Ethanol yields (g.L−1.day−1) as reported for various constructs using the Zymomonas mobilis (Zm)
Abstract
Photoautotrophic ethanol production using model cyanobacteria is an attractive technology that offers potential for sustainable ethanol production as a biofuel. Model strains of Synechocystis PCC6803 have been metabolically engineered to convert central metabolic intermediates such as pyruvate to acetaldehyde via cloned heterologous pyruvate decarboxylase and from acetaldehyde to ethanol via cloned homologous or heterologous alcohol dehydrogenase. While the technology is now proven, strategies are required to increase the ethanol levels through metabolic and genetic engineering and in addition, production and process strategies are required to make the process sustainable. Here we discuss both genetic and molecular strategies in combination with do wnstream strategies that are being applied while also discussing challenges to future application.
Keywords
- synechocystis
- ethanol metabolic engineering
- challenges industrial production
1. Introduction
As an alternative to ethanol fermentation using carbohydrate substrates, the use of photoautotrophic cyanobacteria metabolically engineered to produce ethanol offers an interesting alternative for sustainable biofuel production. Cyanobacteria or Cyanophyta, the name deriving from their color, are a distinct phylum of bacteria, which are photoautotrophic getting energy from sunlight and carbon from carbon dioxide. They are the only photosynthetic bacteria that can evolve oxygen. Model species such as
The interest in utilizing cyanobacteria as cell factories for ethanol production has been stimulated via flux balance analysis on ethanol yields, which estimate that the stoichiometric energy yield for ethanol compares well with other potential fuel metabolites [3]. The earliest reports of photoautotrophic metabolically engineered ethanol production came in
Genetic construct | Strain | Rate per day (g.L−1.day−1) | References |
---|---|---|---|
ZmPDC and ADH1 P |
0.0082 | [4] | |
ZmPDC and ADH1 P |
0.0766 | [5] | |
ZmPDC and slr1192 | 0.097 | [6] | |
JCC1581 B Isolate | 0.41 | [7] | |
ZmPDC and slr1192 PziaA | 0.236 | [8] | |
ZmPDC and slr1192 PcorT | 0.235 | [8] | |
ZmPDC and slr1192 Prbc | 0.202 | [9] | |
ZmPDC and slr1192 PpetJ | 0.261 | [10] | |
TK504 Plasmid Pco | ABICyanol1 | 0.552 | [11] |
This was followed by reports in several patents from the US biotechnology companies Algenol and Joule Unlimited who further manipulated the system to improve yields (Table 1). The reported yields are represented as a daily yield and often the production cycle can last up to 20 days such that the yields would be multiplied by the production days. However, with potential evaporative loss and degradation of ethanol by contaminants in non-axenic culture these yields are lower than would be needed for commercial production. Thus, much effort has been focusing on improving this yield level by metabolic and strain engineering.
2. The model strain and production of key intermediates
The first
Yields of product such as ethanol are highly dependent on the biomass produced during growth of engineered strains. When growing photoautotrophically at 30°C doubling times of
During photoautotrophic metabolism in
3. Key aspects of the engineered ethanol cassette in Synechocystis
To metabolically engineer
Pyruvate decarboxylase (PDC, EC 4.1.1.1) carries out the decarboxylation of pyruvate to acetaldehyde in alcohol fermentations and requires thiamine diphosphate/pyrophosphate (ThDP) and the divalent cation Mg2+ as cofactors. Several other enzymes in various metabolic pathways also require these cofactors to function and it is believed that each of them use a similar mechanism of action. PDC can be found in fungi, plants and yeast and is not present in humans [21]. PDC genes have been observed and characterized from only a small number of bacterial species as it appears to be rather rare amongst prokaryotes. These include
Bacterial host and enzyme | kM (mM) pyruvate | Optimum pH | Optimum temperature (°C) |
---|---|---|---|
0.06 pH 5.0 0.6 pH 6.0 1.2 pH 7.0 |
5.0–5.5 | 45–50 | |
0.24 pH 6.0 0.71 pH 7.0 |
7.0 | 55 | |
0.39 pH 5.0 5.1 pH 7.0 |
3.5–6.5 | 65 | |
0.43 pH 6.0 0.94 pH 7.0 |
6.0–6.5 | 60 | |
5.7 pH 6.5 4.0 pH 7.0 |
6.3–6.7 | N/A | |
0.12 pH 5.0 2.8 pH 7.0 |
4.5–5.0 | 53 |
While there is a potential choice of PDCs to use, in practice most work so far has been carried out on the
4. Zymomonas mobilis and Synechocystis alcohol dehydrogenase (ADH)
In most reports on engineered ethanol cassettes the source of ADH has been
Unusually
5. Construction of functional ethanol cassettes in Synechocystis
In general, terms the construction of an ethanol cassette follows the basic components as reported [4, 5]. The
In attempts to increase ethanol production, gene dosage has been utilized such that two cassettes have been integrated at different sites giving potentially twice the gene copy number and protein expression level of PDC and ADH [9, 36]. While this strategy has been shown to increase the levels of ethanol produced it may be that given the polyploid nature of
Figure 1 illustrates the construction of an ethanol cassette pUL004 Kan. This cassette [36] consists of the Zm
6. Issues and methodologies to enhance ethanol production levels
6.1. Ploidy as an issue in cloning in Synechocystis
Strains of
Griese et al. using a real time PCR method demonstrated that the motile ‘Moscow strain’ of
6.2. Gene dosage
The initial cloning strategies [4, 5] used one copy of the ethanol cassette inserted into a chromosomal neutral site. To enhance productivity two copies of the cassette where then utilized [9, 36]. This had the effect of increasing productivity all be it at the expense of biomass and indeed stability during production. Attempts in our laboratory to generate strains with three cassette copies per cell have thus far failed. This suggests there could be a limit to the gene dosage that can be utilized for ethanol cassettes at least. This limit may be due to several factors and many of these factors may combine to limit production. There is the ploidy issue such that during growth if the ploidy level is some 50 copies [39] then with one cassette the copy number is already 50, two copies would mean it would be approximately 100 and the cell may not be able to tolerate more. There may also be instability issues with recombination events between similar cassette sequences. In addition, there may be the issue of ability to supply the co-factors ThDP, NADH and NADPH for this level of enzyme expression. There may be additional factors such as limitation of pyruvate for other essential cellular functions if high levels of enzyme activity are utilizing it to react to ethanol. This in turn may affect biomass production and synthesis of essential cell components and thus triggering a stress response. In addition, given the negative effect ethanol has on growth there may be the selective pressure to mutate the cassettes selecting for faster growing strains which do not have the burden of ethanol production. The nature of all these possibilities may need to be examined in more detail to generate optimal strains going forward.
That gene dosage can have an effect on production has been demonstrated by utilizing the small native
6.3. Promoter constructs
Most productivity studies for ethanol in
A number of controllable promoters have also been analyzed [37] with the most useful being the Ni++ Co++ inducible, P
6.4. Knockout of competing pathways as an aid to greater production
Manipulation of carbon flux within the cell factory
Increasing levels of substrate, in this case pyruvate, have also been used to increase yield in metabolic engineered strains. Expressing the enzyme pyruvate kinase (PK), which transfers a phosphate group from PEP to ADP forming Pyruvate [51], has been shown to increase flux to product [52]. Thus, there appears to be some potential for manipulating the flux pathways to and from pyruvate as a means of increasing product yield, which may prove useful when coupled to ethanol production.
6.5. Mutagenesis strategies
Mutagenesis and mutant selection has been developed in
6.6. Improving carbon capture
Several mechanisms of carbon accumulation have been described to operate in
6.7. Neutral sites for integration
As integrative vectors, which utilize homologous recombination into the chromosome, are widespread when metabolic engineering
6.8. Replicative plasmids
Replicative plasmids have been utilized for genetic engineering in
6.9. Tolerance to ethanol
For high level, production of ethanol within
Tolerance has also been examined via transcriptomic analysis following exogenous ethanol addition [67] with 1.2–3% ethanol addition to wild type
7. Linking metabolic engineering of Synechocystis to production
While progress is being made with metabolic engineering for ethanol production and establishing
7.1. Overall process life cycle analysis
Implementation of an industrial process for ethanol production from cyanobacteria will be the next stage of development once the challenges of metabolic engineering have been addressed. Development of the downstream aspects of production will require optimization of several parameters and a more favorable economic outlook. Capital expenditure (CapEx) will be a key driving force with many components needing to be considered. Chief amongst these is the nature of the producing organisms being a recombinant strain. This poses potential safety and containment considerations, which would add to the economics of plant construction and operation. The need for sunlight (which may limit location of production facilities) or continuous LCD exposure again adds costs with either cyclic day exposure in high light climates or continuous growth with added light, which would come with an added energy cost. Equally, calculations of volumes that would be needed suggest large CapEx expenditure on plant, large water requirements and effluent processing costs. Many geographical areas that have high sunlight with marginal land, such as desert areas, at first sight might seem suitable but will suffer from water limitations. Other issues that are related to CapEx relate to the growth of the production strains themselves and the provision of optimal conditions for growth and production. Currently as one diverts photosynthetic intermediates to ethanol, one is affecting the flux to biomass. The more ethanol that is produced the slower the growth and the less biomass that can be produced. This impinges significantly on the growth rate and hence competitiveness of production strains. Given that, growth under sterile conditions in photobioreactors would be economically unsustainable (due to cost and the low value of the product ethanol); competitor contamination would need to be built into the growth cycle. Thus, slow growth of producers would have two major potential consequences that could affect the process. Firstly, there may be mutational selection for faster growers, which have lost the engineered ethanol cassette reducing the yield during production, and secondly given that axenic conditions could not be maintained during aseptic but non-sterile culturing, contaminants could easily outgrow the engineered strains. Strategies that might mitigate this could be the addition of mutualistic consortia, which might stimulate the production strains by providing vitamins or co-factors while limiting the growth of contaminants [68]. Thus, strategies that would aid production at large scale would need to be factored in at the initial stages of metabolic engineering.
7.2. Reactor design for large-scale economic production
The need for significant scale up of photoautotrophic ethanol production in a high light environment can add significantly to initial CapEx. Within the reactor system itself, several components may need significant attention. It is impractical for low value ethanol products, at least in comparison to current fuel costs, for growth and production to be carried out in sterile photobioreactors (PBRs) with full control over light, and key physiological conditions. Although the technologies for such photobioreactors are well developed their practicality can reasonably only be considered suitable for high value products [69]. In addition to containment issues, there are issues with inoculum development for non-axenic culturing to insure that initial inoculum is stable, productive and clonal. Depending on the plant size, this may require significant CapEx.
The most frequent types of PBRs proposed are non-sterile horizontal tubular or vertical flat panel PBRs, which have several limitations including: (a) cost, which have been estimated at €2400 m2 for small scale, reducing in cost slightly with scale [70]. This would result in a cost of some €12.6 kg−1 [71], (b) High energy consumption [72] from mixing, CO2 supply, pumping, separations, cleaning, and (c) Maintenance, cleaning and labor costs [70], (d) The reactor design must be able to withstand photo-oxidation, prevent evaporative loss of product, while maintaining axenic conditions as long as possible. Given the generally slow growth, rates of cyanobacterial species, largely because of the photoautotrophic lifestyle, need to manufacture most of their metabolites, maintain a polyploid genome because of the high sunlight and UV exposure the design of PBRs suitable for low value ethanol production from cyanobacteria is a challenge.
In production terms once one moves away from a controlled PBR design one halves the production cycle and level of photosynthetic production due to the night-day diurnal cycle and in addition there is less process control over the operation. Many approaches have been taken in an attempt to reduce cost; this has included use of bicarbonate-based systems for supply of carbon following carbon capture [73]. This may have significant cost savings in terms of CO2 sparging, transport costs and CO2 loss due to outgassing. Bag type culturing [70] which can be once off or be reusable can offer another potential solution. This may mitigate against some of the limitations of more traditional PBRs. Controlling contaminants in non-axenic culture might be carried out by use of pH as a control mechanism for limiting contamination, however this may necessitate use of more alkaliphilic cyanobacterial species [73]. Indeed adapting the production strain to the process or vice versa may offer a way forward in developing optimal reactor configurations with reduced CapEx. Thus incorporating knowledge of the production cycle, the types of conditions required for growth into a metabolic engineering strategy can be important during initial development of strains and strategies.
7.3. Temperature control, energy and evaporative loss
By virtue of the fact that ethanol-producing cyanobacteria will be recombinant strains, the current experimental systems tend to be enclosed due to regulatory constraints with GMO’s. In geographical locations which are suitable for maximal sunlight and hence photosynthesis, enclosing a facility may raise issues with temperature control unless this is designed into the build. Direct exposure to air circulation or venting may also not be feasible due to safety issues while heat buildup beyond optimal growth temperatures, such as 30°C for
Thus to ensure maximal production and recovery of ethanol, systems may need to be engineered to trap and recover ethanol during production which again may add considerably to CapEx. Jorquera et al. estimated, in a comparative analysis of power consumption of different photosynthetic reactors that horizontal tubular PBRs consumed 2500 W.m3, which reduced to 54 W.m3 for flat panel PBRs and to 3.7 W.m3 for raceway systems [74]. However mixing rates are quite different in the different systems such that in raceway systems there is little mixing, which effects movement of producing cells into light and poor mass transfer limiting overall productivity. Thus, power consumption unless linked to wind or solar in an integrated way may be a key hurdle to overall process efficiency and economy.
7.4. Ethanol recovery from production media
Lignocellulose based fermentations tend to be more dilute than starch based systems due to the presence of hemicellulose which increases viscosity and the presence of fermentation based inhibitors [75, 76]. This is currently similar in terms of cyanobacterial production of ethanol, which is also dilute and low in terms of yield. Recovery of ethanol from dilute production streams in an energy efficient and economical manner poses significant technical difficulties. Traditionally ethanol is recovered via distillation, however in the case of biofuel ethanol from cyanobacterial production the energy costs of distillation would be far too high particularly from dilute streams. It has been estimated that in a well-integrated lignocellulose to ethanol plant the process would require 4350 MJ.m−3 equivalent to approximately 20% of the energy content of the ethanol produced [76]. Thus, alternatives to distillation are needed to drive economy from cyanobacterial production systems. A number of techniques are available which may be suitable for the recovery of ethanol from cyanobacterial production such as membrane permeation or pervaporation, vacuum stripping, gas stripping, solvent extraction, adsorption and various hybrid processes [76]. However, the efficiency is dependent on the initial ethanol concentration (which is currently low for metabolically engineered
8. Conclusions and perspectives
The basic proof of concept for photoautotrophic ethanol production from model cyanobacteria such as
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