Chemical lysis of cyanobacteria
© Mehta et al. 2015
Received: 17 March 2015
Accepted: 19 May 2015
Published: 5 June 2015
We have developed a mixture of enzymes and chemicals that completely lyse cyanobacteria. Since the treatment involves only readily-available chemicals and simple proteins that degrade the components of the cyanobacterial cell wall, it can easily be used in high-throughput applications requiring lysis for subsequent intracellular measurements. Our lysis technique consistently enables complete lysis of several different cyanobacterial strains, and we demonstrated that DNA, mRNA, and proteins are preserved in the lysates. Chemical lysis can be superior to existing techniques because of its convenience, reliability, and amenability to a variety of downstream applications.
Significant efforts have been directed towards the fundamental goal of using photosynthesis to sustainably produce chemicals and fuels from sunlight and water, usually with either cyanobacteria or algae as hosts . As the simplest photosynthetic organisms, cyanobacteria have attracted particular attention  for a range of applications including harvesting their remarkably wide library of natural products [3–5] and metabolic engineering for chemical production [6–10]. However, these advances have been limited by lower productivities in cyanobacteria than in traditional hosts. Two recent efforts to engineer cyanobacteria to produce chemicals by photosynthesis achieved productivities of 137 mg/L/day for isobutryaldehyde  and 391 mg/L/day for sucrose . To our knowledge, these are the best productivities achieved from cyanobacteria in lab-scale cultures; however, they are almost two orders of magnitude lower than, for example, 14 g/L/day for succinate in E. coli  and 3.6 g/L/day for artemisinic acid in S. cerivisiae .
We believe high-throughput genetic engineering of cyanobacteria will be essential to overcome these limitations and enable the use of photosynthesis to sustainably synthesize chemicals. Applications for high-throughput genetic approaches include screening large libraries of parts  to optimize pathways and genetic mapping via insertional mutagenesis  to help determine genetic influences on observed phenotypes.
Cell lysis to enable intracellular measurements is a fundamental requirement in the strain engineering cycle. It is particularly challenging in cyanobacteria compared to traditional hosts. A recent assessment of several existing methods for cyanobacterial lysis demonstrated effective lysis only by using low-throughput methods such as sonication, mechanical disruption, or lyophilization . A controllable lysis system has also been developed wherein a lytic casette from a bacteriophage, including a lysozyme and accessory proteins, was expressed in Synechocystis sp. PCC 6803 under the control of a nickel-inducible promoter , but this requires prior genetic engineering and is not generally adaptable. Thus, new and convenient techniques amenable to high-throughput analysis are needed.
We sought to develop a simple, one-step chemical process that would degrade each of these barriers systematically. Some reagents for degrading cell wall components are well-established, such as lysozyme for peptidoglycan and detergents to permeabilize membranes. However, since other components of the cyanobacterial cell wall have not been well characterized, we tested a variety of other reagents to ensure all the layers of the cell wall could be breached. Here, we describe the cloning and testing of a novel lysozyme from a cyanobacteria-specific bacteriophage and the development of a chemical treatment to fully lyse cyanobacteria in a high-throughput manner.
We first tested a lysozyme from a cyanobacteria-specific phage  for its ability to more effectively degrade the cyanobacterial peptidoglycan layer relative to currently used enzymes. Our hypothesis was that a lysozyme from cyanobacterial phage might be uniquely suited to degrading the cyanobacterial peptidoglycan layer and might even be capable of breaking through other barriers as well. A novel lysozyme gene (GenBank accession number: KR259643), selected from a family of 23 identified by homology matching of ORFs in DNA from a cyanobacteria-phage community collected in Yellowstone National Park , was cloned and purified in E. coli.
We initially sought to optimize reaction conditions for cyanobacterial lysis. In these experiments, we compared a chemical lysis process using cyanophage lysozyme to a mechanical lysis procedure using vortexed glass beads as the best established procedure for cyanobacterial lysis. We tested a range of temperatures (4–60 °C), reaction pH values (5.5–9.2), and reaction times (20–180 min). In the initial tests, to achieve any lysis, we had to complement the lysozymes with BugBuster, a commercial lysis reagent which we have previously used with E. coli. Lysis was quantified by measuring the absorbance at 660 nm of chlorophyll released from the cells during the reaction; the “extent of lysis” is plotted in terms of raw absorbance units measured in the supernatant after centrifugation.
A treatment with lysozyme and BugBuster at 30–42 °C and pH 6–8 for 90 min achieved the best results, although it did not enable complete lysis (see Additional file 1). In these initial experiments, we estimated that the maximum extent of lysis corresponded to approximately 50 % of the cells present, although the variability was significant.
Our initial tests revealed that lysozyme had to be supplemented with BugBuster to enable lysis. Since BugBuster is a proprietary mixture of unknown reagents, we sought to discover commercially available components that complement T4 lysozyme in breaching the outer barriers of cyanobacteria. Moreover, we found that BugBuster did not enable complete lysis under the optimized conditions mentioned above.
Removing EDTA or cellulase had a minimal effect on lysis efficiency, but spermine and DTT were both important, with less than 50 % lysis activity without either. EDTA was originally included to help destabilize the outer membrane by chelating divalent cations that stabilize negatively-charged sugars; its lack of effect in both Synechocystis (Fig. 3a) and E. coli (Additional file 1) suggests that the detergents by themselves enable sufficient permeabilization of the membranes.
Removing cellulase, which was included to degrade the secreted exopolysaccharide layer, also had a minimal effect on lysis efficiency. This suggests that exopolysaccharide is not a significant barrier to lysozyme and the other additives. However, these experiments were performed on cells in exponential phase, and we suspected that exopoysaccharide would be thicker in stationary phase. Thus we also tested the effect of cellulase on late-stage cultures. Even in this case, cellulase did not increase the extent of lysis (Additional file 1), suggesting the EPS in general does not pose a significant barrier to other components. Based on these results, subsequent experiments were performed with T4 lysozyme, detergents, spermine, and DTT.
We next performed a design-of-experiments (DOE) analysis to optimize the concentrations of these four reagents (Fig. 3b) and potentially discover interactions between cyanophage lysozyme, detergents, spermine, and DTT. It was interesting that several of the combinations tested gave virtually complete lysis. We thought this might suggest an interaction between some of the components in the mixture, but the DOE analysis did not reveal any significant interactions. Marginal means analysis of the DOE results predicted that the optimal combination is 50 μg/mL cyanophage lysozyme, 2 X detergents, 400 μM spermine, and 200 mM DTT; subsequent experiments were done with this cocktail, except that 10 μg/mL T4 lysozyme was used instead of 50 μg/mL cyanophage lysozyme based on a separate optimization for that omponent (see Additional file 1).
Since the requirement for DTT at such a high concentration was particularly interesting, we titrated it separately while keeping the concentrations of the other components constant (Fig. 3c). This confirmed the results of the design-of-experiments, with a significant increase in lysis activity even from 100 to 200 mM. We examine the implications further in the Discussion.
Our chemical treatment uses detergents to permeabilize membranes in the cyanobacterial cell wall, spermine and DTT to weaken the external layer, and lysozyme to break down peptidoglycan. Initially, we thought a lysozyme from a cyanobacteria-specific phage might be uniquely suited to lysing cyanobacteria, but this was not the case.
Lysozymes cleave the linkage between N-acetylmuramic acid and N-acetylglucosamine in peptidoglycan, which is consistent across almost all bacteria. This could explain why T4 lysozyme worked as well as cyanophage lysozyme against cyanobacteria: Although the cyanobacterial peptidoglycan is thicker and more highly cross-linked than in other species, the actual target of the lysozyme is the sugar chain sequence, which is similar. Nevertheless, the overall cyanobacterial cell wall clearly presents a unique challenge, and a systematic approach was required to breach it.
DTT was the most surprising additive required for full lysis (Fig. 3c). It is generally used as a reducing agent. We initially thought the target might be disulfide bonds in the S-layer, but the Synechocystis S-layer protein has no cysteine residues . Certain cell wall components in yeast are disulfide bonded to each other ; it is possible that similar disulfide bonds exist between other components of the Synechocystis cell wall – for example, between membrane proteins, particularly in the outer membrane. T4 lysozyme does not contain disulfide bonds ; combined with the fact that DTT does not significantly promote E. coli lysis, we do not believe DTT acts on the lysozyme. Moreover, the concentration of DTT that was required – at least 50 mM, with up to 200 mM for the full benefit – is far higher than that typically needed to reduce disulfide bonds. We note that the results of the DTT titration (Fig. 3c) indicating a requirement for 200 mM DTT seem to disagree with some of the individual conditions in the design-of-experiments (Fig. 3b). However, the requirement for 200 mM DTT was consistently reproducible over several independent experiments.
We considered the possibility that the requirement for a high concentration of DTT implies a physical effect, perhaps an intercalation in one of the layers of the cell wall, rather than a chemical effect. We tested this by substituting DTT (redox potential of −330 mV at pH 7) with β-Mercaptoethanol (redox potential of −260 mV) and TCEP (a stronger reducing agent than DTT). β-Mercaptoethanol was essentially as effective as DTT. TCEP also effectively contributed to cyanobacterial lysis, but the lysates had a red color instead of the yellow-green color normally seen; this decreased the absorbance measured at 660 nm and suggests that TCEP may reduce other molecules in the lysate, perhaps the pigments (data in Additional file 1). The fact that other reducing agents could replace DTT suggests that the function of DTT in the cocktail is indeed a reducing one (as opposed to a physical effect); further work is needed to determine why such a high concentration is required.
Spermine was also required for complete cyanobacterial lysis. Previous work with T4 lysozyme against E. coli  suggested that spermine directly enhances the muramidase activity of T4 lysozyme at spermine concentrations below 100 μM, and stabilizes spheroplasts at higher concentrations, counteracting that enhancement. However, we achieved virtually complete lysis of E. coli with lysozyme and detergent only, with small additional benefits from either EDTA or DTT. Since spermine is required for lysis of cyanobacteria but not E. coli using our cocktail, we believe its benefit here is through an interaction with a unique feature of the cyanobacterial cell wall and not with lysozyme. At our reaction pH, spermine is fully protonated and carries four positive charges, so it probably does not pass through either cell membrane; most likely, it acts on the exopolysaccharide or S-layer.
To verify biological activity in lysates produced by our cocktail, we tested enzyme activity (using ferredoxin-NADPH reductase as an example enzyme), DNA amplification by PCR, and isolation and reverse transcription of mRNA. We quantified lysis by measuring the release of pigments into the lysate; this necessarily means that the pigments were not damaged by the lysis cocktail. For proteins in particular, adsorption onto the large surface area presented by glass beads can be a problem, so a chemical procedure could provide a significant advantage for protein analysis. We note a potential limitation of our procedure in that the high concentration of DTT could break disulfide bonds in proteins.
At the scales we have used (reaction volumes ~ 100 μL; ~1 μg cells), we found that chemical lysis was significantly more effective than mechanical disruption with beads. We believe this is because mechanical disruption, especially for cells with such a strong cell wall, requires tremendous force often only provided by specialized equipment such as a BeadBeater homogenizer.
We examined a fundamental operation in the strain engineering cycle, cell lysis, from the perspective of the unique cyanobacterial cell wall. As a result, we developed a chemical treatment that fully lyses several strains of cyanobacteria at physiologically relevant conditions and does so more effectively than mechanical disruption with glass beads.
Our optimized cocktail consists of T4 lysozyme at 10 μg/mL, the detergents C7Bz0 at 0.2 % w/v and SB3-14 at 2 % w/v, spermine at 400 μM, and DTT at 200 mM. This reliably enabled full lysis of three different cyanobacterial strains.
Our lysis procedure uses only chemical components, making it easily adaptible to high-throughput applications, and preserves genomic DNA, mRNA, and enzyme activity for further analysis. We believe this will expand the scope of intracellular measurements that are feasible in cyanobacteria.
Strains and plasmids
Plasmids constructed for this work
Protein expression and purification
Proteins were expressed in vivo from BL21 (DE3) strains using IPTG induction. Briefly, 1 L cultures in terrific broth (Invitrogen) supplemented with ampicillin at 100 μg/mL were grown at 37 °C to OD ~1.5 and induced with IPTG at 100 μM. At this point, the temperature was reduced to 25 °C and proteins were expressed overnight. Cultures were harvested by centrifugation at 3000 × g and cells were lysed in a single pass through a high-pressure homogenizer (Avestin) at >18,000 PSI. Debris and insoluble material were removed by centrifugation at 23,000 × g. The soluble supernatant was purified on a streptactin column (IBA Life Sciences) according to the manufacturer’s protocol. After purification, elution fractions were pooled, concentrated ~30 X using a centrifugal filter device (Amicon Ultra, 10 kDa cutoff, Millipore), formulated in 20 % sucrose, flash frozen in liquid nitrogen, and stored at − 80 °C. Typical yields were 1 mg of purified protein from 3 (wet) grams of cells.
Cyanobacterial lysis reaction
Unless otherwise stated, lysis reactions were carried out on Synechocystis sp. PCC 6803 (ATCC 27184) cells grown in BG-11 medium (Sigma) under constant illumination from fluorescent lamps at ~200 μE/m2/s while sparging a mixture of 5 % CO2 in air. Cells were harvested in exponential phase (culture OD750 of 0.5–1; doubling time 10–15 h). For lysis experiments, cells were pelleted by centrifugation and resuspended to OD 1.5 in MES buffer at pH 6.2 with lysozyme and either the specified lysis components (see Results) or 1 X BugBuster protein extraction reagent (Novagen). Reactions were conducted in either microcentrifuge tubes (300 μL reactions, shaken on a vortexer (vortex genie, VWR)) or deep-well 96-well plates (100–200 μL reactions, shaken on a Thermo Scientific titer plate shaker (shake setting 7)). Wells in the plate were sealed with a custom PDMS gasket to prevent cross-contamination while shaking. After the reaction, insoluble debris were removed by centrifugation at 10,000 × g for 5 min, and the degree of lysis was quantified by measuring absorbance of the supernatant at 660 nm, corresponding to a chlorophyll absorbance peak. Total lysis was confirmed by the absence of green color in the debris pellets, indicating that all cells had lysed and released their chlorophyll into the supernatant. Mechanical lysis via glass beads (150–200 μM in diameter; Sigma-Aldrich) was used as a comparison in many experiments, as this is the current standard for cyanobacterial lysis. In these reactions, beads at 25 % of the reaction volume [30, 31] were added to the cell suspension in buffer and detergents (to solubilize chlorophyll); these reactions were incubated and otherwise processed as described above.
All chemicals were obtained from Sigma-Aldrich. T4 lysozyme was obtained from MClab. Cellulases from Trichoderma viride, Trichoderma reesei, and Aspergillus niger (Sigma) were a kind donation from Lynette Cegelski. Design-of-experiments calculations were done in SPSS (IBM). The orthogonal design function was used to generate the set of experimental conditions, and a univariate general linear model was used to analyze the results.
We gratefully acknowledge Devaki Bhaya and Michelle Davison for the gene sequences of cyanophage lysozymes and a sample of Synechocystis OS-B, Rick Zuzow and the Bhaya group for many stimulating discussions, and the Bio-X Interdisciplinary Initiatives Program at Stanford for funding. KKM was also supported by a DOE Office of Science Graduate Fellowship, a Siebel Scholarship, and an ARCS Fellowship.
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