A low-cost, multiplexable, automated flow cytometry procedure for the characterization of microbial stress dynamics in bioreactors
© Brognaux et al.; licensee BioMed Central Ltd. 2013
Received: 14 March 2013
Accepted: 30 October 2013
Published: 31 October 2013
Microbial cell population heterogeneity is now recognized as a major source of issues in the development and optimization of bioprocesses. Even if single cell technologies are available for the study of microbial population heterogeneity, only a few of these methods are available in order to study the dynamics of segregation directly in bioreactors. In this context, specific interfaces have been developed in order to connect a flow cytometer directly to a bioreactor for automated analyses. In this work, we propose a simplified version of such an interface and demonstrate its usefulness for multiplexed experiments.
A low-cost automated flow cytometer has been used in order to monitor the synthesis of a destabilized Green Fluorescent Protein (GFP) under the regulation of the fis promoter and propidium iodide (PI) uptake. The results obtained showed that the dynamics of GFP synthesis are complex and can be attributed to a complex set of biological parameters, i.e. on the one hand the release of protein into the extracellular medium and its uptake modifying the activity of the fis promoter, and on the other hand the stability of the GFP molecule itself, which can be attributed to the protease content and energy status of the cells. In this respect, multiplexed experiments have shown a correlation between heat shock and ATP content and the stability of the reporter molecule.
This work demonstrates that a simplified version of on-line FC can be used at the process level or in a multiplexed version to investigate the dynamics of complex physiological mechanisms. In this respect, the determination of new on-line parameters derived from automated FC is of primary importance in order to fully integrate the power of FC in dedicated feedback control loops.
Clonal populations of microbial cells exhibit phenotypic heterogeneities due to environmental factors and even in homogenous environments, considering stochastic fluctuations at the level of biochemical reactions . Microbial heterogeneity is governed by a complex set of intrinsic, extrinsic and external noise components that have been thoroughly studied at the fundamental level [2–4] but only partially applied in the field of bioprocess engineering . Indeed, even though the heterogeneity of microbial populations has received a lot of attention over the past few years, the real impact of this phenomenon on microbial bioprocesses remains poorly understood. The main reason behind this lack of knowledge is the difficulty in monitoring microbial population heterogeneity in dynamic process conditions. In this respect, flow cytometry (FC) is a very powerful tool for following physiological properties of microbial cells under process-related conditions . The main advantage of this method is to provide information about the phenotypic heterogeneity of a microbial population. This information is critical from a bioprocess improvement perspective since the appearance of an unwanted phenotype can impair its efficiency. Such phenomena have been recently been noted during culture of recombinant Pichia pastoris with the appearance of a non-secreting phenotype . However, if many techniques are available for the determination of cell physiology at a given moment in a culture, dynamic evolution of microbial resistance to stress and adaptation is still poorly described. For this purpose, an automated version of FC has been proposed for on-line monitoring of cell population heterogeneity under process-related conditions [8, 9]. Specific interfaces comprising a mixing chamber where microbial cells are diluted prior to analysis and stained (if required) have been developed and commercialized. In this way, FC could be used for feedback control at the level of a bioreactor control. Such application is of particular importance for the optimization of bioprocesses based on recombinant microorganism. Indeed, the induction of protein synthesis is often performed by adding a high concentration of inducer molecule at a given moment of the culture. However, this kind of protocol is known to induce a strong physiological stress at the level of the host cell which in turn impairs bioprocess productivity . In this case, on-line FC can be used in order to monitor the intracellular protein synthesis at the single-cell level and the resulting signal can be exploited in order to trigger the feeding of an inducer. This kind of strategy has been previously investigated at the lab-scale level, but no practical applications are available at this time . However, the use of automated FC is still largely underexploited in view of its power in the context of bioprocess optimization. No application of this technique for bioreactor control could be found, except for one very basic application where automated FC was used to control cell density inside a chemostat (cytostat) . The main reason limiting the application of automated FC to microbial bioprocesses is the necessity for a complex interface between the FC and the bioreactor to be sampled. Commercial systems, such as the Flowcytoprep (MSP corp, MN) device, are available but are generally expensive . In this work, we propose to use a benchtop Accuri flow cytometer as the basis for the design of an automated FC. This apparatus was recently tested on microbiological samples and led to reliable results . In addition, fluid displacement is ensured by peristaltic pumps, facilitating the set-up of an interface with a bioreactor since no pressurization of the sample is needed. The development of previous systems was indeed impaired by the need to maintain pressure at the level of the sample unit [15–17]. Under this condition, FC can be easily interfaced to a bioreactor by using additional peristaltic pumps operated by a microcontroller. Sample dilution and staining is carried out in line in the tubing between the FC and the bioreactor. This automated FC system was tested by following the dynamics of an Escherichia coli pfis::gfpAAV fluorescent bio-reporter . The reporter system consisted of an E.coli strain carrying a growth dependent promoter, in this case the fis promoter, fused to a gene expressing an unstable variant of GFP. The pfis promoter is induced in early stationary phase or when cells are shifted from low to high substrate availability . This reporter system is thus a good indicator of the nutrient status of the cells. Indeed, in a mixing-deficient bioreactor, zones with high and low nutrient availability can coexist and significantly affect microbial physiology . In order to increase the responsiveness of the reporter system, a destabilized gfpAAV variant exhibiting a half-life of less one hour was used . The use of automated FC is thus particularly useful for this application since off-line sampling would affect the quality of the results. The combined use of the pfis::gfpAAV bio-reporter with automated FC was investigated for the first time. In a second study, the automated FC interface was multiplexed in order to monitor a platform of parallelized bioreactors. The high-throughput effectiveness of multiplexed FC was demonstrated by extracting dynamic data, giving new insights about the behaviour of the gfpAAV molecule under bioprocess conditions.
Results and discussion
Following fis::gfpAAVactivity by automated FC
A comparison between off-line and on-line samples was performed and no significant differences were observed at the level of GFP synthesis (Additional file 1: Figure S1). Moreover, off-line analyses of extracellular proteins in supernatants confirmed the occurrence of protein leakage during the chemostat phase at D = 0.14 h-1 (Additional file 2: Figure S2), potentially explaining the rise in GFP synthesis during this phase according to our hypothesis. Another hypothesis that can be advanced in order to explain the GFP over-expression during the chemostat phase involves the characteristics of the GFP variant itself. Indeed, GFPAAV exhibits a C-terminal tag recognized by the ATP-dependent ClpXP machinery . The stability of the GFPAAV molecule is thus dependent on the protease content of the cells as well as on ATP availability. These two factors, and particularly the ATP level , are known to fluctuate during bioreactor operations, altering the degree of stability of the GFP. The rise in the GFP signal observed during the chemostat phase could thus also be explained by a decrease in the intracellular ATP pool fuelling the ClpXP protease complex. Unfortunately, both hypotheses are strengthened by the fact that the GFP signal decreases during the second phase of the chemostat phase (Figure 1, after 48 hours of culture). This observation can be explained either by down-regulation of the fis promoter and/or a decrease in the stability of the GFPAAV molecule after the ATP pool is refueled. An explanation about the respective contributions of the different above-mentioned mechanisms could thus be experimentally validated by a complex set of proteomic, transcriptomic and metabolomic profiling of the chemostat phase. In the context of this work, we demonstrate a multiplexed version of the automated FC platform could also be used in order to decipher these dynamics. Prior to that, observations about secondary cytometric variables, i.e. PI uptake by the cells, will be considered in the next section.
Substrate limitation induces a significant segregation of the population according to PI uptake
Multiplexing potentialities of the automated FC
In order to test the multiplexing potential of our automated FC, additional cultivation tests were carried out on a mini-bioreactors platform. This platform comprised, in our case, three parallel stirred vessels with a working volume of 200 mL each and fully equipped with standard controls. It must be pointed out that, in general, the number of reactors in parallel tends to be increased as much as possible and mini-bioreactor platforms comprising 10 to 24 vessels are common [32, 33]. These experiments also allowed exploring the second hypothesis to explain the GFP accumulation under prolonged substrate limitation: a loss of capacity by the cells to degrade the GFP protein. The aim of the experiments was to highlight the equilibrium between GFP caused by fis induction, GFP degradation by proteases and the activity of these proteases that are fuelled by ATP.
This set of experiments demonstrated the value of using a simple flow cytometer interface that can be multiplexed with several bioreactors in order to perform high throughput experiments and gain insights at the level of complex biological processes under dynamic conditions.
Towards a definition of variables for on-line characterization of population segregation in bioreactors
where %R1 is the percentage of PI negative cells and %R2 the percentage of PI intermediate cells, whereas || stands for the absolute value of the difference between the two percentages.
A low-cost FC platform has been designed in order to follow both GFP synthesis and PI uptake inside bioreactors. This system is simple, robust and gives reliable results compared with off-line analysis. However, one of the main limitations of FC is its requirement for fluorescent tag for physiologically relevant analysis. In the context of this work, a pfis::gfpAAV bio-reporter was used in order to track nutrient status at the single cell level. Automation of FC analysis is particularly useful in the case of a destabilized reporter, since off-line analysis requires sample processing that can affect the quality of the results. However, our results have shown that the response of the pfis::gfpAAV system is altered by unknown physiological mechanisms attributable either to up-regulation of the fis promoter or to the destabilized GFP itself. In this context, a multiplexed version of automated FC allowed us to demonstrate that the degree of stabilization of the GFPAAV is involved in this process and is correlated to the intracellular ATP content of the cells. Finally, MMR has been proposed as a useful parameter for on-line detection of microbial population segregation. Overall, this work demonstrates that a simplified version of on-line FC can be used at the process level or in a multiplexed version to investigate the dynamics of complex physiological mechanisms. In this respect, the determination of new on-line parameters (e.g. the MMR) is of primary importance in order to fully integrate the power of FC into dedicated feedback control loops.
Microbial GFP reporter strain and medium
Escherichia coli K12 MG1655 bearing a pGS20 plasmid with a pfis::gfpAAV gene and a chloramphenicol resistance gene was used in this work. Additional details about strain construction can be found in . pGS20PfisBAAV was digested with NotI and NdeI to replace the rrnB promoter region with the E. coli fis promoter that encodes a DNA binding protein, FIS. The fis promoter was amplified from the E. coli chromosome using primers FisP-SD (this primer contains an optimized SD sequence) and FisP-up. The 300 bp PCR product was subsequently digested with NotI and NdeI and ligated into the vector fragment. The resulting plasmid was called pGS20FisGFPAAV (10–15 copies/cell, chloramphenicol resistant marker, 25 μg/mL) .
The strain was maintained at -80°C in working seed vials (2 ml). Pre-cultures and cultures were grown on a defined mineral salt medium containing (in g/L): K2HPO4 14.6, NaH2PO4.2H2O 3.6, Na2SO4 2.0, (NH4)2SO4 2.47, NH4Cl 0.5, (NH4)2-H-citrate 1.0, glucose 5.0, thiamine 0.01, and chloramphenicol 0.05. Thiamine and kanamycin were sterilized by filtration (0.2 μm). The medium was supplemented with 3 mL/L trace solution, 3 mL/L FeCl3.6H2O solution (16.7 g/L), 3 mL/L EDTA solution (20.1 g/L) and 2 mL/L MgSO4 solution (120 g/L). The trace solution contained (in g/L): CoCl2.H2O 0.74, ZnSO4.7H2O 0.18, MnSO4.H2O 0.1, CuSO4.5H2O 0.1, and CoSO4.7H2O 0.21. Before operating in the bioreactor, a pre-cultivation step was performed in 100 mL of the above-mentioned medium in a baffled shaker flask at 37°C under orbital shaking at 140 rounds per minute during 16 h.
Automated FC protocol
Bioreactor operating conditions
Chemostats were carried out in a lab-scale stirred bioreactor (Biostat B-Twin, Sartorius with a total volume of 3 L; working volume of 1 L; mixing provided by a standard Rushton disk turbine with 6 blades) in remote control mode interfaced with the MFCS/win 3.0 software. During the experiments, pH was maintained at 6.9 (regulation by ammonia and phosphoric acid) the stirrer rate at 800 rpm, the air flow rate at 1 L/min and the temperature at 37°C. The chemostat phase was stabilized during 43 h with a feeding solution containing 5 g/L of glucose solution (in the same minimal medium as previously described) at a dilution rate of 0.14 h-1(corresponding to 6 residence times). Glucose pulses were performed during 24 h at the same dilution rate by adding pulses of 8 mL of a glucose solution (30 g/L made up in distilled water) each 15 minutes. Additional batch experiments were performed on a Dasgip mini-bioreactor platform. The mini-bioreactors were filled with 200 mL of the defined medium previously described. Stirring was provided by two Rushton turbines with 6 blades at an agitation rate of 900 min-1. The air flow was maintained at 100 mL/min and the pH was kept at 7.0. The first experiments involved three bioreactors at three different temperatures: 30°C, 37°C and 42°C. A second experiment was also performed. The three reactors were started at 30°C. After 4 h, the temperature was increased in the reactors to 42°C to induce a thermal shock and ATP-dependent protease synthesis. In the second and the third bioreactors, an acetate pulse and a glucose pulse, respectively, were delivered at the same time (2 g added in total). All these experiments were carried out with two replicates. At both scales, cell growth was monitored by optical density (OD) at a wavelength of 600 nm (Genesys 105 UV–VIS spectrophotometer, purchased from Thermo Scientific). Cell dry weight was determined on the basis of filtered samples (0.45 μm) dried during 24 h at 105°C.
Glucose concentration was monitored by a YSI (Yellow Spring Instrument Co) electro-enzymatic system. Samples were injected into a chamber filled with a buffer solution and the sample was diffused through a polycarbonate membrane that limited the reaction rate. Membrane-immobilised glucose oxidase produced hydrogen peroxide that diffused through a cellulose acetate membrane and was oxidized by a palatine electrode. The signal was proportional to the glucose concentration up to 2.5 g/L. More concentrated samples were diluted first. The total amount of protein in the extracellular medium was measured on the basis of the Folin-Lowry method .
ATP assays were performed on supernatants from the mini-bioreactors according to the CellTiteGlo® luminescent assay. The assay procedure involved adding a simple reagent directly to the cells. This resulted in cell lysis and generation of a luminescent signal proportional to the amount of ATP present, measured by a V3 Wallac luminometer (Perkin Elmer). This relies on the properties of a thermostable luciferase that generates a stable luminescent signal for 5 hours: mono-oxygenation of luciferin is catalysed by luciferase in the presence of Mg2+, ATP and molecular oxygen. One hundred microlitres of reagent was added to 100 μl of culture medium (the cell concentration has to be less than 5.104 cell/ml). A calibration curve was done over the same measurement range. All analyses were performed in triplicates and expressed as specific ATP, i.e. the ATP concentration divided by cell optical density.
AB is the recipient of a PhD FRIA grant provided by the Belgian Fund for Scientific Research (FRS-FNRS). The authors gratefully acknowledge Samuel Telek and Rudy Schartz for their excellent technical assistance.
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