Metabolic engineering of Caldicellulosiruptor bescii yields increased hydrogen production from lignocellulosic biomass
© Cha et al.; licensee BioMed Central Ltd. 2013
Received: 15 April 2013
Accepted: 28 May 2013
Published: 3 June 2013
Members of the anaerobic thermophilic bacterial genus Caldicellulosiruptor are emerging candidates for consolidated bioprocessing (CBP) because they are capable of efficiently growing on biomass without conventional pretreatment. C. bescii produces primarily lactate, acetate and hydrogen as fermentation products, and while some Caldicellulosiruptor strains produce small amounts of ethanol C. bescii does not, making it an attractive background to examine the effects of metabolic engineering. The recent development of methods for genetic manipulation has set the stage for rational engineering of this genus for improved biofuel production. Here, we report the first targeted gene deletion, the gene encoding lactate dehydrogenase (ldh), for metabolic engineering of a member of this genus.
A deletion of the C. bescii L-lactate dehydrogenase gene (ldh) was constructed on a non-replicating plasmid and introduced into the C. bescii chromosome by marker replacement. The resulting strain failed to produce detectable levels of lactate from cellobiose and maltose, instead increasing production of acetate and H2 by 21-34% relative to the wild type and ΔpyrFA parent strains. The same phenotype was observed on a real-world substrate – switchgrass (Panicum virgatum). Furthermore, the ldh deletion strain grew to a higher maximum optical density than the wild type on maltose and cellobiose, consistent with the prediction that the mutant would gain additional ATP with increased acetate production.
Deletion of ldh in C. bescii is the first use of recently developed genetic methods for metabolic engineering of these bacteria. This deletion resulted in a redirection of electron flow from production of lactate to acetate and hydrogen. New capabilities in metabolic engineering combined with intrinsic utilization of lignocellulosic materials position these organisms to provide a new paradigm for consolidated bioprocessing of fuels and other products from biomass.
Keywordsldh Metabolic engineering Switchgrass Biohydrogen Caldicellulosiruptor
Fuel production from plant biomass offers the opportunity to generate energy from a sustainable feedstock, reduce dependence on petroleum, and reduce the negative environmental impact of increased CO2 emissions. The major obstacle in the use of lignocellulosic feedstocks is the recalcitrance of the biomass itself. Plants have evolved to resist deconstruction by microbes, and plant cell wall components such as cellulose, hemicellulose, and lignin play a major role in recalcitrance [1–3]. Industrial conversion of plant biomass to fuels currently relies on thermal and chemical treatment of biomass to remove hemicellulose and lignin, followed by enzymatic hydrolysis to solubilize the plant cell walls to generate a fermentable substrate for fuel-producing organisms [4–6]. However, these methods add cost, produce hydrolysates that are toxic to microorganisms  and are destructive to the sugars in the biomass . An alternative approach is to use consolidated bioprocessing (CBP), in which the fermentative organism is also responsible for production of the biomass-solubilizing enzymes . Members of the genus Caldicellulosiruptor are able to ferment all primary C5 and C6 sugars from plant biomass and are the most thermophilic cellulolytic bacteria known, with growth temperature optima between 78°C ~ 80°C . They can also grow on and degrade biomass containing high lignin content as well as highly crystalline cellulose without conventional pretreatment [11–13], raising the possibility of further economic improvement of biofuel production from plant biomass by reducing or eliminating the pretreatment step.
While Caldicellulosiruptor species are attractive platforms for fuel and chemical production from plant biomass, the dearth of genetic tools for this genus has prevented rational strain development. Recent advances have enabled genetic transformation of Caldicellulosiruptor bescii, opening the possibility of metabolic engineering for improved biofuel production in this genus.
We hypothesized that by developing the necessary tools to delete genes from the C. bescii chromosome [14, 27, 28], we would enable metabolic engineering to increase H2 production. Here, we demonstrate the utility of gene deletion in the pyruvate metabolic pathway for rational strain engineering of C. bescii while simultaneously creating a platform for further strain modification for advanced production of fuels and chemicals from renewable plant feedstocks.
Deletion of lactate dehydrogenase (ldh) from the C. bescii chromosome
Deletion of ldh eliminates lactate production and increases acetate and H2production
To compare the production of lactate, acetate and hydrogen, C. bescii wild-type and mutant strains were grown in LOD medium  with soluble cellodextrans (cellobiose) or plant biomass (switchgrass) as carbon source. When grown on 0.5% cellobiose for 30 hours, JWCB017 showed 29% and 21% more acetate production and 37% and 34% more hydrogen production than wild type and parent strains, respectively (Figure 3D). Cells grown for 120 hours on LOD medium supplemented with 0.5% switchgrass as the sole carbon source showed a similar profile to that on cellobiose, with the Δldh strain producing 38% and 40% more acetate and 55% and 70% more hydrogen than wild-type and parent strains (Figure 3E).
Growth yield increases upon deletion of ldh
We have built upon recent advances in the genetic manipulation of Caldicellulosiruptor[14, 27, 28] to delete the gene encoding lactate dehydrogenase. While the wild type strain produced roughly equimolar amounts of acetate and lactate, the JWCB017 mutant strain no longer produced lactate, instead rerouting carbon and electron flux to acetate and H2, respectively. JWCB001 (~1.8 mol/mol of glucose) and JWCB005 (~1.7 mol/mol of glucose) appeared a bit lower in hydrogen yield than reported values for C. saccharolyticus (~2.5 mol/mol of glucose) . However, in this report, C. saccharolyticus was grown in culture media with added yeast extract, which improves yields . JWCB017 (~3.4 mol/mol of glucose) was more than that reported for C. saccharolyticus. Yield and titer of acetate and H2 were increased in the C. bescii ldh deletion strain from both model soluble substrates and real-world plant biomass. A similar approach has been applied to other thermophilic biomass-degrading bacteria, including xylanolytic Thermoanaerobacterium saccharolyticum and cellulolytic Clostridium thermocellum[30–32], with the goal of increasing ethanol production.
Members of the genus Caldicellulosiuptor offer special advantages for biomass conversion to products of interest in that they are hyperthermophiles with optimal growth temperatures between 70°C ~ 80°C and they are capable of using biomass without conventional pretreatment.
Interestingly, deletion of ldh resulted in a higher cell yield and longer exponential growth phase relative to the wild type. The increase in cell density is likely caused by an increase in acetate production, which should increase ATP production per glucose via acetate kinase providing more energy for biosynthesis and growth. The evolutionary pressures that selected for maintenance of ldh are not clear, though it may be related to the partial pressure of H2 found in the environment. Further, the molecular mechanism by which C. bescii switches from production of acetate + H2 to lactate is unknown. C. thermocellum encodes a lactate dehydrogenase that is allosterically activated by fructose-1,6-bisphosphate , such that lactate is only produced when the rate of substrate uptake exceeds glycolytic flux. It would be interesting to examine whether C. bescii utilizes a similar mechanism for flux control at the pyruvate node of glycolysis. Independent of mechanism, the fact that C. bescii JWCB017 grows to a higher density without an obvious effect on growth rate suggests that further engineered strains may be able to compete well with the wild type strain and thrive in an industrial setting.
Recent progress in genetic tool development opened the possibility of more advanced metabolic engineering strategies to increase the utility of C. bescii for industrial applications. In addition to the construction of gene deletions, this will enable gene insertion into the chromosome (so called gene knock-ins), simplifying the process of heterologous gene expression by eliminating the need for plasmid maintenance and increasing the number of genes that can be stably expressed. Thus, we have created a new platform for rational strain design for lignocellulosic bioconversion, enabling future efforts to increase the titer of H2, express heterologous pathways for production of liquid fuels and chemicals, increase robustness, and improve upon the native ability of Caldicellulosiruptor species to deconstruct and convert biomass without conventional pretreatment.
Characterization of the JWCB017 mutant also sheds light on the basic physiology of C. bescii, which will inform future metabolic modeling and engineering efforts. For instance, this strain has now been engineered to produce only acetate and H2 from sugars, providing further evidence that C. bescii uses a bifurcating hydrogenase to funnel all the electrons to H2. Examination of the C. bescii genome sequence reveals that glycolysis likely yields NADH from glyceraldehyde-3-phosphate oxidation, based on the presence of glyceraldehyde-3-phosphate dehydrogenase (Cbes1406) and lack of glyceraldehyde-3-phosphate: ferredoxin oxidoreductase. Conversion of pyruvate to acetyl-CoA, on the other hand, presumably reduces ferredoxin, based on the presence of pyruvate: ferredoxin oxidoreductase (Cbes0874-0877) and the lack of pyruvate dehydrogenase and pyruvate-formate lyase. Because H2 evolution from NADH is thermodynamically unfavorable except in extremely low H2 partial pressures , this implies that the favorable production of H2 from ferredoxin drives the unfavorable NADH-dependent H2 production. Further genetic modification will increase our understanding of metabolic flux in C. bescii, allowing better metabolic models and further informing metabolic engineering efforts.
Here we show the first application of recently developed genetic methods for metabolic engineering of a member of the genus Caldicellulosiruptor. The method for creating a deletion of the ldh gene in the C. bescii chromosome was efficient enough to allow targeted marker replacement using non-replicating plasmids. The resulting mutant grew to a higher cell density and produced more hydrogen than the wild-type strain. Using the tools developed here, C. bescii JWCB017 will serve as a platform for additional rational strain engineering for production of fuels and chemicals from lignocellulosic feedstocks.
Strains, growth conditions and molecular techniques
Plasmids and C. bescii strains (JWCB) used in this study
Oligonucleotides used in this study
Sequence 5’- 3’
Construction of pDCW121
To construct a plasmid for deletion of the ldh gene (Cbes1918), three cloning steps including overlapping polymerase chain reactions were used. All PCR amplifications were performed using Pfu Turbo DNA polymerase (Agilent Tech., Santa Clara, CA). A 1,009 bp fragment containing a KpnI site upstream of the ldh gene was amplified using primers DC348 and DC349. A 1,011 bp fragment containing an EcoRI site downstream of ldh, was amplified using primers DC350 and DC351. The two fragments were joined by overlapping PCR using primers DC348 and DC351 to generate a 2,020 bp product that was cloned into pDCW88  using the Kpnl and EcoRI sites. The resulting plasmid, pDCW121, was transformed into E. coli DH5α by an electrotransformation via a single electric pulse (1.8 kV, 25 μF and 200 Ω) in a pre-chilled 1 mm cuvette using a Bio-Rad gene Pulser (Bio-Rad, Hercules, CA). Transformants were selected on LB solid medium containing apramycin (50 μg/ml final).
Competent cells, transformation and mutant selection in C. bescii
To prepare competent cells, a 50 ml culture of JWCB005 was grown in LOD minimal medium at 75°C for 18 hours (to mid exponential phase) and 25 ml of the culture was used to inoculate a 500 ml culture of LOD (low osmolarity defined growth medium) supplemented with 40 μM uracil and a mixture of 19 amino acids (5% inoculum, v/v) . The 500 ml culture was incubated at 75°C for 5 hours and cooled to room temperature for 1 hr. Cells were harvested by centrifugation (6000 × g, 20 min) at 25°C and washed three times with 50 ml of pre-chilled 10% sucrose. After the third wash, the cell pellet was resuspended in 50 μl of pre-chilled 10% sucrose in a microcentrifuge tube and stored at −80°C until needed. Before transformation, plasmids from E. coli cells were methylated in vitro with C. bescii methyltransferase (M.CbeI, ) and methylated plasmid DNAs (0.5-1.0 μg) were added to the competent cells, gently mixed and incubated for 10 minutes in ice. Electrotransformation of the cell/DNA mixture was performed via single electric pulse (1.8 kV, 25 μF and 350 Ω) in a pre-chilled 1 mm cuvette using a Bio-Rad gene Pulser (Bio-Rad, Hercules, CA). After pulsing, cells were inoculated into 10 ml of LOC medium (low osmolarity complex growth medium, ) and incubated for 4 hours at 75°C. 100 μl of the culture was transferred into 20 ml of defined medium without uracil. After 18 hours incubation at 75°C, cells were harvested by centrifugation (at 6000 × g for 20 min) and resuspended in 1 ml of 1× basal salts. 100 microliters of the cell suspension was plated onto solid defined media with 40 μM uracil and 8 mM 5-FOA (5- fluoroorotic acid monohydrate).
Analytical techniques for determining fermentation end products
Batch fermentations were conducted in stoppered 125 ml serum bottles containing 50 ml LOD medium with 5 g/l maltose, cellobiose or switchgrass. Cultures of JWCB005 and JWCB017 were supplemented with 40 μM uracil. Triplicate bottles were inoculated with a fresh 2% (v/v) inoculum and incubated at 75°C without shaking. Total cell dry weight (CDW) was determined by concentrating 25 ml of each culture on dried, preweighed 47 mm Supor membrane filters (0.45, Pall Corp., Ann Arbor, MI) and washed with 10 ml of ddH2O. Cell retentates were dried for 16 hours at 85°C and weighed on an analytical balance. Culture supernatants were analyzed via HPLC using a Waters Breeze 2 system (Waters Chromatography, Milford, MA) operated under isocratic conditions at 0.6 ml/min with 5 mM H2SO4 as a mobile phase. Analytes were separated on an Aminex HPX-87H column (Bio-Rad Laboratories, Hercules, CA) at 60°C and monitored via refractive index (RI) using a Waters 2414 RI detector. Total peak areas were integrated using Waters Breeze 2 software and compared against peak areas and retention times of known standards for each analyte of interest. H2 was measured using an Agilent Technologies 6850 Series II Gas Chromatograph equipped with a thermal conductivity detector at 190°C with a N2 reference flow and a HP-PLOT U Column (30 m * 0.32 mm). To measure organic acid production, Nuclear magnetic resonance (NMR) analysis was performed. One-dimensional 1H-NMR spectra were recorded at 298 K with a Varian Inova-NMR operating at 600 MHz for 1H and equipped with a 5-mm NMR cold probe. Samples (500 μL) of cell free culture media were mixed with 150 μL of D2O as internal lock and immediately analyzed. 128 scans were recorded for each sample using a pre-saturation method to suppress the water resonance. The amounts of the most abundant components in the samples were calculated by integration of the proton signals in the spectra. The data were normalized to the amount of acetic acid in each sample.
Air-dried switchgrass (Panicum virgatum, Alamo variety) was reduced to 60 mesh using a Wiley Mini-Mill (Thomas Scientific, Swedesboro, NJ, USA). The ground switchgrass was subjected to a hot water treatment similar to that described by Yang et. al.  however the biomass was boiled in distilled H2O (2% w/v) for 1 hour rather than treating overnight at 75°C. The switchgrass was then washed and dried overnight at 50°C before dispensing into serum bottles as previously described .
High-performance liquid chromatography
Nuclear magnetic resonance
Low osmolarity defined growth medium
Low osmolarity complex growth medium
Cell dry weight
We thank Jennifer Copeland for outstanding technical assistance, Bob Kelly and Sara Blumer-Schuette for providing the Caldicellulosiruptor species, Maria Pena and William York for NMR analysis, Li Tan and Debra Mohnen for assistance with the HPLC analysis and Jonathan Mielenz for providing the switchgrass used in this study. The BioEnergy Science Center is a U.S. Department of Energy Bioenergy Research Center supported by the Office of Biological and Environmental Research in the DOE Office of Science.
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