Optimizing cyanobacteria growth conditions in a sealed environment to enable chemical inhibition tests with volatile chemicals

https://doi.org/10.1016/j.mimet.2016.05.011Get rights and content

Highlights

  • 0.5 g/L NaHCO3 is optimal for cultivating cyanobacteria in a sealed environment.

  • Adding NaHCO3 in a fed-batch manner marginally improves growth.

  • Sealed environment is necessary to maintain volatile chemicals in solution.

  • SYTO® 9 fluorescence assay accurately monitors cyanobacterial growth with minimal sample.

Abstract

Cyanobacteria are currently being engineered to photosynthetically produce next-generation biofuels and high-value chemicals. Many of these chemicals are highly toxic to cyanobacteria, thus strains with increased tolerance need to be developed. The volatility of these chemicals may necessitate that experiments be conducted in a sealed environment to maintain chemical concentrations. Therefore, carbon sources such as NaHCO3 must be used for supporting cyanobacterial growth instead of CO2 sparging. The primary goal of this study was to determine the optimal initial concentration of NaHCO3 for use in growth trials, as well as if daily supplementation of NaHCO3 would allow for increased growth. The secondary goal was to determine the most accurate method to assess growth of Anabaena sp. PCC 7120 in a sealed environment with low biomass titers and small sample volumes. An initial concentration of 0.5 g/L NaHCO3 was found to be optimal for cyanobacteria growth, and fed-batch additions of NaHCO3 marginally improved growth. A separate study determined that a sealed test tube environment is necessary to maintain stable titers of volatile chemicals in solution. This study also showed that a SYTO® 9 fluorescence-based assay for cell viability was superior for monitoring filamentous cyanobacterial growth compared to absorbance, chlorophyll α (chl a) content, and biomass content due to its accuracy, small sampling size (100 μL), and high throughput capabilities. Therefore, in future chemical inhibition trials, it is recommended that 0.5 g/L NaHCO3 is used as the carbon source, and that culture viability is monitored via the SYTO® 9 fluorescence-based assay that requires minimum sample size.

Introduction

Fossil fuels are a finite resource, and it is well established that the massive use of fossil fuels has led to pollution and detrimental health effects in many organisms (Chen et al., 2011). Beyond the long-recognized negative environmental impacts of smog formation and ozone depletion, global warming is a more recently recognized effect of fossil fuel use (von Blottnitz and Curran, 2007). Due to these environmental concerns, it is urgent to develop efficient, clean, and secure systems for the production of biofuels from sustainable sources (Becerra et al., 2015, Gu et al., 2012).

One potential source of renewable biofuels is the photoautotrophic, diazotrophic cyanobacterium Anabaena sp. PCC 7120 (herein referred to as Anabaena sp. 7120). This microbe is capable of being genetically engineered to produce next-generation biofuels and high-value chemicals such as linalool (Gu et al., 2012), limonene (Halfmann et al., 2014b), and farnesene (Halfmann et al., 2014a). These chemicals are insoluble or have low solubility in water, and this hydrophobic nature leads to high bio-concentration in aquatic organisms such as cyanobacteria, making these chemicals quite toxic at low concentrations (Mayer et al., 2000). Thus it is important to increase the microbe's tolerance to these chemicals to improve productivity and the industrial potential of these photoautotrophs.

The chemicals previously mentioned are highly volatile, thus chemical inhibition tests must be conducted in a sealed environment with minimal headspace. This will enable maintenance of desired titers of the chemical of interest during the incubation period. Unfortunately, this means that sparging with CO2 enriched air, or even simple exposure to atmospheric CO2 cannot be used to supply carbon for cell growth. An alternative carbon source for cyanobacteria is NaHCO3. Many cyanobacterial species are capable of taking up HCO3 from the environment via transport across the plasma membrane into the cytosol. There, CO2 is derived from HCO3 by carbonic anhydrase maintaining a steady flux to ribulose-1,5-bisphosphate carboxylase/oxygenase for photosynthesis (White et al., 2013).

Studies have been conducted on algal chemical inhibition tests with volatile chemicals, but to the best of our knowledge no studies have used cyanobacteria. Mayer et al. (Mayer et al., 2000)) used 0.3 g/L NaHCO3, while Herman et al. (Herman et al., 1990)) used 4 g/L NaHCO3 for algal chemical inhibition tests. Mayer et al. (Mayer et al., 2000)) also supplemented the medium with 2% CO2 which was adapted from a study by Hailing-Sørensen et al. (Hailing-Sørensen et al., 1996)). CO2 was used both for carbon enrichment and to act as a pH buffering agent. However, supplementing the medium with a physiological buffer, such as HEPES could also suit this purpose.

While Herman et al. (Herman et al., 1990) did evaluate different NaHCO3 concentrations, the vessels used had a significant volume of headspace. Also, both Herman et al. (Herman et al., 1990) and Mayer et al. (Mayer et al., 2000) performed these trials with algal rather than cyanobacterial strains. Thus it is necessary to determine the optimal concentration of NaHCO3 for growth of a cyanobacteria strain (Anabaena sp. 7120), and if supplementing with NaHCO3 in a fed-batch manner would further increase growth in a sealed environment.

An additional challenge with chemical inhibition tests with hydrophobic chemicals is that the chemicals can have sorption interactions with the cyanobacterial biomass itself and/or the walls of the culture vessel, thereby altering the effective concentration exposed to the cells (Mayer et al., 2000). To minimize this problem, it was recommended that trials are conducted with low biomass levels (Mayer et al., 1997, Nyholm and Peterson, 1997, Peterson and Nyholm, 1993). However, at low biomass levels, classical methods of monitoring culture biomass are less accurate. For example, optical density is considered to have borderline sensitivity and precision at the biomass levels of standard algal toxicity tests (Mayer et al., 1997). Another issue to consider is that optical density and chlorophyll α (chl a) content can be easily affected by biomass debris formation (Robertson et al., 1998).

A fluorescence viability assay has previously been shown by Johnson et al. (Johnson et al., 2016) to be a superior method of monitoring viability of Anabaena sp. 7120 at low biomass titers when compared to optical density and chl a content. Determining if there is a strong correlation between the viability assay and absorbance, chl a content, and biomass content would provide further evidence that the viability assay is an accurate means of monitoring cell viability.

For next-generation biofuels and high-value chemical production from cyanobacteria to become industrially feasible, it is essential to develop strains with increased tolerance to the chemicals that they will be engineered to produce. Because many of these compounds are highly volatile, a sealed environment will be necessary to maintain the chemical titer in solution. Biomass levels must also be minimized to ensure constant chemical-to-biomass concentrations. Therefore, the objectives of this study were to: 1) determine the most accurate and reproducible methods to monitor cyanobacterial growth and viability in a sealed environment, 2) determine the optimal initial concentration of NaHCO3 and if fed-batch addition of NaHCO3 would enhance growth, and 3) compare cyanobacterial growth in the sealed test tube environment optimized in the previous objective to growth in test tubes that are not sealed.

Section snippets

Microbial strains, maintenance, and culture conditions

Anabaena sp. PCC 7120, a model species for filamentous cyanobacteria (Bryant, 2006, Rippka et al., 1979), was obtained from the Pasteur Culture Collection of Cyanobacteria (Paris, France). For long term storage, strains were frozen at − 80 °C in 5% v/v methanol. For short term maintenance the cyanobacteria were grown on BG11 agar (Allen and Stanier, 1968) (1.5% agar) at pH 7.1, incubated at room temperature under constant illumination of 24 μmol m 2 s 1, and then stored at room temperature. Light

Correlation of growth parameters in sealed test tubes

As the first step of this study, we monitored growth of Anabaena sp. 7120 in sealed test tubes using 0.5 g/L NaHCO3 as the carbon source to determine which growth measurements provided accurate and reproducible results at low biomass levels and small sample volumes. This was necessary, since low cell biomass levels are recommended for chemical toxicity testing (Mayer et al., 1997, Nyholm and Peterson, 1997, Peterson and Nyholm, 1993). At high biomass levels the chemical can bind to the biomass

Conclusions

The purpose of this study was to optimize conditions for filamentous cyanobacterial growth in a sealed environment. Growth in a sealed environment is necessary in order to assess the tolerance of strains to volatile chemicals, as well as to develop mutants with increased tolerance via constant exposure to the chemicals. Several research groups are engineering cyanobacteria to produce high-value chemicals and next-generation biofuels directly from CO2 and sunlight. If the tolerance of the

Acknowledgements

This work was supported by the South Dakota Agricultural Experiment Station under grant SD00H398-11. This work was also supported by NASA under award No. NNX11AM03A. We acknowledge use of the South Dakota State University Functional Genomics Core Facility supported in part by NSF/EPSCoR Grant No. 0091948 and by the State of South Dakota. The authors would like to acknowledge the guidance and assistance of Charles Halfmann, and Dr. Huilan Zhu throughout this study. The authors would also like to

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