bacterial hosts. The widespread occurrence of thioredoxin genes in phages from different families and infecting a range of bacterial taxa, whether acquired in the distant past or more recently, suggests that thioredoxins confer an evolutionary advantage for diverse phages. The predicted structure of phage thioredoxins is very similar to that of bacterial thioredoxins ( Fig. 2A ). In particular, the sequence, position and conformation of the catalytic site is conserved between the bacterial and phage thioredoxins ( Fig. 2A , Fig. S1). However, the majority of the phage thioredoxins differ at the N-terminus, lacking approximately 20 aa that form a beta strand and alpha helix in bacterial thioredoxins ( Fig. 2A , Fig. S1). In contrast, the phage thioredoxins that were likely recently acquired from their hosts still contain the bacterial-like thioredoxin N-terminus ( Fig. 2B ). The overall structural similarity yet differences at the N-terminus raises the question as to whether phage thioredoxins have similar or different functions to those of bacterial thioredoxins. Download figure Open in new tab Figure 2: Alignment of predicted structures of phage and bacterial thioredoxins. AlphaFold3 predicted structural alignment of E. coli K12 (blue) and Syn5 (red) thioredoxins ( A ) and of Burkholderia cenocepacia (purple) and Burkholderia phage BcepSauron (green) thioredoxins. Cysteine residues of the active sites are shown in grey (carbon) and yellow (sulfur). Phage or bac are used to indicate the location of phage or bacterial part of the protein. The g26 gene is directly downstream of the putative thioredoxin gene in our model Syn5 phage. Therefore, we investigated the prevalence of g26 in the non-redundant database. Different to thioredoxin, we found this gene to be limited to cyanophages. It is, however, very common in T4-like cyanophage genomes. It is also present in some T7-like cyanopodoviruses, being found in two of four T7-like cyanophages from clade A that carry a thioredoxin gene (Table S2) and has an overlapping start codon in both cases. The gene is also present in TIM5-like cyanomyoviruses and P-SSP2-like cyanosiphoviruses (Table S2). Interestingly, we found a homologue of this gene in a complete PSS2-like cyanopsiphovirus putative prophage that we detected in the genome of Synechococcus sp. SYN20. The restricted presence of this gene to cyanophages, while being quite prevalent among them, raises the question as to whether it has a function during infection of cyanobacteria. Cyanophage thioredoxin is catalytically active and negatively impacts host growth The structural differences between the phage and bacterial thioredoxin genes raised the question as to whether the phage copies of the gene have a similar function to those in bacteria. We assessed whether the Syn5 thioredoxin gene displays the typical catalytic activity of cellular thioredoxins using an in vitro assay. For this, we expressed the Syn5 thioredoxin in E. coli cells lacking the trxA gene using an inducible expression vector and measured redox activity in the cell lysate. We found that Syn5 thioredoxin successfully reduced the fluorescently labeled substrate ( Fig. 3A ), indicating that it is catalytically active. We also tested whether this catalytic activity is facilitated by the classical CGPC thioredoxin active site conserved in the phages. For this we exchanged the two cysteines, known to be essential for redox activity, to serine residues, following Huber et al. (1986). This abolished the catalytic activity ( Fig. 3B ), indicating that the CGPC of Syn5 thioredoxin is the active sites of the protein and that the cysteines are essential for the redox reaction. Taken together, these results indicate that Syn5 carries a functional thioredoxin gene with a conserved catalytic site despite the lack of the N-terminus conserved in cellular thioredoxins. Download figure Open in new tab Figure 3: Syn5 thioredoxin is catalytically active. Comparison of Syn5 wild type (WT) and mutated thioredoxin catalytic activity in induced (purple) and non-induced control (black) cultures of E.coli Δ trxA cell lysates carrying inducible expression vectors of thioredoxin ( A ) catalytically inactive thioredoxin ( B ) or empty vector ( C ). Average and standard deviation of 3-4 biological replicates. The effect of the inducer was tested using the repeated measures ANOVA. P-values are represented by: *** p-value<0.001. Next, we wondered if the catalytic activity of cyanophage thioredoxin affects the metabolism of the host. For this purpose, we attempted to express the cyanophage thioredoxin in the host from the previously described pDS-proCAT replicative plasmid [ 67 ]. However, we were unable to insert a plasmid expressing the trxA gene into the host. In contrast, we were successful in inserting an inactivated version, in which we inserted a stop codon and a frameshift into the gene. These results suggested that Syn5 thioredoxin is toxic to the host. To directly test the effect of the cyanophage thioredoxin on the metabolism of the host we adapted a theophylline-dependent riboswitch inducible expression system previously used in different cyanobacterial species [ 54 , 55 ]. We expressed catalytically active and inactive thioredoxin and measured their effect on host growth ( Fig. 4A-C ). We found that induction of the catalytically active thioredoxin resulted in a decline in host growth ( Fig. 4A ). In contrast, host growth was not affected by expression of the catalytically inactive thioredoxin ( Fig. 4B ). This indicates that catalytic activity of the Syn5 thioredoxin impacts host physiology. Importantly, this result indicates that Syn5 thioredoxin is detrimental to the host, and therefore affects host cell metabolism. Download figure Open in new tab Figure 4: Catalytic activity of Syn5 thioredoxin affects host physiology. Effect of wild-type thioredoxin ( A ), catalytically inactive thioredoxin ( B ), or empty vector ( C ) expression on host growth. Average and standard deviation of 5 biological replicates of Syn5 wild type (WT) and mutated induced (purple) and non-induced control (black) cultures of Synechoccucus WH8109 carrying theophylline-dependent inducible expression vectors. The effect of the inducer was tested using the repeated measures ANOVA. P-values are represented by: *** p-value<0.001. Thioredoxin and gp26 increase competitive fitness The prevalence of thioredoxin and gp26 in phages led us to hypothesize that these genes play a functional role in the infection of the cyanobacterial host and increase phage fitness. To test this, we used our recently developed genetic manipulation method, REEP (Shitrit et al. 2022), to delete each of these genes from the Syn5 phage and to compare their fitness and infection properties to that of the wild-type phage. Since the stop codon of the trxA gene overlaps the start codon of the g26 gene, the translation of gp26 may be dependent on translation of thioredoxin. Thus, the deletion of trxA may result in the absence of both thioredoxin and the gp26 protein. To test this, we measured peptide levels of the two proteins in the Δ trxA mutant and compared them to those in the wild-type phage. Neither thioredoxin nor gp26 peptides were detected in the Δ trxA mutant phage (Fig. S2B). However, g26 transcripts were at a similar level in the Δ trxA mutant and the wild-type phages (Fig. S2A). Thus, while transcription of g26 is not affected in Δ trxA mutant, this protein was not translated, indicated that the translation of gp26 is coupled to that of thioredoxin. Thus, the Δ trxA cyanophage mutant is a functional double mutant lacking both the thioredoxin and the gp26 proteins. We will thus refer to this mutant as Δ trxA /gp26 S , with the “s” referring to silencing. We now turned to assessing the importance of these two genes on phage fitness, both together and individually. To do this we compared the mutant phages to that of the wild-type phage and to each other. The difference between the Δ trxA /gp26 S and Δg26 mutants provides insight into the effect of thioredoxin alone on phage fitness. A comparison of the ΔtrxA /gp26 S and the wild-type phage revealed ∼6-fold wild-type than mutant cyanophages ( Fig. 5A ). This result indicates that both genes together contribute significantly to competitive phage fitness. Download figure Open in new tab Figure 5: Direct competition of Syn5 strains infecting the same host culture. Competitive phage fitness between the wild-type and the ΔtrxA /gp26 S mutant ( A ), the wild-type and the Δ g26 mutant ( B ), and the ΔtrxA /gp26 S and the Δ g26 mutants ( C ). The wild-type strain is shown in black, ΔtrxA /gp26 S in green and Δ g26 in red. Average and standard deviation of 4 biological replicates. P-values of paired, two tailed student t-test are represented by: *** p-value<0.001. n.s., no significance. Next, we compared the Δg26 mutant and the wild-type phage and found a ∼2-fold more of the wild-type phage ( Fig. 5B ). Therefore, gp26 contributes to competitive phage fitness independently of thioredoxin. Finally, we compared the ΔtrxA /gp26 S and Δg26 mutants and found ∼5-fold more of the Δg26 mutant ( Fig. 5C ), indicating that the phage thioredoxin has a distinct contribution to competitive fitness. The greater fold difference between the wild-type and ΔtrxA /gp26 S mutant than between the wild-type and the Δg26 mutant suggests that trxA contributes more to phage fitness than g26 . Thus, while trxA and g26 both have a distinct contribution to competitive phage fitness, the effect of trxA is greater. Thioredoxin and gp26 alter phage infection properties differently Phage fitness is impacted by different properties of phage infection such as the rate of adsorption, the length of time to produce new phage progeny (the latent period), the number of infective progeny produced per cell (the burst-size) and the percentage of cells lysed during infection (the virulence) [ 68 , 69 ]. Thus, we set out to elucidate the impact of trxA and g26 genes on phage infection properties using the same comparative as described above. All experiments were carried out with an infective phage to host ratio of 1 at a cell and phage concentration of 5 x10 7 cells and phages per ml. To test the effect of the genes on the rate of adsorption and the length of the latent period, we used phage growth curve experiments. We found that neither gene changed the rate of adsorption or the length of the latent period ( Fig. 6A ). Download figure Open in new tab Figure 6: Effects of Syn5 trxA and g26 mutants on latent period and infection cycle. Comparison of adsorption and the latent period ( A ) and phage progeny production ( B - D ) between Syn5 wild-type ΔtrxA /gp26 S mutant ( B ), the ΔtrxA /gp26 S and the Δ g26 mutants ( C ), and the wild-type and Δ g26 mutant ( D ) during infection. Average and standard deviation of 3-4 biological replicates. P-values of significantly different PFU at a given time point (repeated measures ANOVA), are represented by: *p-value<0.05,*** p-value<0.001. Next, we quantified the virulence of our phage strains. The virulence of the wild-type phage was ∼40%, indicating that 40% of the cells in the culture were lysed under the conditions used ( Fig. 7A ). In comparison, the virulence of the Δg26 mutant was ∼34% ( Fig. 7A ), significantly lower than that of the wild-type by 6% ( Fig. 7A,B ), resulting in an increase of ∼15% in the number of cells that are lysed due to g26 in the wild-type phage. However, no additional difference was observed between the Δ g26 and ΔtrxA /gp26 S strains ( Fig. 7A,B ), indicating that trxA has no significant contribution to phage virulence in the wild-type phage. Download figure Open in new tab Figure 7: Effects of Syn5 trxA and g26 mutants on virulence and burst size. Virulence ( A ), burst size ( C ) and the contribution of trxA and g26 to virulence ( B ) and burst size ( D ) of Syn5 wild type (black), ΔtrxA /gp26 S (green) and Δ g26 (red) strains. Average and standard deviation of 8 biological replicates. P-values of significantly different virulence or burst size (Wilcoxon signed rank test or paired t-test) are represented by: * p-value<0.05, ** p-value<0.01 *** p-value<0.001. Strains without significant differences are represented by n.s. (no significance). Next, we tested the effect of trxA and g26 on phage progeny production. For this purpose, we assessed the burst size, of our phage strains using a population level burst size assay [ 70 ]. The burst size of the wild-type phage was ∼157 phages per cell ( Fig. 7C ). Intriguingly, the burst size of the Δg26 mutant was significantly more than that (∼217 phages per cell) ( Fig. 7D ), indicating that g26 reduces the burst size of the Syn5 wild-type phage by ∼60 phages per cell, decreasing Syn5 progeny production by ∼28% relative to the wild-type phage. However, the burst size of ΔtrxA /gp26 S was similar to that of the wild-type phage ( Fig. 7C ) suggesting that the negative effect of g26 on phage progeny production is mitigated by a positive effect of the trxA gene. Indeed, a direct comparison of the ΔtrxA /gp26 S and Δ g26 mutants, showed that the lack of thioredoxin gene resulted in a lower burst size, i.e. that trxA gene in the wild-type phage increases the burst size by ∼51 phages per cell ( Fig. 7D ), increasing progeny production by ∼33% relative to the wild-type phage. Therefore, our results indicate that g26 decreases, while trxA increases, phage progeny production by a comparable amount, each cancelling the effect of the other gene on burst size. A summary of all the infection property results indicates that trxA had a positive effect on burst size but did not alter other infection properties. In contrast, our results show that g26 both reduced the burst size and increased phage virulence. The increased burst size due to thioredoxin is negated by the negative effect of g26 on this infection property, such that the g26 virulence effect appears to be the major contributor to the increased fitness conferred by both genes together. This indicates that virulence is an important factor affecting phage fitness. In fact, the fitness of the Δ g26 mutant is lower even when burst size is increased ( Fig. 5B ), indicates that virulence can be an extremely important properties influencing phage fitness. Cyanophage thioredoxin increases phage genome replication The Syn5 thioredoxin gene and g26 are located within the gene cluster responsible for phage DNA replication [ 21 ]. Therefore, we hypothesized that one or both genes may contribute to phage genome replication. To test this hypothesis, we compared intracellular phage DNA during infection between the wild-type, Δ g26 and ΔtrxA /gp26 S mutant Syn5 phages. We found that intracellular DNA was lower in the ΔtrxA /gp26 S mutant compared to both the wild-type phage and the Δ g26 mutant phage ( Fig. 8A,B ). Moreover, there was no difference in genome replication between the wild-type and Δ g26 mutant phages ( Fig. 8C ). Thus, the trxA gene is response for the increased phage genome replication out of the two genes affected in the ΔtrxA /gp26 S mutant. These findings further suggest that the Syn5 thioredoxin increases phage progeny production by increasing phage genome replication. Download figure Open in new tab Figure 8: The effect of trxA and g26 on phage DNA replication. Two-way comparisons of intracellular DNA replication during infection of Syn5 wild type (black) ( A, C ), ΔtrxA /gp26 S (green) ( A, B) and Δ g26 (red) ( B, C ). Average and standard deviation of 4 biological replicates. P-values of significantly different intracellular phage DNA copy numbers at a given time point (paired, two tailed student t-test), are represented by: * p-value<0.05. We and others have hypothesized that the phage-encoded thioredoxin could have a dual function in Syn5. One possibility is that it reduces the phages ribonucleotide reductase [ 21 ], facilitating the increase the recycling of host ribonucleotides to dexoyribonucleotides. A higher concentration of deoxyribonucleotides would increase the availability of nucleotide substrates for the phage DNA polymerase. In turn, this could lead to an increase in the rate of genome replication and therefore progeny production. Alternatively, the mode of function of the phage thioredoxin may be independent of its redox activity and function through direct binding to DNA polymerase (DNAP) as a processivity factor [ 21 ]. This would be similar to the function of the E. coli thioredoxin during T7 infection, in which the host-encoded thioredoxin binds the phage DNA polymerase, greatly increasing its processivity [ 71 – 73 ]. Since this process is essential for T7 genome replication and phage progeny production [ 73 ], it is possible that a T7-like cyanophage like Syn5 would use its own thioredoxin as a processivity factor for DNA replication. We next explored whether the Syn5 thioredoxin might function as a processivity factor for the phage DNA polymerase. We evaluated the ability of T7 to infect a deletion mutant of E. coli trxA ( E. coli Δ trxA ) expressing Syn5 or Synechococcus WH8109 thioredoxins and found that neither restored T7 infectivity. This provides some support for the possibility that the cyanophage thioredoxin does not function as a processivity factor. However, such negative findings could also result from other factors such as the lack of compatibility between the cyanophage thioredoxin and the T7 DNA polymerase. Next, we used AlphaFold3 [ 44 ] to model the predicted structure of phage DNA polymerases with host and phage thioredoxins. First, we modeled the predicted structure of T7 DNAP with E. coli thioredoxin and found that it resembles the experimentally determined structure of their complex ( Fig. 9A ) [ 74 ], providing confidence in the ability of AlphaFold3 to reconstruct such an interaction when it occurs. We then moved to model the structure of Syn5 cyanophage DNA polymerase with thioredoxins coded by two different genes from the Synechococcus WH8109 host and the Syn5 cyanophage thioredoxin. We found that both Synechococcus thioredoxins are predicted to bind the Syn5 DNA polymerase similarly to the way that the E. coli thioredoxin binds to the DNA polymerase of T7 ( Fig. 9A,B ,C). This suggests that host thioredoxins from marine cyanobacteria could act as processivity factors for T7-like cyanophage DNA polymerases. In stark contrast, the Syn5 cyanophage thioredoxin is predicted to bind at a completely different site on the cyanophage DNA polymerase ( Fig. 9D ). Moreover, the cyanophage thioredoxin had a ∼4 times lower interface predicted template modelling (ipTM) score compared to the other models, suggesting a lower likelihood of complex formation. These findings argue against the cyanophage thioredoxin functioning as a processivity factor for the cyanophage DNA polymerase. Thus, we consider the possibility that the cyanophage thioredoxin reduces the phage ribonucleotide reductase, or some other redox dependent function, to be the more likely mode of action of the Syn5 cyanophage thioredoxin in enhancing phage genome replication. Download figure Open in new tab Figure 9: AlphaFold3 predicted structures of phage DNA polymerase interacting with different thioredoxins. ( A ) Predicted fold of T7 DNA polymerase (brown) with E. coli K12 thioredoxin (orange), ( B ) Syn5 DNA polymerase (purple) with Synechococcus WH8109 thioredoxin 1 (blue), ( C ) Syn5 DNA polymerase (purple) with Synechococcus WH8109 thioredoxin 2 (teal) and ( D ) Syn5 DNA polymerase (purple) with Syn5 thioredoxin (green). Predicted template modelling (pTM) score and interface predicted template modelling (ipTM) are provided for each structure. GenBank accession numbers are AHF64381.1 and AHF63033.1 for Synechococcus WH8109 thioredoxin 1 and 2 respectively. Materials and methods Cyanobacterial growth and cyanophage propagation Synechococcus sp. WH8109 was grown in artificial sea water medium (ASW) with NO 3 - as the nitrogen source [ 31 ]. The light intensity was 20-25 µmol photons m -2 sec -1 under a 14/10 day/night cycle and the growth temperature was 21 +/-1°C. Growth was monitored using chlorophyll a fluorescence of the cells, with excitation at 440±20 nm and emission detected at 680±20 nm using a Synergy Mx microplate reader (BioTek, Winooski, CA, USA). Synechococcus WH8109 colonies and lawns were grown using the pour plate method with ASW+NO 3 - +SO 3 2- medium supplemented with 0.28% low melting point agarose [ 32 , 33 ] (UltraPure, Invitrogen, Carlsbad, CA, USA). In order to achieve high plating efficiencies, an antibiotic resistant ‘helper’ bacterium, Alteromonas sp. strain EZ55, was added to the medium when plating Synechococcus WH8109 after conjugation to reduce oxidative stress (see below) [ 34 ]. Syn5 lysates were prepared by infecting exponentially growing Synechococcus WH8109 at a concentration of 7*10 7 -1*10 8 cells/mL with a preexisting lysate. After 24 hours the resulting lysate was filtered over a 0.22µm Millex GV syringe filter (Millipore, Cork, Ireland) to remove cellular debris and was stored at 4°C in glass tubes. Plaque assay The plaque assay was used to quantify the number of infective phages (plaque forming units - PFU). Filtered lysates were serially diluted, mixed with Synechococcus WH8109 culture in a petri dish, pour-plated and grown as described above to produce plaques but without the addition of the ‘helper’ bacterium. Phylogenetics Gene trees were built for the thioredoxin and a translationally coupled protein, gp26, using the NGPhylogeny.fr web server [ 35 ]. Protein sequences were identified by sequence homology using Position-Specific Iterated BLAST (PSI-BLAST) [ 36 ] and retrieved from the National Center for Biotechnology Information (NCBI) database. Thioredoxin homologues included in our analyses all contained the canonical active cite CGPC or CAPC [ 37 ], except for two T7-like cyanophages that had …. in the appropriate position. Protein sequences were aligned using the MAFFT program [ 38 ]. Aligned sequences were examined before and after alignment curation using TrimAI [ 39 ]. For thioredoxin sequences a gap threshold (the fraction of sequences with a gap allowed) of 0.3 was set to avoid losing a bacterial N-terminal amino acid sequence not found in phages. Maximum likelihood and neighbor-joining analyses were performed using the PhyML+SMS and FastME programs [ 40 , 41 ]. Maximum likelihood and neighbor-joining topologies were compared using TreeViewer [ 42 ] to ensure overall similarity in topologies. To obtain confidence estimates for the inferred tree topology, 500 and 400 bootstrap resamplings were performed for thioredoxin and gp26 respectively. The maximum likelihood tree topology was drawn and edited using the iTOL software [ 43 ]. Structural predictions and alignment Protein structure models were predicted using AlphaFold 3 web server [ 44 ]. Structures were uploaded and visualized using the ChimeraX software [ 45 ]. Structural alignment was performed using the matchmaker function with default parameters. Vector construction and bacterial conjugation Vectors were constructed for expression of trxA in E. coli K12 BW25113 and Synechococcus WH8109, and for the cloning of recombination templates for gene inactivation (Table S1). The Syn5 thioredoxin gene was codon optimized for expression in E. coli K12 and synthesized (Azenta life sciences) with BspHI and PmeI restriction sites. This construct was used to introduce a CGPC → SGPS mutation in Syn5 thioredoxin catalytic site using the PCR overlap extension method [ 46 ]. Two PCR fragments were generated by amplifying the codon optimized Syn5 thioredoxin. The mutation was introduced into the overlapping region of the PCR fragments placed in the active site region of Syn5 thioredoxin. Constructs were ligated into the pBAD plasmid (pBAD TOPO TA Expression Kit, Thermo Fisher Scientific) using the NcoI and PmeI restriction sites, made possible by the compatibility of BspHI and NcoI overhangs following restriction enzyme digestion. The vector was and inserted into E. coli K12 BW25113 Δ trxA cells (Keio collection strain ECK3773 [ 47 ]) lacking the endogenous thioredoxin gene by electroporation. Selection of transformed colonies was done on LB-agar plates containing 50 µg/mL ampicillin. Successful transformation of the plasmids was verified by colony PCR [ 48 ] and Sanger sequencing of the inserted region. All vectors used in Synechococcus WH8109 were based on pDS-proCAT, a broad host range replicative vector optimized for cyanobacterial chloramphenicol (Cm) resistance [ 49 ]. The different inserts were constructed using the PCR overlap extension method [ 46 ]. The cloning of the different inserts (Table S1) was performed by digesting the pRL1342-proCAT backbone and the insert at BamHI and BsiWI restriction sites. Conjugation was performed using the method described by Brahamsha (1996) with modifications as described previously [ 49 ]. E. coli S17.1 carrying the λpir gene [ 52 ] was used as a conjugative donor strain for vectors into Synechococcus WH8109. Plasmids were inserted into E. coli S17.1 by electroporation. Successful conjugation of Synechococcus WH8109 was verified by colony PCR [ 48 ] and Sanger sequencing of the inserted region of the conjugated vector. Successfully conjugated colonies were grown in ASW+NO 3 - medium containing chloramphenicol for selection of plasmid-carrying cyanobacteria. Thioredoxin activity assays We used the thioredoxin fluorometric activity assay kit (Cayman chemical) to measure the reduction of eosin-labeled insulin substrate by thioredoxin in a cell lysate over time. We expressed Syn5 thioredoxin in E. coli lacking the endogenous thioredoxin gene, using an inducible expression plasmid. Protein expression was induced using a 0.1% (v/v) L-arabinose (Sigma-Aldrich). Induced cultures were grown with vigorous shaking overnight (15 hours) at 37°C. Cultures were centrifuged to a pellet for 5 min, 5467g at 4°C, washed twice with ice-cold 100mM Tris-HCl, 1mM EDTA pH7.5 supplemented with protease inhibitor (Cayman chemical), resuspended in 0.5mL and added to a tube containing ∼0.5mL of ice-cold, acid-washed glass beads ≤106μm (Sigma-Aldrich). The culture was lysed using a bead-beater (Mini-BeadBeater, Biospec) for 1 minute (3450 oscillations/min) followed by 1 minute cooling on ice. Cell lysate was cleared by centrifugation for 3 minutes, 20817g at 4°C and the supernatant was used immediately for the thioredoxin activity assay. Thioredoxin activity in the cell lysate was tested by measuring the reduction of eosin-label insulin. The reduction of disulfide bonds in the labeled insulin by thioredoxin [ 53 ] releases the eosin label and causes an increase in eosin fluorescence (excitation at 520nm and emission at 560nm). Thioredoxin is oxidized in this reaction and is recycled to the reduced state by thioredoxin reductase using NADPH. To ensure that similar protein concentrations were used in the activity assay, we measured protein concentrations using the Pierce™ BCA Protein Assay Kit (Thermo Scientific). Inducible expression system Here we adapted an inducible expression system at the stage of translation for marine Synechococcus following that used previously for freshwater cyanobacteria [ 54 , 55 ]. This system utilizes a theophylline-dependent riboswitch of type E* (also known as F) and was engineered to consist of the Prochlorococcus MED4 rnpB promoter constitutively transcribing the riboswitch and the downstream gene of interest in the pDS-proCAT self-replicating plasmid [ 49 ]. Gene expression was induced by adding ASW-dissolved theophylline to an exponentially growing Synechococcus culture containing the expression plasmid. To evaluate the timing and changes in the level of gene expression from the theophylline-dependent riboswitch system, we employed the β-glucuronidase (GusA) reporter gene assay [ 56 ]. We used the GusA assay previously described before for the filamentous cyanobacterium Fremyella diplosiphon UTEX 481 [ 57 , 58 ]. We induced GusA expression in Synechococcus WH8109 using 0-0.3mM theophylline and measured GusA catalytic activity using a quantitative assay at several time points between 0.5 and 72.5 hours post induction (Fig. S3). Cells were harvested by centrifugation in room temperature at 5467g for 5 minutes. Cell pellets were resuspended in ice-cold GUS assay buffer (1mM EDTA, 50mM NaPO 4 pH7) containing 0.001% SDS and 6.25μg/mL chloramphenicol, then lysed by bead-beating and cleared by centrifugation as described for the thioredoxin activity assay. 20μL of cell lysates were mixed with 180μL GUS assay buffer containing 1.125mM 4-Nirophenyl β-D-glucuronide (PNPG, Sigma-Aldrich). The degradation of PNPG by GusA, producing a yellow pigment, was measured by absorbance at 410nm every 2 minutes for 30 minutes using a Synergy Mx microplate reader (BioTek, Winooski, CA, USA). Protein concentration in the cell lysates was measured using the BCA protein determination kit (Thermo Fisher Scientific) according to the manufacturer’s protocol. GusA activity was quantified as nmol of PNG product per mg of protein per minute. Four biological replicates were performed for each theophylline concentration. Based on the GusA assay findings, we used theophylline at a final concentration of 0.1mM to test the effect of expression of different versions of the Syn5 trxA gene on the growth of Synechococcus WH8109. Induced cultures were compared to non-induced cultures to determine the effect of thioredoxin expression on bacterial growth. To control for the potential effect of theophylline on growth, expression experiments included an empty vector control culture carrying the pDS-proCAT plasmid containing the riboswitch but lacking a downstream gene. Five biological replicates were performed for each expression experiment. Phage gene inactivation Phage gene inactivation mutants were generated using the method described by Shitrit et al. (2021). In brief, a culture of Synechococcus WH8109 carrying a vector with a recombination template for gene inactivation was infected with the wild type Syn5 phage. The recombination template consisted of a short tag sequence (23bp) inserted between two regions of homology in the phage genome (∼250bp long) that flank the region to be deleted. Infection of Synechococcus WH8109 carrying the recombination vector resulted in a double homologous crossover between the phage genome with the inactivation template producing a deletion of the gene. The mutant phage was then isolated by enrichment and PCR screening. Full genome sequencing was performed for all phage strains, including the wild type strain to verify an identical genetic background. Phage infection properties The length of the latent period was determined using phage growth curves. Synechococcus WH8109 cultures were infected with Syn5 phage strains at an MOI of 1. Samples were collected periodically, diluted 10-fold in ASW and filtered over a 0.22µm Millex GV syringe filter (Millipore, Cork, Ireland) to remove cells and to collect phages in the extracellular medium. The number of infective phages in the filtrate was then quantified using the plaque assay (see above). Average burst size and virulence was determined using a burst size assay. Synechococcus WH8109 (∼1·10 8 cells/mL) cultures were infected with Syn5 phage strains at an MOI of 1. At maximal phage adsorption, 60 minutes post infection, the infected culture was diluted 10 5 -fold, to prevent additional infection steps. The number of infected cells was subsequently calculated using the infectious centers assay on samples at the point of maximal phage adsorption and prior to cell lysis [ 59 ]. The plaque assay was used to determine the total number of infectious units in both filtered (0.22µm) and unfiltered samples, the latter of which contains both infected cells and free phages. The number of infected cells is calculated by subtracting the number of free phages (filtered) from the total number of infected cells and free phages (unfiltered). The number of free phages produced by the infected cells was determined 24 hours post infection in the 0.22 µm filtrate using the plaque assay. The total number of living cells was determined by colony plating at the point of maximal phage adsorption from an uninfected control culture. Virulence was calculated by dividing the number of infectious centers by the total number of living cells. Burst size was calculated by dividing the number of infective phages at the end of the infection by the number of infected cells. Competition assay A direct competition assay was used to assess the contribution of trxA and g26 to competitive phage fitness (Schwartz & Lindell 2017). Equal concentrations of two phage strains were used to infect the same Synechococcus WH8109 culture (10 8 cells/mL) and multiple infection cycles of infection were allowed by infecting at an MOI of 10 -4 . The relative abundance of each phage strain was determined before and after 24 hours of competition by plating plaques and screening each plaque by PCR to determine its genotype (n=96). PCR primers were designed to amplify across the region of deletion, such that wild-type plaques had full length fragments whereas mutant plaques had shorter fragments. The PCR was carried out in 20µL reactions using 10µL Taq PCR MasterMix (Tiangen) and 0.4µM primers (see Table S1 for primer pairs and their use). PCR cycling consisted of an initial step at 95°C for 5 minutes, followed by 34 cycles of denaturation at 94°C for 30 seconds, annealing at 61°C for 30 seconds, elongation at 72°C for 1 minute and a final elongation step at 72°C for 5 minutes. Plaque fragments were identified and counted after gel electrophoresis in 1.25% agarose gel containing ∼0.2µg/mL ethidium bromide. Phage gene expression Transcription of specific phage genes were determined for the wild-type and mutant Syn5 phage strains during infection of Synechococcus WH8109. Infection was performed at a cell concentration of ∼7.5*10 7 cells/mL and at an MOI of ∼3. Approximately 7.5*10 7 infected cells were collected by centrifugation at 15,000g for 2 minutes at 4°C. Cell pellets were flash-frozen in liquid nitrogen and stored at −80 °C prior. RNA was extracted as previously described by Zborowsky & Lindell (2019). Briefly, cell pellets were thawed, resuspended in 10 mM Tris⋅HCl (pH 8) with 100 units of RNase inhibitor (Applied Biosystems) and treated with 15000 units of lysozyme (Sigma-Aldrich) for 30 min at 37°C to lyse the cells. RNA was isolated using the Quick RNA Mini Prep Kit (Zymo) and residual DNA was removed using 2 units of TURBO DNase™ (Invitrogen). Reverse transcription (RT) was conducted using LunaScript™ RT SuperMix Kit (New England Biolabs). 6µM of random hexamer primers were annealed to RNA at 25°C for 2 minutes, followed by cDNA synthesis at 55°C for 10 minutes and heat inactivation at 95°C for 1 minute. The cyanobacterial rnpB gene was used as a positive control for RT for all samples. No RT controls were performed on all samples to ensure that reported transcript levels were not from residual phage DNA. Samples were diluted 2-fold in Nuclease-free Water (New England Biolabs) and stored at −20 °C prior to real-time qPCR (see below). Phage protein translation was determined using the infection procedure described above. Approximately 3.75*10 9 Synechococcus WH8109 cells were harvested at 60 min post infection by centrifugation at 5407g for 5 minutes at 21°C. The resulting cell pellets were flash-frozen in liquid nitrogen and stored at −80°C. Proteins were extracted using the method previously described by Zborowsky & Lindell (2019). Briefly, proteins were extracted in 2% sodium deoxycholate (SDC) and 50mM ammonium bicarbonate by 2 cycles of sonication. They were then reduced with 3mM dithiothreitol, modified with 10mM iodoacetamide, and digested twice with modified trypsin (Promega) at a 1:50 enzyme to protein ratio, in 1% SDC and 50mM ammonium bicarbonate. The deoxycholate was removed by centrifugation, 1% formic acid was added, and the samples were centrifuged again as described above. The tryptic peptides in the supernatant were desalted using C18 tips (Ultra-Micro), dried, and resuspended in 0.1% formic acid. The peptides were resolved by reverse-phase chromatography and mass spectrometry (MS) was performed with a Q Exactive Plus Mass Spectrometer (Thermo Fisher Scientific). MS data were analyzed using MaxQuant 1.5.2.8 software against the proteomes of Synechococcus WH8109 and the Syn5 phage from the UniProt database. A protein was considered to be expressed if the log 2 of LC-MS/MS signal intensity was above the detection limit of 18. Phage DNA replication Intracellular phage DNA replication was assessed by qPCR after infecting Synechococcus WH8109 at a concentration ∼7.5·10 7 -1·10 8 cells/mL at an MOI of ∼3. Samples were collected at various time points after infection and subjected to a quantitative DNA extraction procedure following Zinser et al. (2006) with modifications by Zborowsky & Lindell (2019). Briefly, 200μL of cell culture was collected on 25mm, 0.2μm pore-sized polycarbonate filters (General Electric) by filtration. Cells were washed 3 times with 3mL of ASW medium and once with 3mL of preservation solution (10mM Tris⋅HCl [pH 8], 100mM EDTA, 0.5M NaCl). The cells were then frozen in liquid nitrogen and stored at −80 °C. Cells were removed from filters by immersion in 10mM Tris⋅HCl (pH 8) and agitation in a bead-beater (Mini-BeadBeater, Biospec) for 2 minute (3450 oscillations/min) without beads. DNA was extracted from cells by heat lysis at 95°C for 15 min. Phage DNA was quantified by real-time qPCR (see below) using primers for the DNA polymerase gene of Syn5 (Table S1). Real-time quantitative PCR (qPCR) Real-time quantitative PCR (qPCR) was used for quantification of phage intracellular DNA and phage transcription as previously described by Zborowsky & Lindell (2019). Reactions were carried out on a LightCycler 480 Real-Time PCR System (Roche) with a cycling program as previously described and annealing temperature as described in Table S1. The cycle threshold (Ct) in which the fluorescence of a sample rose above background fluorescence was calculated using LightCycler 480 software (release 1.5.0) using the absolute quantification/second-derivative maximum analysis package [ 64 ]. Specificity of the amplified PCR product was verified by performing melting curve analysis. The following modifications were made to the original method: Each qPCR reaction contained 1X LightCycler 480 SYBR Green I Master mix (Roche), 500nM desalted primers (Table S1), and 2μL of template in a total reaction volume of 20μL. Standard curves were generated from phage genomic DNA to calculate the number of gene copies in samples. Phage genomic DNA was extracted using the Promega Wizard® DNA Clean-Up system and its concentration measured using Qubit TM dsDNA HS kit in the Qbit 3 fluorometer. DNA concentrations were converted to genome copies per milliliter by entering the length of the phage genome into the URI Genomics & Sequencing Center calculator ( http://cels.uri.edu/gsc/cndna.html ). Statistical analyses All statistical analysis was performed using MathWorks Inc. MATLAB software, R2024a. For time-series data, we used repeated measures ANOVA. Sphericity (equality of variance of differences) for time-series analysis was tested using the Mauchly’s test. When the assumption of sphericity was violated the Greenhouse-Geisser correction was applied to calculate the p-value of the repeated measures ANOVA. For pairwise comparison of means, either the Student’s t -test (parametric) or Mann–Whitney U test (nonparametric) was used depending on the data distribution . Normality was tested using the Shapiro–Wilk test with the Shapiro-Wilk and Shapiro-Francia normality tests package [ 65 ]. To determine the latent period, we performed a change point analysis. We used the ischange MATLAB function to detect the initial rise in the log-transformed number of infective viruses. Log transformation was performed to increase the accuracy of latent period time determination by reducing the impact of sampling variability. Download figure Open in new tab Figure S1: Multiple alignment of phage and bacterial thioredoxins. The scale represents the number of amino acids from the N-terminus of the multiple alignment. The putative catalytic site of thioredoxin is marked by a dashed red box. Download figure Open in new tab Figure S2: The effect of Syn5 trxA deletion on the translation of downstream genes. ( A ) Transcription of Syn5 trxA , g26 , DNA polymerase in wild-type and mutant phage. ( B ) Relative protein abundance in cells infected by the wild type and the mutant Syn5. Wild type is shown in black and the trxA mutant in green. Relative protein abundance values are LC-MS/MS log2 average normalized intensity values of Syn5 TrxA, gp26, DNA polymerase 1 hour post infection. Genes are shown from left to right in their order in the Syn5 genome. Error bars represent standard deviation of 3 independent experiments. **p-value<0.01, ***p-value<0.001. Download figure Open in new tab Figure S3: Theophylline-dependent riboswitch induced translation of beta-glucuronidase (GUS). Beta-glucuronidase activity under different concentrations of theophylline, normalized to 0 mM. GUS activity 0.5 hours (blue), 7.5 hours (orange), 24.5 hours (yellow), 48.5 hours (purple), 72.5 hours (green) after theophylline addition. Bars represent averages of 4 biological replicates, error bars represent standard deviations. Individual points are shown by grey circles. View this table: View inline View popup Table S2 thioredoxin and gp26 occurrence and overlap in cyanophages Acknowledgments We thank Idan Yalin and Shirly Larom for their suggestions and discussion on the thioredoxin activity assay; David Keho for his suggestions and guidance with the GusA assay; Udi Qimron for the idea to try and complement T7 with Syn5 and host thioredoxin; Oded Beja and the Lindell lab members for their support and discussions. This research was funded by the European Research Council, ERC Consolidator Grant 646868, and the Simons Foundation Life Grant 735081 to DL. 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