Genome Engineering in Cyanobacteria: Where We Are and Where We Need To Go.

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Genome Engineering in Cyanobacteria: Where we are and where we need to go C. Josh Ramey, Ángel Barón-Sola, Hanna R. Aucoin, and Nanette R. Boyle ACS Synth. Biol., Just Accepted Manuscript • DOI: 10.1021/acssynbio.5b00043 • Publication Date (Web): 18 May 2015 Downloaded from http://pubs.acs.org on May 24, 2015

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ACS Synthetic Biology

Genome Engineering in Cyanobacteria: Where we are and where we need to go C. Josh Ramey, Ángel Barón-Sola, Hanna R. Aucoin, Nanette R. Boyle1 Chemical and Biological Engineering Department, Colorado School of Mines, Golden, CO 1

Corresponding Author [email protected]

Abstract Genome engineering of cyanobacteria is a promising area of development in order to produce fuels, feedstocks and value added chemicals in a sustainable way. Unfortunately, the current state of genome engineering tools for cyanobacteria lags far behind those of model organisms such as Escherichia coli and Saccharomyces cerevisiae. In this review, we present the current state of synthetic biology tools for genome engineering efforts in the most widely used cyanobacteria strains and areas which need concerted research efforts to improve tool development. Cyanobacteria pose unique challenges to genome engineering efforts because their cellular biology differs significantly from other eubacteria; therefore, tools developed for other genera are not directly transferrable. Standardized parts, such as promoters and ribosome binding sites, which control gene expression, require characterization in cyanobacteria in order to have fully predictable results. The application of these tools to genome engineering efforts is also discussed; the ability to do genome wide searching and introducing multiple mutations simultaneously is one area that needs additional research in order to enable fast and efficient strain engineering.

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Introduction The dramatic decrease in the cost of DNA synthesis and sequencing in the last decade coupled to advances in molecular biology have enabled the development of synthetic biology and led to the sophisticated re-engineering of cellular machinery. The standardization of biological parts, such as promoters and other regulatory elements, allows researchers to edit and re-write genomes in a fast, efficient and predictable way. It is not currently possible to redesign entire genomes de novo; however, in the two most widely researched model organisms (E.coli and S. cerevisiae), this goal is within reach. Subsets of cellular machinery have been re-engineered in some organisms, one notable example is the complete refactoring of the nitrogen fixing gene cluster in Klebsiella oxytoca1. Despite the significant progress which has been achieved in re-writing genomes using synthetic biology techniques; the application of synthetic biology methods and techniques needs to be broadened to a larger group of organisms to fully reach its potential. One major area that needs concerted research effort is the development and use of synthetic biology tools in cyanobacteria. Cyanobacteria hold great promise as platform strains for the production of chemicals and feedstocks, especially since they require only carbon dioxide and light as substrates. Unfortunately, the current state of knowledge of these organisms lags far behind other model systems. Recently, research in this area has improved the ability to engineer cyanobacteria, but we are still in need of continued effort to enable more predictable and efficient redesign of these organisms as industrial production strains. In this review, we will discuss the current state of synthetic biology and genome engineering in cyanobacteria and future directions which need additional research. Standardized Parts One goal of synthetic biology is the development of standard genetic elements which can be ‘pulled off the shelf’ and combined to control gene expression within the cell of any organism. Some regulatory elements are conserved in bacteria and thus a few fully characterized parts from E. coli can be used in cyanobacteria, however, the majority of these parts are not directly transferrable. Below, we will discuss the current state of knowledge for standard parts in cyanobacteria and where improvements can be made. Promoters and terminators The first step in designing or discovering new methods to control transcriptional regulation is to understand the differences between cyanobacteria and typical eubacteria. Cyanobacteria have different consensus promoter sequences and their RNA polymerase holoenzyme (RNAP)2, 3 only functions with sigma (σ) factors belonging to the σ70 –family4, 5. Three different types of σ70 have been described that bind three different promoter sequences (I-III) in cyanobacteria. Group 1 σ70 factors prefer RpoD consensus promoters (Type I) similar to other eubacteria (-35 and -10 elements as TTGACA and TATAAT)6. Group 1 σ70 factors are responsible for the expression of common housekeeping genes and thus are indispensable to the cell. Group 2 and 3 σ70 factors have been reported to increase in concentration in response to environmental stresses6 such as heat7-9, salt10, oxidative11, or osmotic stress12 or nutrient limitation6, 13. Type II promoter sequences have only the -10 hexamer or enhancer motif sequences associated with transcriptional activator proteins; Type III promoters are distinct from both Type I and II but little else is known about their specificity and interaction with σ factors. Recently, Kaczmarzyk


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et al. improved tolerance to butanol and high temperatures (up to 45˚C - 48˚C) by increasing expression of a group 2 cyanobacterial sigma factor (SigB) which binds to type II promoter sequences14. Engineering the transcription factors themselves can also be a successful approach to improved complex phenotypes, such as improved solvent tolerance. In yeast, Alper et al. used random mutagenesis to engineer transcription factors which resulted in improved glucose/ethanol tolerance 15. Similar approaches can be used in cyanobacteria to elicit global changes in transcription resulting in improved growth in a variety of stressful growth conditions. The ability to tune gene expression is critical for any re-design of cellular machinery; thus, the identification and/or engineering of promoter sequences which span large dynamic ranges and are both constitutive and inducible are necessary steps to further develop synthetic biology in cyanobacteria. The majority of endogenous promoters currently used to drive heterologous gene expression in cyanobacteria are derived from genes associated with the light harvesting complex of photosynthesis (Table 1). These promoters induce high levels of gene expression; however, as these promoters are light induced, there is little or no gene expression when cells are grown in the dark16. Recently, a light-driven super-strong promoter (Pcpc560) reported in the cyanobacterium Synechocystis sp. PCC6803 has been used efficiently for heterologous gene expression resulting in protein levels similar to that produced using the strong tac promoter in E. coli 17-19 with low or medium-copy plasmids. However, one drawback of using these strong promoters is that they are either ‘on’ or ‘off’, which can lead to unpredictable (and deleterious) physiological changes and does not allow tuning of gene expression. There are a number of tunable promoters which are responsive to environmental cues (such as nutrient concentrations) in cyanobacteria which can be used to overcome this hurdle. Most of the currently known inducible promoters are responsive to micronutrient concentrations (see Berla et al. for a comprehensive review 20), such as metal ion concentrations. However, due to the sensitive nature of these promoters (some are responsive to concentrations as low as 50µM), they are difficult to use in the laboratory or industrial settings due to additional experimental considerations, such as acid washing glassware, the use of trace metal grade chemicals and growing cells in nutrient limited conditions for several generations which may result in non-negligible physiological stresses or low growth rates. Due to the dearth of easily tunable promoters in cyanobacteria, Markley et al. engineered promoter libraries based on the cyanobacterial cpcB promoter in Synechococcus sp. PCC700221. Using YFP as a reporter gene, the resulting library showed a 3log range in promoter strength. The cpcB promoter based library was used to control gene expression in E. coli and results varied dramatically from what was measured in Synechococcus, further evidence that promoter performance in E. coli cannot be used to predict promoter response in cyanobacteria. Engineered E. coli promoter sequences have also been used in cyanobacteria with varying degrees of success23, 24 (Table 1). A transcriptional modulation system based on TetR regulated promoters allowed for the induction of a 290-fold change of eYFP (Yellow Fluorescent Protein) levels25. Despite elevated levels of gene expression using the TetR regulated promoter, the high light sensitivity of the inducer aTc (anyhydrotetracycline) restricts its use to low light conditions (30µmol photons m-2 s-1). Isopropyl β-D-1-thiogalactopyranoside (IPTG) inducible promoters have also been used in cyanobacteria with a wide range of effectiveness. Utilization of the same IPTG inducible promoter, such as Ptrc , in different cyanobacteria strains can result in a high level of variability. For example, the use of Ptrc in Synechococcus elongatus PCC7942 resulted in an increase in gene expression up to 36-fold26 while in Synechocystis sp. PCC6803 there was little



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to no change in gene expression27. A possible explanation for this difference could be the capacity of different cyanobacteria to transport the IPTG molecules into cells. Markley et al. engineered a constitutive promoter library based on well-characterized E.coli promoters using YFP as a reporter gene21 and reported a range of YFP expression of 2.5-log. By combining the cpcB (discussed in previous paragraph) and the E.coli-based promoter libraries, Markley et al.21 engineered an IPTG induced system yielding an approximately 48 fold increase in YFP expression in Synechococcus sp. PCC7002. These results are impressive when compared to the previously reported isiAB-Fe3+ repressor system, which has a 2-fold induction; however, they are still one order of magnitude lower than those obtained with the Ptrc E. coli promoter20. Promoter engineering in cyanobacteria is a powerful tool for controlling gene expression however, it is not without its challenges. These examples illustrate the complexity of this endeavor as these promoters are affected by a multitude of factors such as growth conditions, growth phase, competence to inducer uptake, inducer type and concentration, sigma factors, and promoter type. In addition to promoter sequences, terminator sequences are also extensively used in synthetic biology to control gene expression at the transcriptional level. In prokaryotes, terminator sequences control gene expression via two different mechanisms: intrinsic termination, also known as Rho-independent termination, which occurs when the formation of a stem-loop structure in mRNA causes RNAP to stall. Rho-dependent termination relies on the binding of the Rho protein to the nascent mRNA causing RNAP to be released. Although homologs of E. coli Rho proteins are found in a majority of bacteria, they have not been found in two of the most widely used cyanobacteria (either Synechocystis sp. PCC6803 or Synechococcus elongatus). A high resolution transcriptome map created for Synechococcus elongatus PCC7942 has been used to identify termination sequences. Secondary structures at the 3' end of mRNA transcripts were found to have lower free energy, and therefore a preference to form stem-loop structures just prior to the end of the transcripts as is found in Rho-independent transcription termination28. Additionally, several terminator sequences were found to occur between open reading frames (ORFs) within the same operon suggesting promoter-independent transcription level regulation occurs in cyanobacteria. This mechanism of transcriptional regulation has been used in cyanobacteria by inserting terminator sequences between ORFs to prevent background transcription effects of upstream and/or downstream genes on the gene of interest. The endogenous cyanobacterial Rubisco terminator29, E. coli rrnB strong terminator30, and transcription terminator sequences derived from T723 bacteriophage have also been used in cyanobacteria. Although these terminator sequences are used in synthetic constructs, it is important to note that to date, there have been no concerted efforts to test the effectiveness of terminator sequences in cyanobacteria specifically. Ribosome binding site Ribosome binding sites (RBSs) are also well characterized genetic tools used to control protein expression in model organisms. There are several computational tools available to predict the effect of altering a RBS sequence, such as the RBS Designer Model31, the UTR Designer32 and the RBS Calculator33, 34. The RBS Designer can be used to predict protein translation efficiencies of a given mRNA based on the RBS sequence, spacer length and homology to the Shine-Dalgarno (SD) sequence. The UTR Designer is based on the characterization of specific E.coli mRNA regions that produce dynamic changes in mRNA folding32. Use of this model showed that the 5´ RNA region of the coding sequence is critical for modulating protein


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expression levels. Finally, the RBS calculator is based on thermodynamic analysis of mRNA sequences in E.coli. The effects of different RBS sequences and surrounding nucleotides are considered in this model along with the start codon sequence and putative mRNA secondary structures. In E. coli, the RBS calculator is quite robust and can predict translation rates over a range of 100,000 within a factor of roughly 2. These three RBS prediction software packages were used by Markley et al.21 to design and evaluate a RBS library in Synechococcus sp. PCC 7002. Unfortunately, the predicted levels of YFP expression (up to 213-fold range) were poorly correlated with experimental YFP fluorescence data (30-fold range). The poor correlation between RBS calculator predictions and experimental evidence was also reported in a study to optimize 2,3-butanediol in Synechococcus elongatus sp. PCC 7942 35. Although several RBS models have been developed to improve RBS efficiencies in E.coli, the use of these predictive models in cyanobacteria does not yield predictable results. Thus, protein translation is regulated via a different mechanism in cyanobacteria and deserves further research to develop specific predictive RBS tools for these organisms. It is important to note that while the above studies report changes in gene expression due to changes in promoter and/or RBS, the use of the same expression construct within different genetic context may result in different results36, 37. To our knowledge, there have been no rigorous studies which examine the effect of genetic context on the promoter strength in cyanobacteria, therefore it is difficult to know how much variation will be seen. In recent work by Mutalik et al., approximately 500 transcription and translation initiation units were engineered to minimize effects of secondary structure38. These genetic elements can be used with any downstream interest and expression levels should be within twofold of the target level; this is an 87% reduction in expression error typically reported. In order to achieve the goal of synthetic biologists and have truly interchangeable parts, more research should be done to create fully insulated parts; the results of Mutalik et al. indicate that this indeed can be achieved with enough effort.

RNA Regulators and Post-translational Control Yet another mechanism to control gene expression and/or protein concentrations is by altering RNA conformation or base pairing with DNA or other RNA sequences. These so called RNA regulators can be used to engineer genetic networks. One type of RNA regulator that cyanobacteria rely on is non-coding RNA (ncRNA). Antisense RNA (asRNA), a type of ncRNA, has been reported to regulate gene expression in cyanobacteria in several studies39-41 suggesting an important role of this genetic control in the general metabolism of these organisms42. For instance, an asRNA was found to bind AU-rich box and RBS regions of the psbA2 to prevent the degradation caused by RNAse E in Synechocystis sp. PCC6803. Interestingly, both asRNA and psbA2 transcript abundance were regulated by light and expression levels were correlated43. Riboregulators are specific RNA sequences that respond to signal molecules and can act on the translation process by releasing silencing RNA or activating ribozymes. This system has been successfully adapted from E. coli45 for use in cyanobacteria46 as a method for posttranscriptional control of gene expression. Abe et al.46 designed and tested different riboregulators in E. coli based on the cis-repressed (cr)/ trans-activating (ta) RNA system developed by Isaacs et al.45. The crRNA sequence represses gene expression by binding RBS region of the mRNA. Then,



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taRNA transcript can bind crRNA by disrupting the inactivation of the mRNA RBS thus, activating gene transcription. The authors further evaluated the described riboregulator system using a GFPuv gene as a reporter in Synechocystis sp. PCC6803 and report comparable results to that obtained in E. coli. These RNA-based genetic tools can be used in cyanobacteria to design specific transcriptional approaches to block or promote mRNA degradation allowing for the modulation of metabolic pathways for the optimal production of target compounds. Regulation of mRNA stability can also be conducted using a plasmid based approach controlling the transcript levels of ncRNA using an inducible/repressible promoter. This allows for the targeting of multiple transcripts from the multiple copies of a gene simultaneously by ncRNA transcripts addressing concerns derived from polyploidy nature of cyanobacteria (see Multiple Chromosomes and Associated Challenges). Riboswitches are yet another approach to altering protein levels in a cell. Riboswitches are functional non-coding RNA molecules which have a sensing domain (aptamer) and a regulating domain. A conformational changes occurs when the aptamer domain binds to a specific ligand and thereby affects the expression of the adjacent genes. A recent study in Synechocccus elongatus PCC 7942 reports the use of a theophylline-dependent riboswitch which is capable of inducing protein expression 190 fold47. The riboswitch was also very effective at repressing translation in the absence of theophylline, unlike leaky gene expression when IPTG inducible promoters are used 23, 48, 49. Another advantage of riboswitches is that protein levels of genes under control of the riboswitch can be tuned by altering the concentration of the ligand. Since 16s rRNA is conserved amongst related cyanobacteria, this implies that similar approaches can be used for other model cyanobacteria, such as Synechocystis sp. PCC 6803 and Anabaena sp. PCC 7120. Protein stability is also an important point of regulation; protein degradation tags control the abundance of protein and may prove useful for altering cellular mechanisms. Natural protein degradation mechanisms have been reported in Synechocystis sp. PCC6803 which protect the organism from heat and light stress conditions50. Landry et al.51 evaluated a set of different cyanobacterial and E. coli protein degradation tags fused to the reporter enhanced Yellow Fluorescence Protein (eYFP) in Synechocystis sp. PCC6803. Fluorescence levels showed a wide a range (from 1 to 50% of untagged eYFP) indicating that this system of regulation has the potential to fine-tune enzyme levels in cyanobacteria. Harnessing the natural mechanisms of metabolic control described in cyanobacteria for use as synthetic biology tools can be used to optimize a desired metabolic pathway however, plenty remains to be discovered and understood about transcriptional and translational regulation in cyanobacteria and each should be considered fields to be explored to determine the limitations and advantage of every approach. Genome Engineering Multiple Chromosomes and Associated Challenges A major challenge in any genome engineering effort, regardless of the organism used is obtaining clones/mutants which are homogenous. From early studies in E. coli, it was assumed that prokaryotes contain only a single copy of their genome (monoploid); however, in E. coli, this is only true when growth is extremely slow (td = 16 hours)52. The process of replicating a chromosome in E. coli takes approximately 60 – 90 minutes to complete53; therefore, in order to replicate at faster growth rates which are typically seen in rich medium (td = 20 minutes), E. coli


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must initiate multiple rounds of replication before the previous replication machinery is complete. This results in an average chromosome copy number of 6.8 in normal growth conditions 54, 55. Despite having much slower growth rates, cyanobacteria have also been reported to have multiple copies of their chromosome when grown under ‘normal’ conditions and it can vary from a handful to several hundred. One of the most widely used species for metabolic engineering and synthetic biology is Synechocystis sp. PCC 6803, which is highly polyploid; having chromosome copy numbers ranging from 60 - 225 copies of its chromosome per cell 56 depending on the strain being used (Table 1). The number of chromosomal copies can also vary depending on the growth phase, Griese et al. report that Synechocystis sp. PCC 6803 has an average of 218 chromosome copies in exponential phase which drops to 58 and 57.6 in linear and stationary phase respectively56. Another widely used species, Synechococcus elongatus sp. PCC 7942 is reported to have on average 4 copies of its chromosome per cell in both linear and stationary growth phases 56, 57. Griese et al. also compiled information on several other species, which can range from 1 to 218 chromosomes per cell56. There are several evolutionary advantages to carrying multiple copies of the chromosome, such as efficient repair of double strand breaks, gene redundancy, lower rates of spontaneous mutations and rapid cell divisions. Despite the numerous evolutionary advantages multiple chromosomes provides the cell, they also create a grand challenge for genome engineering, both in terms of creating mutant libraries and tracking them on a chromosome by chromosome basis. Even in organisms with advanced genome engineering tools, such as E. coli, the challenge of working with multiple chromosome copies has not been overcome (see58 for a detailed discussion of how multiple chromosomes affects recombination efficiencies). Desired mutations must either be integrated into every chromosome or continuously selected for until a homogenous population is achieved; if this is not possible (as many mutations are not selectable) the mutation will be diluted out significantly due to further replication and division of WT cells. This is a significant problem in E. coli, where the copy number is only 6.8; in cyanobacteria, where some species have hundreds of copies of their chromosome present it becomes especially problematic. If desired mutations are introduced on one or even a few chromosomes, they will quickly be diluted by growth during the recovery period. No reports to date address this problem specifically in E. coli or cyanobacteria, however if ploidy can be finely controlled without significant defects on growth, recombination efficiency can be increased tremendously. The fewer chromosomes present, the fewer recombination events need to occur (or shorter outgrowth period) to have homogenous populations. As discussed earlier, cells grown more slowly have lower chromosome copy numbers, however this significantly decreases the ability to introduce mutations quickly. Therefore, control of ploidy is a major challenge that needs to be addressed in any genome engineering effort and has yet to be addressed in many of the genome engineering literature. Genomic, Expression and Deletions Libraries in Cyanobacteria The use of genomic libraries to understand complex genetic phenotypes such as increased production of bio-products and understanding tolerance mechanisms are well established59, 60. Cyanobacteria have been engineered for the production of ethylene, sucrose and 2, 3 butanediol; however, yields still need to be improved for economic feasibility61, 62. The development of genetic tools such as genomic libraries, expression libraries and gene deletion collections will facilitate the development of cyanobacteria into an important production organism. The



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construction of genomic libraries in cyanobacteria requires the continued improvement of shuttle vectors between cyanobacteria and E. coli (current vectors for cyanobacteria have recently been reviewed16). Plasmid vectors currently used in cyanobacteria either use a replicon of a broad host range plasmid or an endogenous cryptic plasmid. Important considerations for the development of an effective genomic library in cyanobacteria are the following: plasmid copy number, plasmid partitioning into daughter cells, library insert size, genetic coverage and available selectable markers. One concern that is specific for using endogenous plasmids is any effect caused by endogenous genes or DNA sequence elements present on the plasmid. Use of shuttle vectors for the construction of a genomic library in E. coli allows for all cyanobacteria genomic sequences to be represented. Construction in E. coli permits the identification of genes and DNA sequences whose overexpression is toxic under normal conditions in cyanobacteria, as well as genes and DNA sequences that promote increased biosynthesis of chemical products by increasing tolerance or flux through important biochemical pathways. Cyanobacteria genomic library approaches and resources applied thus far are listed in Table 1. To date genomic libraries have been utilized for complementation analysis, genome sequencing or as a starting point for constructing a knockout library using transposon mutagenesis of genomic library plasmids in E. coli (Table 1). Genomic libraries are a simple overexpression at the level of DNA sequence, therefore when developing genomic libraries, an important consideration is the plasmid copy number. Due to the polyploidy nature of cyanobacterium discussed above, the effectiveness of a simple overexpression at the DNA level might be limited. To overcome this limitation, an expression library of all known open reading frames (ORFs) of a particular cyanobacterium needs continued development; this requires a sequenced and annotated genome (Table 1). Also listed in Table 1 are expression libraries created in cyanobacteria species and strains. Expression libraries have been reported for Synechocystis sp. PCC 6803, Synechococcus elongatus and Anabaena. The expression system used for Synechococcus elongatus was a bacteriophage lambda based system that utilizes the lac promoter to identify carotenoid binding proteins by isolating immunopositive clones63. This library was designed to express Synechococcus elongatus proteins in E. coli and thus cannot be utilized to elucidate genes which confer complex phenotypes in the endogenous host. A cosmid based expression library has been reported in Synechocystis for identification of enzymes involved in light dependent chlorophyll biosynthesis by complementing a defect in a R. capsulatus strain64. In Anabaena, an expression library was created by ligating (2-10kb) genomic fragments into a plasmid containing the highly expressed endogenous rbcL promoter65. These reported expression systems are insufficient for understanding complex phenotypes because they do not control gene expression over a dynamic range and the genomic fragments inserted into the expression vectors are not defined. Ideally, genomic libraries would be constructed in shuttle vectors with interchangeable pieces (promoters, RBS, and terminators) so that different levels of gene expression could be tested in the native host to aide in identifying gene targets for further engineering. Defined expression library collections using inducible promoters and optimized ribosome binding site sequences are an important tool for the development of increased production of bioproducts and to facilitate the understanding of tolerance mechanisms. One advantage of expression libraries is that protein levels can be controlled over a much greater dynamic range than a simple genomic library. A current limitation of expression libraries in cyanobacteria is the lack of an effective and easy to use cyanobacteria-specific inducible promoter that does not


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affect the overall physiology of cyanobacteria (see 20 for a comprehensive list of promoters used in cyanobacteria). As discussed previously, the majority of inducible promoters native to cyanobacteria are trace metal responsive; these promoters require the removal of inducer metal ions which have deleterious effects to the overall physiology of the cell and growth rate. There are a number of E. coli based promoters which have been engineered to function in cyanobacteria which can be used for this purpose but their range of expression can be limited or problematic. For example, IPTG inducible promoters are known to be leaky in the absence of the IPTG. Advanced metabolic engineering requires the ability to fine tune entire metabolic pathways; to test different levels of gene expression in different genetic backgrounds, genomic or expression libraries require the availability of promoters whose strength can be tuned easily. Expression library construction should allow for easy shuffling of vector backbones controlling plasmid copy number and promoters controlling transcript levels. The construction of an expression library could be accomplished through traditional means or take advantage of homologous recombination cloning methods in cyanobacteria or Saccharomyces cerevisiae67, 68. Important considerations for the development of expression libraries are the following: plasmid copy number, promoter strength, translation efficiency, ribosome binding sequences, terminators, selectable markers available and downstream techniques for introducing the overexpression into the genome. Multiple methods have been described for the construction of knockout libraries in cyanobacteria 20, 69, 70 including: homology based recombination of selectable markers, markerless knockout using selection/counter selection strategies, transposon mutagenesis of genomes and transposon mutagenesis of plasmid libraries in E. coli that are then integrated back into the host genome71. Cyanobacteria are an attractive model for genetic manipulation because they are naturally competent, easily taking up foreign DNA and integrating it into the genome through homologous recombination pathways. An important limitation for the construction of deletion libraries is the polyploid genome of cyanobacteria. This is overcome via continued selection over multiple generations, thus greatly increasing the time needed to create a single gene knockout. By traditional methods of constructing a knockout library, each gene knockout takes approximately three weeks. One caveat to this approach is that obtaining homogeneous strains that contain only the gene knockout and not the wild type gene is selected against if the gene knockout negatively affects the cell. Using a plasmid shuffle approach can solve this problem. A covering plasmid containing the wild type gene with a counter selectable maker allows for all the genomic copies of the gene to be deleted while the gene function is covered by the plasmid copy of the gene, followed by counter selection against the plasmid. Another important consideration when constructing a knockout library in cyanobacteria is species/strain selection. For example, if the genetic target or molecular process being targeted is involved in photosynthesis, it is important to determine if your species is an obligate photoautotroph; if it is, this would preclude any efforts to knockout or reduce activity of those essential genes. This is one reason why Synechocystis sp. PCC 6803 is used extensively as a model organism – it can be grown purely autotrophically, mixotrophically or heterotrophically. It is important to note that it has been recently shown that the introduction of sugar transport systems allow for Synechococcus elongatus PCC7942, an obligate photoautotroph, to grow using saccharides such as glucose, xylose and sucrose in the absence of light72. We believe that constructing a knock-out library in cyanobacteria is a worthwhile effort that the community should work on in a massive parallel fashion while next generation knockout approaches such as Crisper/Cas9 and TALEN systems are developed. Examples of collaborative efforts for the construction of a deletion collection in Saccharomyces cerevisiae and E. coli are available to model an approach to making a deletion



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collection in cyanobacteria73, 74 67. The construction of a knockout library is an excellent opportunity to integrate research into undergraduate education. Current knockout libraries in species and strains of cyanobacteria are listed in Table 1. Multiplexing As discussed above, the generation of a single gene knockout in a fast-growing cyanobacterium can take approximately three weeks using conventional methods. Therefore, in order to design and create production strains in a timely and cost-effective manner, it is imperative to develop methods which allow the introduction of several genetic mutations at once, thus streamlining the process (Figure 1). The first step required for multiplexing in cyanobacteria is to identify the genes to target for a given phenotype. In E. coli, genes associated with a given phenotype (for example, solvent tolerance) can be identified using genomic library enrichment strategies. For example, MultiScalar Analysis Library Enrichment (SCALE)75 uses clones from a plasmid-based genomic library to identify individual genes or group of genes responsible for the desired phenotype 76-78. The enrichment of clones which are able to grow in the given condition is quantitatively evaluated by hybridization or next generation sequencing. A similar method that uses barcodes to track genetic modifications is TRackable Multiplex Recombineering (TRMR)79. This approach merges the yeast barcoding method80 with homologous recombination processes in order to integrate and track mutations in the chromosome. These types of genome-wide searching tools have not be developed for use in cyanobacteria, but they are necessary in order to rationally engineer production strains by targeting genes which are known to increase or improve growth in the desired conditions. It has been shown that tuning the expression of genes in an entire pathway can lead to higher productivities when compared to just overexpressing genes encoding rate limiting enzymes81. In order to tune the expression level of several genes simultaneously, methods such as Multiplex Automated Genome Engineering (MAGE) need to be developed for cyanobacteria. MAGE, which was developed in E. coli, enables the simultaneous generation of mutants with multiple genetic modifications. Based on evolutionary processes within a cell population, MAGE is able to generate up to 15 billion genetic variants per day in E. coli81. This approach involves a large-scale introduction of synthetic single stranded DNA (ssDNA) oligos which encode desired mutations into a genome using a short cycling automation process (2 - 2.5 h). With the longer doubling times of even the fastest growing cyanobacteria (Table 1), achieving similar library sizes would take much longer (1 week to 1 month); however, would still be faster than creating mutants with desired gene expression levels one at a time. One of the most challenging aspects of creating a MAGE-like method for creating mutant libraries is how to design an effective screen or selection to identify mutants with the desired phenotype. This is exacerbated by the high chromosomal copy number in cyanobacteria; it is a critical step to reduce the heterogeneity of the population and to minimize the number of clones which need to be tested further in order to identify the ‘winner’. Multiplex genome engineering methods, such as MAGE and TRMR, would enable researchers to design, build and test production strains (Figure 1) faster and more efficiently than current methods allow. This expanded ability to generate mutants would also have the additional benefit of generating increased knowledge of processes within the cell which differ from model bacteria (such as E. coli). CRISPR-Cas systems


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Clustered regularly interspaced short palindromic repeats (CRISPR) along with CRISPR associated proteins (Cas proteins) have been described in Eubacteria and Archaea as an adaptive immune system against foreign genetic elements86, 87. This immune systems has been used as a basis to design genome editing tools in a variety of organisms, from bacteria88 and yeast89 to plants90 and human cells90, 91. The three major CRISPR/Cas systems described in prokaryotes92 share a common mechanistic process in order to acquire immunity against foreign genetic material (viruses and plasmids). The first stage is the incorporation of external nucleic acids into the host genome CRISPR/Cas regions. The next step involves the transcription of DNA into small pieces of RNA (crRNA) that will be further used as an interference element for targeting and degrading the complementary foreign genetic material. All of the processes are performed by the Cas proteins which function as helicases and nucleases. Unlike other previous genome editing technologies, such as zinc finger nucleases and transcription activator like effector nucleases (TALENs), CRISPR/Cas systems do not rely on DNA-protein interaction but simply work with RNA-DNA complementarity rules. This feature simplifies the protocol by avoiding the complex steps required for protein engineering. Type II CRISPR/Cas system can make insertions and/or deletions (indels) as well as site directed point mutations. This mechanism involves various elements for the maturation of a pre-crRNA into crRNA: a short trans-encoded RNA known as transactivating crRNA (tracrRNA), the nuclease Cas9 (Csn1) and host RNA polIII. Additionally, multiple efforts to improve the efficiency of CRISPR/Cas systems have been performed to optimize the specificity of Cas nucleases93 and prevent reported off-target effects in eukaryotic cells94. CRISPR/Cas systems are becoming a preferred tool as a highly efficient and reliable method for genome editing applications. The presence of CRISPR/Cas repeat sequences have been reported in different cyanobacteria. Analysis of 126 cyanobacterial genomes found that only marine cyanobacterial strains lack CRISPR/Cas sequences95. Genomic studies in the cyanobacterium Synechocystis sp. PCC680396 have described three different types of CRISPR/Cas systems in the endogenous pSYSA plasmid. Each CRISPR system bears different Cas proteins and uses independent processing pathways. Authors found high transcript abundance of CRISPR/Cas sequences and a strong dependency between crRNA accumulation and Cas6 protein activity. Further analysis in the same strain revealed that crRNA accumulation was affected by environmental conditions and the presence of genes encoding putative transcriptional regulators97. These results offer a promising genome editing strategy to improve the design and creation of cyanobacterial production strains. For example, CRISPR/Cas systems can be designed to cut all wild type sequences, therefore eliminating the need for a screen or selection to identify mutants; this makes it an incredibly powerful tool for overcoming the presence of multiple chromosomes in cyanobacteria. CRISPR/Cas systems designed to cut wild type DNA also eliminates the need for long periods of segregation to ensure homogeneous populations. By way of illustration, the CRISPR/Cas9 nuclease system has been successfully used for genome editing polyploid yeast Sacharomyces cerevisiae98. In a single step of mutagenesis, authors obtained a mutant which increased cellobiose fermentation rates by over 10-fold. CRISPR/Cas can be designed to cut any nonmutant chromosomes, CRISPR/Cas also presents an additional advantage – that of immunity to bacteriophages. At the industrial level, bacteriophages can cause huge economic losses if they are unwittingly introduced to a production fermentation processes. Phages are usually considered one of the most significant threats in the bioproduct industry and the application of genome editing



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technologies based on CRISPR/Cas can prevent bacteriophage infections86, 99, 100. Cells can also be engineered to have immunity against cyanobacteria specific phages (cyanophages)101, which addresses one concern of large scale production: crop-protection. CRISPR/Cas genome editing systems are a promising tool to overcome many of the challenges of working with cyanobacteria; it has the potential to reduce the time needed to produce mutants or mutant libraries and can overcome the limitations associated with multiple chromosomes. Conclusion Cyanobacteria are a promising group of organisms to engineer as platform strains using synthetic biology techniques for the production of biochemicals and biofuels. Because cyanobacteria can convert light energy and CO2 to high value chemical products they are a truly sustainable platform for bioproduction. Current limitations for the development of cyanobacteria into industrial productions strains are: increasing CO2 fixation, increasing light capture, reducing intracellular oxidative stress, and the improvement and development of sophisticated genetic tools. In this review, we have focused on the current state of synthetic biology tools and areas where the development of new genetic tools can greatly increase our understanding of the physiology of cyanobacteria as well as the development of strains with increased production and tolerance of bioproducts. It has already been demonstrated that cyanobacteria are capable of producing high value bioproducts; the development of sophisticated genomic tools will help make these bioproducts commercially viable. The development of well-defined genetic libraries such as: genomic libraries, expression libraries and knockout libraries will facilitate a better understanding of complex phenotypes in cyanobacteria. Genetic libraries will also provide the basis for more advanced genome searching techniques such as SCALES and TRMR. In addition to genetic library based techniques there are new exciting synthetic biology techniques that need to be utilized such as Crisper/Cas9 and MAGE. One important limitation of genetic analysis in cyanobacteria is their polyploidy nature. Depending on the species and strain DNA copy number can vary between 2 and more than 200, making it difficult of obtain homozygous strain. Recently the Crisper/Cas9 system has been utilized to obtain homologous mutants in a polyploid industrial production strain of yeast, indicating that Crisper/Cas9 genomic editing technique has promise in ployploid strains of cyanobacteria. The leverage of new synthetic biology techniques and application of knowledge discovered in model organisms will allow for the development of cyanobacteria into an important green production platform for bioproducts.


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Table 1. Cyanobacterial strains for synthetic biology

Strains

Synechococcus sp. PCC 7002

Synechococcus elongatus PCC 7942

Synechocystis sp. PCC 6803

Leptolyngbia sp. BL0902

Representative shuttle vectors

Representative promotersa

pAQE17102 pAQ1Ex-Pcpc 22

cpc22 (E,NI) isiAB103 (E,I)

pUC303105 pCB4106 pSG111107

idiA108 (E,I) smt109 (E,I) trp-lac110 (F,I) trc26 (F,I)

pFC1114 pSL1211115

pAM5052/5057/5059122

isiAB+GFP116 (E,I) trc2023 (F,I) A1lacO-127 (F,I)

trc122 (F,I)

tac126 (F,I) petE127 (E,I) psbA1128 (E,NI) nir129 (E,I) a Endogenous(E)/ Foreign (F)/ Inducible (I)/ Non Inducible (NI) b GeneBank Assembly Accession number

Anabaena (Nostoc) sp. PCC7120

pRL1125 pPMQAK123

Polyploidy (chromosomes/cell)

622

3-6111

60-22556

-----------

8-10130

Doubling time (h)

3.522

12112

624

Genomic, Expression or Knockout Libraries

Sequenced annotated Genomeb

Genomic library104 RBS library21

GCA_000019485.1

Genomic and knock out libraries113 Expression library63 Genomic library117, 118 Customized amplification library119 Random insertion DNA library of mutagenesis120 cDNA library121 Expression library64

~20123

Genomic library124

>24131

Genomic library132 Expression library65


GCA_000012525.1

GCA_000009725.1

------------

GCA_000009705.1


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Figure 1. Development and use of genome engineering tools in cyanobacteria. In order to design and rewrite genomes in cyanobacteria, genome-wide search tools (such as SCALEs, MAGE, TRMR and CRSIPR/Cas) need to be developed to aide in gene discovery. Once genes associated with particular phenotypes are known, their gene expression can be re-designed using standardized parts. The design can then be tested in vivo and the strain can be improved using iterative cycles. It is important to note here that molecular tools depicted in each step of the cycle are not necessarily restricted to the step depicted but can be used in any step of the design-buildtest cycle.


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This figure is for the abstract 482x482mm (96 x 96 DPI)


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Figure 1. Development and use of genome engineering tools in cyanobacteria. In order to design and rewrite genomes in cyanobacteria, genome-wide search tools (such as SCALEs, MAGE, TRMR and CRSIPR/Cas) need to be developed to aide in gene discovery. Once genes associated with particular phenotypes are known, their gene expression can be re-designed using standardized parts. The design can then be tested in vivo and the strain can be improved using iterative cycles. It is important to note here that molecular tools depicted in each step of the cycle are not necessarily restricted to the step depicted but can be used in any step of the design-build-test cycle. 1422x1422mm (96 x 96 DPI)


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