How long does it take for E. coli to shift feedstocks?

How long does it take for E. coli to shift feedstocks?

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With our fermentations we're noticing that it takes an appreciable amount of time for E. coli (K12 variant) to change from being metabolically streamlined on amino acids to being metabolically streamlined to glucose.

How long does it take for such microorganisms to adjust to a new feedstock?

This must depend upon the conditions in question, but I think it would not be very long. The length of a generation for e coli can be 12 minutes or 24 hours, so that gives some idea of a typical time.

I did find this interesting case in the literature. They found that even as you subject e coli to minimal nutrients they are expressing genes that prepare them for a richer nutrient mix. The bacteria are primed for a flood of carbohydrates succinate or amino acids.

This study incubated the e coli overnight before taking their readings, so this puts an upper bound of about 12 hours. The implication is that if you are moving the bacteria to rich media it might be just a short time before they are in log phase growth.

In practice moving from LB plates to log phase growth is usually just like 4 hours. This paper interestingly shows that cold adapted E coli moving to other media react in a few hours, but can show behavioral changes days out.

I would say that 4 hours might be a typical expectation. 12 is typically used in the literature.

Chapter 6 Normal Flora

Significance of the Normal Flora

The normal flora influences the anatomy, physiology, susceptibility to pathogens, and morbidity of the host.

Skin Flora

The varied environment of the skin results in locally dense or sparse populations, with Gram-positive organisms (e.g., staphylococci, micrococci, diphtheroids) usually predominating.

Oral and Upper Respiratory Tract Flora

A varied microbial flora is found in the oral cavity, and streptococcal anaerobes inhabit the gingival crevice. The pharynx can be a point of entry and initial colonization for Neisseria, Bordetella, Corynebacterium, and Streptococcus spp.

Gastrointestinal Tract Flora

Organisms in the stomach are usually transient, and their populations are kept low (10 3 to 10 6 /g of contents) by acidity. Helicobacter pylori is a potential stomach pathogen that apparently plays a role in the formation of certain ulcer types. In normal hosts the duodenal flora is sparse (0 to 10 3 /g of contents). The ileum contains a moderately mixed flora (10 6 to 10 8 /g of contents). The flora of the large bowel is dense (10 9 to 10 11 /g of contents) and is composed predominantly of anaerobes. These organisms participate in bile acid conversion and in vitamin K and ammonia production in the large bowel. They can also cause intestinal abscesses and peritonitis.

Urogenital Flora

The vaginal flora changes with the age of the individual, the vaginal pH, and hormone levels. Transient organisms (e.g., Candida spp.) frequently cause vaginitis. The distal urethra contains a sparse mixed flora these organisms are present in urine specimens (10 4 /ml) unless a clean-catch, midstream specimen is obtained.

Conjunctival Flora

The conjunctiva harbors few or no organisms. Haemophilus and Staphylococcus are among the genera most often detected.

Host Infection

Many elements of the normal flora may act as opportunistic pathogens, especially in hosts rendered susceptible by rheumatic heart disease, immunosuppression, radiation therapy, chemotherapy, perforated mucous membranes, etc. The flora of the gingival crevice causes dental caries in about 80 percent of the population.

Protein Expression and Purification Core Facility

To speed up protein production, we have adopted a strategy of parallel expression of a protein from a variety of vectors containing different tags and/or fusion partners, and a variety of E. coli host strains. This approach should not only gain us a lot of time but also result in a larger number of successfully expressed proteins.

The expression strategy consists of the following two sets of experiments:

1. The expression of a protein in a basic E. coli host strain from a variety vectors with different tags and/or fusion partners.

Our first screen is to express a protein in BL21 (DE3) from modified pET-vectors with the following a selection of tags and fusion partners:

Tag fusion partner
N-terminal His6-tag
N-terminal His6-tag thioredoxin
N-terminal His6-tag glutathione-S-transferase (GST)
N-terminal His6-tag maltose binding protein (MBP)
N-terminal His6-tag disulfide oxidoreductase (DsbA)
N-terminal His6-tag NusA
C-terminal His6-tag

2. The expression of a protein from a standard vector in a number of different E. coli host strains.

The choice of the host strains depends more on the nature of the heterologous protein. The following considerations should be made:

  • If the protein contains a high number of rare E. coli codons, it is worthwhile trying to express it in a strain that co-expresses the tRNAs for these rare codons. There are several strains commercially available:
  • If the protein contains one or more disulfide bonds, proper folding is stimulated in host strain with a more oxidizing cytoplasmic environment. Two strains are commercially available from Novagen:

  • If the protein is toxic to the cell, expression in a strain containing the pLysS or pLysE vector tightens regulation of expression systems using the T7 promoter. These vectors express lysozyme, which binds to and inactivates T7 RNA polymerase. Strains are commercially available from different manufacturers.

Our first screen is to express a protein from a modified pET-vector with an N-terminal His6-tag in the following host strains:

Host strain
BL21 (DE3)
BL21 (DE3) pLysS
BL21 (DE3) CodonPlus-RIL (-RP) or Rosetta (DE3)
Origami (DE3)

For the rapid screening of expression levels and protein solubility from the different vectors and in the different strains we have developed a small scale method using chromatography on magnetic beads.

The results of the first screen will be a starting point for further experiments in case no satisfactory expression conditions have been found. How to optimise expression levels, improve protein solubility, improve protein stability, and decrease protein toxicity will be discussed in the other chapters in this website.

Expression method

A typical expression experiment consists of the following step:

  • Picking of a single colony from a freshly streaked plate of the expression host containing the recombinant vector. When the heterologous protein is toxic for the cells, higher expression levels are obtained by using the so-called "plating" method.
  • Growing of a starter culture. Inoculate with the picked colony up to 50 ml of rich medium (such as LB or 2xYT) containing the appropriate antibiotic. When a larger starter culture is required, inoculate 4 ml of rich media with the single colony grow for 4-8 hours at 37°C and use this to inoculate the starter culture.

Do not let cultures grow at 37°C overnight! It is better to grow overnight cultures at 30°C or lower. Alternatively, the culture can be incubated at 37°C until the OD600 is approx. 1. Then store the culture at 4°C overnight. The following morning, collect the cells by centrigfugation, resuspend them in fresh medium and use this to inoculate the main culture.

The use of ampicillin requires special care. The selectable marker, b -lactamase, is secreted into the medium where it hydrolysis all of the ampicillin. This point is already reached when the culture is barely turbid. From here on, cells that lack the plasmid will not be killed and could overgrow the culture (which can be tested using a plasmid stability test). Some possible solutions are:

  • grow overnight cultures at 30°C or lower.
  • spin overnight cultures and resuspend the pellet in fresh medium to remove b -lactamase.
  • use the more stable carbenicillin instead of ampicillin.
  • Inoculation of the main culture and incubation until OD600 reaches 0.4-1. The optimal OD value depends on the culture method and the medium. For flask cultures using LB-medium an OD600of 0.6 is recommended. To increase the growth rate, we carry out the cultures at 37°C until the OD for induction is reached. Then the cultures are cooled to the induction temperature in ice-water.

Remark: For good aeration, don't use more medium than 20% of the total flask volume.

  • Induction of protein expression. Protein expression is induced by the addition of the proper inducer or by changing the growth conditions. From this point on the cells will use most of their resources for the production of the target protein and will not grow much further.

For the most used promoters induction conditions are listed below.

Promoter induction typical condition range
trc (hybrid) addition of IPTG 0.2 mM 0.05 - 2.0 mM
araBAD addition of l-arabinose 0.2% 0.002 - 0.4 %
PL shifting the temperature from 37 to 42°C
T7-lac operator addition of IPTG 0.2 mM 0.05 -2.0 mM

After induction the cultures are incubated from 3 hours to overnight depending on the induction temperature. Guide lines are given below.


In general, proteins possess more or less the ability to coordinate metal ions on their surface, and it is possible to separate proteins by chromatography making use of the difference in their affinity. This is the immobilized metal ion affinity chromatography announced in 1975. [2] Subsequent studies have revealed that among amino acids constituting proteins, histidine is strongly involved in the coordinate bond with metal ions. [3] Therefore, if a number of histidines are added to the end of the protein by genetic engineering, the affinity of the protein for the metal ion is remarkably increased and the basic idea is that purification can be easily carried out. When a protein having a His-tag is brought into contact with a carrier on which a metal ion such as nickel is immobilized under the condition of pH 8 or higher, the histidine residue chelates the metal ion and binds to the carrier. Since other proteins do not bind to the carrier or bind only very weakly, they can be removed by washing the carrier with an appropriate buffer. Thereafter, by removing imidazole or the like from the carrier, it is possible to recover the protein having the His-tag with high purity.

Practical choice Edit

Carrier Edit

Various carriers such as Ni - NTA agarose (nickel - nitrilotriacetic acid) are on the market. It is packed in a column and used in combination with centrifugation and magnetic separation in a test tube.

Metal ions Edit

As the metal ion, copper has the highest affinity, and the affinity decreases in the order of nickel, zinc, and cobalt [ citation needed ] . Nickel is often used for ordinary purposes, and cobalt is used when it is desired to increase the purity of purification.

Elution method Edit

In order to elute His-tagged protein from the carrier, there is a plurality of methods as follows and it will be used properly according to the purpose. In order to avoid denaturation of proteins, it is desirable to have as mild as possible, and imidazole addition is often used from this viewpoint.

Competition with analogs Edit

When a compound having a structure similar to the histidine residue is added at a high concentration, the protein competes with the coordination of the metal ion, so that the protein is separated from the carrier. Imidazole is a compound constituting the side chain of histidine and is frequently used at a concentration of 150 mM or more. In addition, histidine and histamine may be used in some cases.

Decrease in pH Edit

When the pH decreases, the histidine residue is protonated and can no longer coordinate the metal tag, allowing the protein to be eluted. When nickel is used as the metal ion, it is eluted at around 4 and cobalt at around 6.

Removal of metal ions Edit

When a strong chelating agent is added, the protein is detached from the carrier because the metal ion immobilized on the carrier is lost. EDTA is used exclusively.

Protein purification Edit

Polyhistidine-tags are often used for affinity purification of polyhistidine-tagged recombinant proteins expressed in Escherichia coli [4] and other prokaryotic expression systems. Bacterial cells are harvested via centrifugation and the resulting cell pellet lysed either by physical means or by means of detergents and enzymes such as lysozyme or any combination of these. At this stage, raw lysate contains the recombinant protein among many other proteins originating from the bacterial host. This mixture is incubated with an affinity resin containing bound divalent nickel or cobalt ions, which are available commercially in different varieties. Nickel and cobalt have similar properties and as they are adjacent period 4 transition metals (v. iron triad). These resins are generally sepharose/agarose functionalized with a chelator, such as iminodiacetic acid (Ni-IDA) and nitrilotriacetic acid (Ni-NTA) for nickel and carboxyl-methyl aspartate (Co-CMA) for cobalt, which the polyhistidine-tag binds with micromolar affinity. Ernst Hochuli et al. coupled 1987 the NTA ligand and Nickel-ions to agarose beads. [5] The resin is then washed with phosphate buffer to remove proteins that do not specifically interact with the cobalt or nickel ion. With Ni-based methods, washing efficiency can be improved by the addition of 20 mM imidazole (proteins are usually eluted with 150-300 mM imidazole). Generally, nickel-based resins have a higher binding capacity, while cobalt-based resins offer the highest purity. The purity and amount of protein can be assessed by SDS-PAGE and Western blotting. [ citation needed ]

Affinity purification using a polyhistidine-tag usually results in relatively pure protein when the recombinant protein is expressed in prokaryotic organisms. Depending on downstream applications, including the purification of protein complexes to study protein interactions, purification from higher organisms such as yeasts or other eukaryotes may require a tandem affinity purification [6] using two tags to yield higher purity. Alternatively, single-step purification using immobilized cobalt ions rather than nickel ions generally yields a substantial increase in purity and requires lower imidazole concentrations for elution of the his-tagged protein.

Polyhistidine-tagging is the option of choice for purifying recombinant proteins in denaturing conditions because its mode of action is dependent only on the primary structure of proteins. For example, even when a recombinant protein forcibly expressed in E. coli produces an inclusion body and can not be obtained as a soluble protein, it can be purified with denaturation with urea or guanidine hydrochloride. Generally, for this sort of a technique, histidine binding is titrated using pH instead of imidazole binding—at a high pH, histidine binds to nickel or cobalt, but at low pH (

4 for nickel), histidine becomes protonated and is competed off of the metal ion. Compare this to antibody purification and GST purification, a prerequisite to which is the proper (native) folding of proteins involved. On the other hand, it is said that the His tag tends to aggregate and insolubilize more than other affinity tags.

Polyhistidine-tag columns retain several well known proteins as impurities. One of them is FKBP-type peptidyl prolyl isomerase, which appears around 25kDa (SlyD). Impurities are generally eliminated using a secondary chromatographic technique, or by expressing the recombinant protein in a SlyD-deficient E. coli strain. [7] Alternatively, comparing with nickel-based, cobalt-based resins have less affinity with SlyD from E. coli, but in several cases, it is moderately helpful. [8]

Separating one from two polyhistidine tags Edit

Proteins with different numbers of polyhistidine tags elute differently from nickel-affinity resin. For proteins with a single hexahistidine tag, 75 mM imidazole enables elution from Ni-NTA, whereas for proteins with two hexahistidine tags, 100 mM imidazole is required for elution. [ citation needed ] This step-wise elution may be used to isolate specific protein assemblies from a mixture, such as defined heteromultimers (e.g. an AB heterodimer from a mixture including AA and BB homodimers, if only subunit B has a polyhistidine tag). Such an approach was used in isolation of monovalent streptavidin. [9]

Binding assays Edit

Polyhistidine-tagging can be used to detect protein-protein interactions in the same way as a pull-down assay. However, this technique is generally considered to be less sensitive, and also restricted by some of the more finicky aspects of this technique. For example, reducing conditions cannot be used, EDTA and many types of detergents cannot be used. Recent advances in dual polarisation interferometry is amenable to EDTA and a wider use of reagents, and the use of such site-specific tags greatly simplifies the direct measurement of associated conformational change. [ citation needed ]

Fluorescent tags Edit

Hexahistadine CyDye tags have also been developed. These use Nickel covalent coordination to EDTA groups attached to fluorophores in order to create dyes that attach to the polyhistidine tag. This technique has been shown to be effective for following protein migration and trafficking. There has also been recent discoveries that show this technique may be effective in order to measure distance via Fluorescent Resonance Energy Transfer. [10]

Fluorohistidine tags Edit

A polyfluorohistidine tag has been reported for use in in vitro translation systems. [11] In this system, an expanded genetic code is used in which histidine is replaced by 4-fluorohistidine. The fluorinated analog is incorporated into peptides via the relaxed substrate specificity of histidine-tRNA ligase and lowers the overall pKa of the tag. This allows for the selective enrichment of polyfluorohistidine tagged peptides in the presence of complex mixtures of traditional polyhistidine tags by altering the pH of the wash buffers.

The most common polyhistidine tags are formed of six histidine (6xHis tag) residues - which are added at the N-terminus preceded by Methionine or C-terminus before a stop codon, in the coding sequence of the protein of interest. The choice of the end where His-tag is added will depend mainly on the characteristics of the protein and the methods chosen to remove the tag. Some ends are buried inside the protein core and others are important for the protein function or structure. In those cases the choice is limited to the other end. On the other hand, most available exopeptidases can only remove the His-tag from the N-terminus removing the tag from the C-terminus will require the use of other techniques. It is important to take into account that the computer simulation (by molecular dynamics) will help you to choose between options, for example, whether the His-tag must be digested or engineered to the N- or C-terminal. [12]

There are two ways to add polyhistidines. The most simple is to insert the DNA encoding the protein in a vector encoding a His-tag so that it will be automatically attached to one of its ends (See picture). Another technique is to perform a PCR with primers that have repetitive histidine codons (CAT or CAC) right next to the START or STOP codon in addition to several (16 or more) bases from one end of the DNA encoding the protein to be tagged (see primer example below). [ citation needed ]

Example of primer designed to add a 6xHis-tag using PCR. Eighteen bases coding six histidines are inserted right after the START codon or right before the STOP codon. At least 16 bases specific to the gene of interest are needed next to the His-tag. With 6 His, the protein will have an added 1 kDa of molecular weight. Often, a linker (such as gly-gly-gly or gly-ser-gly) is placed between the protein of interest and the 6 His tag in order to prevent the polyhistidine tag from affecting the activity of the protein being tagged. [ citation needed ]

The polyhistidine-tag can also be used to detect the protein via anti-polyhistidine-tag antibodies or alternatively by in-gel staining (SDS-PAGE) with fluorescent probes bearing metal ions. This can be useful in subcellular localization, ELISA, western blotting or other immuno-analytical methods. [ citation needed ]

The polyhistidine-tag can be successfully used for the immobilization of proteins on a surface such as on a nickel- or cobalt-coated microtiter plate or on a protein array. [13]

HQ tag Edit

The HQ tag has alternating histidine and glutamine (HQHQHQ).

HN tag Edit

The HN tag has alternating histidine and asparagine (HNHNHNHNHNHN) and is more likely to be presented on the protein surface than Histidine-only tags. The HN tag binds to the immobilized metal ion more efficiently than the His tag. [14]

HAT tag Edit

The HAT tag is a peptide tag (KDHLIHNVHKEEHAHAHNK) derived from chicken lactate dehydrogenase, and is more likely to be a soluble protein with no bias in charge distribution compared to the His tag. [15] The arrangement of histidines in the HAT tag allows high accessibility compared to the His tag, and it binds efficiently to the immobilized metal ion.


The process was discovered by Joshua Lederberg and Edward Tatum [9] in 1946.

  1. Donor cell produces pilus.
  2. Pilus attaches to recipient cell and brings the two cells together.
  3. The mobile plasmid is nicked and a single strand of DNA is then transferred to the recipient cell.
  4. Both cells synthesize a complementary strand to produce a double stranded circular plasmid and also reproduce pili both cells are now viable donor for the F-factor. [1]

The F-plasmid is an episome (a plasmid that can integrate itself into the bacterial chromosome by homologous recombination) with a length of about 100 kb. It carries its own origin of replication, the oriV, and an origin of transfer, or oriT. [4] There can only be one copy of the F-plasmid in a given bacterium, either free or integrated, and bacteria that possess a copy are called F-positive or F-plus (denoted F + ). Cells that lack F plasmids are called F-negative or F-minus (F − ) and as such can function as recipient cells.

Among other genetic information, the F-plasmid carries a tra and trb locus, which together are about 33 kb long and consist of about 40 genes. The tra locus includes the pilin gene and regulatory genes, which together form pili on the cell surface. The locus also includes the genes for the proteins that attach themselves to the surface of F − bacteria and initiate conjugation. Though there is some debate on the exact mechanism of conjugation it seems that the pili are not the structures through which DNA exchange occurs. This has been shown in experiments where the pilus are allowed to make contact, but then are denatured with SDS and yet DNA transformation still proceeds. Several proteins coded for in the tra or trb locus seem to open a channel between the bacteria and it is thought that the traD enzyme, located at the base of the pilus, initiates membrane fusion.

When conjugation is initiated by a signal the relaxase enzyme creates a nick in one of the strands of the conjugative plasmid at the oriT. Relaxase may work alone or in a complex of over a dozen proteins known collectively as a relaxosome. In the F-plasmid system the relaxase enzyme is called TraI and the relaxosome consists of TraI, TraY, TraM and the integrated host factor IHF. The nicked strand, or T-strand, is then unwound from the unbroken strand and transferred to the recipient cell in a 5'-terminus to 3'-terminus direction. The remaining strand is replicated either independent of conjugative action (vegetative replication beginning at the oriV) or in concert with conjugation (conjugative replication similar to the rolling circle replication of lambda phage). Conjugative replication may require a second nick before successful transfer can occur. A recent report claims to have inhibited conjugation with chemicals that mimic an intermediate step of this second nicking event. [10]

If the F-plasmid that is transferred has previously been integrated into the donor's genome (producing an Hfr strain ["High Frequency of Recombination"]) some of the donor's chromosomal DNA may also be transferred with the plasmid DNA. [3] The amount of chromosomal DNA that is transferred depends on how long the two conjugating bacteria remain in contact. In common laboratory strains of E. coli the transfer of the entire bacterial chromosome takes about 100 minutes. The transferred DNA can then be integrated into the recipient genome via homologous recombination.

A cell culture that contains in its population cells with non-integrated F-plasmids usually also contains a few cells that have accidentally integrated their plasmids. It is these cells that are responsible for the low-frequency chromosomal gene transfers that occur in such cultures. Some strains of bacteria with an integrated F-plasmid can be isolated and grown in pure culture. Because such strains transfer chromosomal genes very efficiently they are called Hfr (high frequency of recombination). The E. coli genome was originally mapped by interrupted mating experiments in which various Hfr cells in the process of conjugation were sheared from recipients after less than 100 minutes (initially using a Waring blender). The genes that were transferred were then investigated.

Since integration of the F-plasmid into the E. coli chromosome is a rare spontaneous occurrence, and since the numerous genes promoting DNA transfer are in the plasmid genome rather than in the bacterial genome, it has been argued that conjugative bacterial gene transfer, as it occurs in the E. coli Hfr system, is not an evolutionary adaptation of the bacterial host, nor is it likely ancestral to eukaryotic sex. [13]

Spontaneous zygogenesis in E. coli

In addition to classical bacterial conjugation described above for E. coli, a form of conjugation referred to as spontaneous zygogenesis (Z-mating for short) is observed in certain strains of E. coli. [6] In Z-mating there is complete genetic mixing, and unstable diploids are formed that throw off phenotypically haploid cells, of which some show a parental phenotype and some are true recombinants.

Conjugation in Mycobacteria smegmatis, like conjugation in E. coli, requires stable and extended contact between a donor and a recipient strain, is DNase resistant, and the transferred DNA is incorporated into the recipient chromosome by homologous recombination. However, unlike E. coli Hfr conjugation, mycobacterial conjugation is chromosome rather than plasmid based. [7] [8] Furthermore, in contrast to E. coli Hfr conjugation, in M. smegmatis all regions of the chromosome are transferred with comparable efficiencies. The lengths of the donor segments vary widely, but have an average length of 44.2kb. Since a mean of 13 tracts are transferred, the average total of transferred DNA per genome is 575kb. [8] This process is referred to as "Distributive conjugal transfer." [7] [8] Gray et al. [7] found substantial blending of the parental genomes as a result of conjugation and regarded this blending as reminiscent of that seen in the meiotic products of sexual reproduction.

Bacteria related to the nitrogen fixing Rhizobia are an interesting case of inter-kingdom conjugation. [14] For example, the tumor-inducing (Ti) plasmid of Agrobacterium and the root-tumor inducing (Ri) plasmid of A. rhizogenes contain genes that are capable of transferring to plant cells. The expression of these genes effectively transforms the plant cells into opine-producing factories. Opines are used by the bacteria as sources of nitrogen and energy. Infected cells form crown gall or root tumors. The Ti and Ri plasmids are thus endosymbionts of the bacteria, which are in turn endosymbionts (or parasites) of the infected plant.

The Ti and Ri plasmids can also be transferred between bacteria using a system (the tra, or transfer, operon) that is different and independent of the system used for inter-kingdom transfer (the vir, or virulence, operon). Such transfers create virulent strains from previously avirulent strains.

Conjugation is a convenient means for transferring genetic material to a variety of targets. In laboratories, successful transfers have been reported from bacteria to yeast, [15] plants, mammalian cells, [16] [17] diatoms [18] and isolated mammalian mitochondria. [19] Conjugation has advantages over other forms of genetic transfer including minimal disruption of the target's cellular envelope and the ability to transfer relatively large amounts of genetic material (see the above discussion of E. coli chromosome transfer). In plant engineering, Agrobacterium-like conjugation complements other standard vehicles such as tobacco mosaic virus (TMV). While TMV is capable of infecting many plant families these are primarily herbaceous dicots. Agrobacterium-like conjugation is also primarily used for dicots, but monocot recipients are not uncommon.


Cellulosic ethanol is a type of biofuel produced from lignocellulose, a structural material that comprises much of the mass of plants and is composed mainly of cellulose, hemicellulose and lignin. Popular sources of lignocellulose include both agricultural waste products (e.g. corn stover or wood chips) and grasses like switchgrass and miscanthus species. [1] These raw materials for ethanol production have the advantage of being abundant and diverse and would not compete with food production, unlike the more commonly used corn and cane sugars. [2] However, they also require more processing to make the sugar monomers available to the microorganisms typically used to produce ethanol by fermentation, which drives up the price of cellulos-derived ethanol. [3]

Cellulosic ethanol can reduce greenhouse gas emissions by 85% over reformulated gasoline. [4] By contrast, starch ethanol (e.g., from corn), which most frequently uses natural gas to provide energy for the process, may not reduce greenhouse gas emissions at all depending on how the starch-based feedstock is produced. [5] According to the National Academy of Sciences in 2011, there is no commercially viable bio-refinery in existence to convert lignocellulosic biomass to fuel. [6] Absence of production of cellulosic ethanol in the quantities required by the regulation was the basis of a United States Court of Appeals for the District of Columbia decision announced January 25, 2013, voiding a requirement imposed on car and truck fuel producers in the United States by the Environmental Protection Agency requiring addition of cellulosic biofuels to their products. [7] These issues, along with many other difficult production challenges, led George Washington University policy researchers to state that "in the short term, [cellulosic] ethanol cannot meet the energy security and environmental goals of a gasoline alternative." [8]

The French chemist, Henri Braconnot, was the first to discover that cellulose could be hydrolyzed into sugars by treatment with sulfuric acid in 1819. [9] The hydrolyzed sugar could then be processed to form ethanol through fermentation. The first commercialized ethanol production began in Germany in 1898, where acid was used to hydrolyze cellulose. In the United States, the Standard Alcohol Company opened the first cellulosic ethanol production plant in South Carolina in 1910. Later, a second plant was opened in Louisiana. However, both plants were closed after World War I due to economic reasons. [10]

The first attempt at commercializing a process for ethanol from wood was done in Germany in 1898. It involved the use of dilute acid to hydrolyze the cellulose to glucose, and was able to produce 7.6 liters of ethanol per 100 kg of wood waste (18 US gal (68 L) per ton). The Germans soon developed an industrial process optimized for yields of around 50 US gallons (190 L) per ton of biomass. This process soon found its way to the US, culminating in two commercial plants operating in the southeast during World War I. These plants used what was called "the American Process" — a one-stage dilute sulfuric acid hydrolysis. Though the yields were half that of the original German process (25 US gallons (95 L) of ethanol per ton versus 50), the throughput of the American process was much higher. A drop in lumber production forced the plants to close shortly after the end of World War I. In the meantime, a small but steady amount of research on dilute acid hydrolysis continued at the USFS's Forest Products Laboratory. [11] [12] [13] During World War II, the US again turned to cellulosic ethanol, this time for conversion to butadiene to produce synthetic rubber. The Vulcan Copper and Supply Company was contracted to construct and operate a plant to convert sawdust into ethanol. The plant was based on modifications to the original German Scholler process as developed by the Forest Products Laboratory. This plant achieved an ethanol yield of 50 US gal (190 L) per dry ton, but was still not profitable and was closed after the war. [14]

With the rapid development of enzyme technologies in the last two decades, the acid hydrolysis process has gradually been replaced by enzymatic hydrolysis. Chemical pretreatment of the feedstock is required to hydrolyze (separate) hemicellulose, so it can be more effectively converted into sugars. The dilute acid pretreatment is developed based on the early work on acid hydrolysis of wood at the USFS's Forest Products Laboratory. Recently, the Forest Products Laboratory together with the University of Wisconsin–Madison developed a sulfite pretreatment to overcome the recalcitrance of lignocellulose for robust enzymatic hydrolysis of wood cellulose. [15]

In his 2007 State of the Union Address on January 23, 2007, US President George W. Bush announced a proposed mandate for 35 billion US gallons (130 × 10 ^ 9 L) of ethanol by 2017. Later that year, the US Department of Energy awarded $385 million in grants aimed at jump-starting ethanol production from nontraditional sources like wood chips, switchgrass, and citrus peels. [16]

The stages to produce ethanol using a biological approach are: [17]

  1. A "pretreatment" phase to make the lignocellulosic material such as wood or straw amenable to hydrolysis
  2. Cellulose hydrolysis (cellulolysis) to break down the molecules into sugars
  3. Microbial fermentation of the sugar solution
  4. Distillation and dehydration to produce pure alcohol

In 2010, a genetically engineered yeast strain was developed to produce its own cellulose-digesting enzymes. [18] Assuming this technology can be scaled to industrial levels, it would eliminate one or more steps of cellulolysis, reducing both the time required and costs of production. [ citation needed ]

Although lignocellulose is the most abundant plant material resource, its usability is curtailed by its rigid structure. As a result, an effective pretreatment is needed to liberate the cellulose from the lignin seal and its crystalline structure so as to render it accessible for a subsequent hydrolysis step. [19] By far, most pretreatments are done through physical or chemical means. To achieve higher efficiency, both physical and chemical pretreatments are required. Physical pretreatment involves reducing biomass particle size by mechanical processing methods such as milling or extrusion. Chemical pretreatment partially depolymerizes the lignocellulose so enzymes can access the cellulose for microbial reactions. [20]

Chemical pretreatment techniques include acid hydrolysis, steam explosion, ammonia fiber expansion, organosolv, sulfite pretreatment, [15] AVAP® (SO2-ethanol-water) fractionation, [21] alkaline wet oxidation and ozone pretreatment. [22] Besides effective cellulose liberation, an ideal pretreatment has to minimize the formation of degradation products because they can inhibit the subsequent hydrolysis and fermentation steps. [23] The presence of inhibitors further complicates and increases the cost of ethanol production due to required detoxification steps. For instance, even though acid hydrolysis is probably the oldest and most-studied pretreatment technique, it produces several potent inhibitors including furfural and hydroxymethylfurfural. [24] Ammonia Fiber Expansion (AFEX) is an example of a promising pretreatment that produces no inhibitors. [25]

Most pretreatment processes are not effective when applied to feedstocks with high lignin content, such as forest biomass. These require alternative or specialized approaches. Organosolv, SPORL ('sulfite pretreatment to overcome recalcitrance of lignocellulose') and SO2-ethanol-water (AVAP®) processes are the three processes that can achieve over 90% cellulose conversion for forest biomass, especially those of softwood species. SPORL is the most energy efficient (sugar production per unit energy consumption in pretreatment) and robust process for pretreatment of forest biomass with very low production of fermentation inhibitors. Organosolv pulping is particularly effective for hardwoods and offers easy recovery of a hydrophobic lignin product by dilution and precipitation. [26] </ref> AVAP® process effectively fractionates all types of lignocellulosics into clean highly digestible cellulose, undegraded hemicellulose sugars, reactive lignin and lignosulfonates, and is characterized by efficient recovery of chemicals. [27] [28]

Cellulolytic processes Edit

The hydrolysis of cellulose (cellulolysis) produces simple sugars that can be fermented into alcohol. There are two major cellulolysis processes: chemical processes using acids, or enzymatic reactions using cellulases. [17]

Chemical hydrolysis Edit

In the traditional methods developed in the 19th century and at the beginning of the 20th century, hydrolysis is performed by attacking the cellulose with an acid. [29] Dilute acid may be used under high heat and high pressure, or more concentrated acid can be used at lower temperatures and atmospheric pressure. A decrystallized cellulosic mixture of acid and sugars reacts in the presence of water to complete individual sugar molecules (hydrolysis). The product from this hydrolysis is then neutralized and yeast fermentation is used to produce ethanol. As mentioned, a significant obstacle to the dilute acid process is that the hydrolysis is so harsh that toxic degradation products are produced that can interfere with fermentation. BlueFire Renewables uses concentrated acid because it does not produce nearly as many fermentation inhibitors, but must be separated from the sugar stream for recycle [simulated moving bed chromatographic separation, for example] to be commercially attractive. [ citation needed ]

Agricultural Research Service scientists found they can access and ferment almost all of the remaining sugars in wheat straw. The sugars are located in the plant's cell walls, which are notoriously difficult to break down. To access these sugars, scientists pretreated the wheat straw with alkaline peroxide, and then used specialized enzymes to break down the cell walls. This method produced 93 US gallons (350 L) of ethanol per ton of wheat straw. [30]

Enzymatic hydrolysis Edit

Cellulose chains can be broken into glucose molecules by cellulase enzymes. This reaction occurs at body temperature in the stomachs of ruminants such as cattle and sheep, where the enzymes are produced by microbes. This process uses several enzymes at various stages of this conversion. Using a similar enzymatic system, lignocellulosic materials can be enzymatically hydrolyzed at a relatively mild condition (50 °C and pH 5), thus enabling effective cellulose breakdown without the formation of byproducts that would otherwise inhibit enzyme activity. All major pretreatment methods, including dilute acid, require an enzymatic hydrolysis step to achieve high sugar yield for ethanol fermentation. [25]

Fungal enzymes can be used to hydrolyze cellulose. The raw material (often wood or straw) still has to be pre-treated to make it amenable to hydrolysis. [31] In 2005, Iogen Coroporation announced it was developing a process using the fungus Trichoderma reesei to secrete "specially engineered enzymes" for an enzymatic hydrolysis process. [32]

Another Canadian company, SunOpta, uses steam explosion pretreatment, providing its technology to Verenium (formerly Celunol Corporation)'s facility in Jennings, Louisiana, Abengoa's facility in Salamanca, Spain, and a China Resources Alcohol Corporation in Zhaodong. The CRAC production facility uses corn stover as raw material. [33]

Microbial fermentation Edit

Traditionally, baker's yeast (Saccharomyces cerevisiae), has long been used in the brewery industry to produce ethanol from hexoses (six-carbon sugars). Due to the complex nature of the carbohydrates present in lignocellulosic biomass, a significant amount of xylose and arabinose (five-carbon sugars derived from the hemicellulose portion of the lignocellulose) is also present in the hydrolysate. For example, in the hydrolysate of corn stover, approximately 30% of the total fermentable sugars is xylose. As a result, the ability of the fermenting microorganisms to use the whole range of sugars available from the hydrolysate is vital to increase the economic competitiveness of cellulosic ethanol and potentially biobased proteins. [ citation needed ]

In recent years, metabolic engineering for microorganisms used in fuel ethanol production has shown significant progress. [34] Besides Saccharomyces cerevisiae, microorganisms such as Zymomonas mobilis and Escherichia coli have been targeted through metabolic engineering for cellulosic ethanol production. An attraction towards alternative fermentation organism is its ability to ferment five carbon sugars improving the yield of the feed stock. This ability is often found in bacteria [35] based organisms. [ citation needed ]

Recently, engineered yeasts have been described efficiently fermenting xylose, [36] [37] and arabinose, [38] and even both together. [39] Yeast cells are especially attractive for cellulosic ethanol processes because they have been used in biotechnology for hundreds of years, are tolerant to high ethanol and inhibitor concentrations and can grow at low pH values to reduce bacterial contamination. [ citation needed ]

Combined hydrolysis and fermentation Edit

Some species of bacteria have been found capable of direct conversion of a cellulose substrate into ethanol. One example is Clostridium thermocellum, which uses a complex cellulosome to break down cellulose and synthesize ethanol. However, C. thermocellum also produces other products during cellulose metabolism, including acetate and lactate, in addition to ethanol, lowering the efficiency of the process. Some research efforts are directed to optimizing ethanol production by genetically engineering bacteria that focus on the ethanol-producing pathway. [40]

Gasification process (thermochemical approach) Edit

The gasification process does not rely on chemical decomposition of the cellulose chain (cellulolysis). Instead of breaking the cellulose into sugar molecules, the carbon in the raw material is converted into synthesis gas, using what amounts to partial combustion. The carbon monoxide, carbon dioxide and hydrogen may then be fed into a special kind of fermenter. Instead of sugar fermentation with yeast, this process uses Clostridium ljungdahlii bacteria. [41] This microorganism will ingest carbon monoxide, carbon dioxide and hydrogen and produce ethanol and water. The process can thus be broken into three steps:

    — Complex carbon-based molecules are broken apart to access the carbon as carbon monoxide, carbon dioxide and hydrogen
  1. Fermentation — Convert the carbon monoxide, carbon dioxide and hydrogen into ethanol using the Clostridium ljungdahlii organism
  2. Distillation — Ethanol is separated from water

A recent study has found another Clostridium bacterium that seems to be twice as efficient in making ethanol from carbon monoxide as the one mentioned above. [42]

Alternatively, the synthesis gas from gasification may be fed to a catalytic reactor where it is used to produce ethanol and other higher alcohols through a thermochemical process. [43] This process can also generate other types of liquid fuels, an alternative concept successfully demonstrated by the Montreal-based company Enerkem at their facility in Westbury, Quebec. [44]

Studies are intensively conducted to develop economic methods to convert both cellulose and hemicellulose to ethanol. Fermentation of glucose, the main product of cellulose hydrolyzate, to ethanol is an already established and efficient technique. However, conversion of xylose, the pentose sugar of hemicellulose hydrolyzate, is a limiting factor, especially in the presence of glucose. Moreover, it cannot be disregarded as hemicellulose will increase the efficiency and cost-effectiveness of cellulosic ethanol production. [45]

Sakamoto (2012) et al. show the potential of genetic engineering microbes to express hemicellulase enzymes. The researchers created a recombinant Saccharomyces cerevisiae strain that was able to:

  1. hydrolyze hemicellulase through codisplaying endoxylanase on its cell surface,
  2. assimilate xylose by expression of xylose reductase and xylitol dehydrogenase.

The strain was able to convert rice straw hydrolyzate to ethanol, which contains hemicellulosic components. Moreover, it was able to produce 2.5x more ethanol than the control strain, showing the highly effective process of cell surface-engineering to produce ethanol. [45]

General advantages of ethanol fuel Edit

Ethanol burns more cleanly and more efficiently than gasoline. [46] [47] Because plants consume carbon dioxide as they grow, bioethanol has an overall lower carbon footprint than fossil fuels. [48] Substituting ethanol for oil can also reduce a country's dependence on oil imports. [49]

Advantages of cellulosic ethanol over corn or sugar-based ethanol Edit

U.S. Environmental Protection Agency
Draft life cycle GHG emissions reduction results
for different time horizon and discount rate approaches [50]
(includes indirect land use change effects)
Fuel Pathway 100 years +
2% discount
30 years +
0% discount
Corn ethanol (natural gas dry mill) (1) -16% +5%
Corn ethanol (Best case NG DM) (2) -39% -18%
Corn ethanol (coal dry mill) +13% +34%
Corn ethanol (biomass dry mill) -39% -18%
Corn ethanol (biomass dry mill with
combined heat and power)
-47% -26%
Brazilian sugarcane ethanol -44% -26%
Cellulosic ethanol from switchgrass -128% -124%
Cellulosic ethanol from corn stover -115% -116%
Notes: (1) Dry mill (DM) plants grind the entire kernel and generally produce
only one primary co-product: distillers grains with solubles (DGS).
(2) Best case plants produce wet distillers grains co-product.

Commercial production of cellulosic ethanol, which unlike corn and sugarcane would not compete with food production, would be highly attractive since it would alleviate pressure on these foodcrops.

Although its processing costs are higher, the price of cellulose biomass much cheaper than that of grains or fruits. Moreover, since cellulose is the main component of plants, the whole plant can be harvested, rather than just the fruit or seeds. This results in much better yields for instance, switchgrass yields twice as much ethanol per acre as corn. [51] Biomass materials for cellulose production require fewer inputs, such as fertilizer, herbicides, and their extensive roots improve soil quality, reduce erosion, and increase nutrient capture. [52] [53] The overall carbon footprint and global warming potential of cellulosic ethanol are considerably lower (see chart) [54] [55] [56] and the net energy output is several times higher than that of corn-based ethanol.

The potential raw material is also plentiful. Around 44% of household waste generated worldwide consists of food and greens. [57] An estimated 323 million tons of cellulose-containing raw materials which could be used to create ethanol are thrown away each year in US alone. This includes 36.8 million dry tons of urban wood wastes, 90.5 million dry tons of primary mill residues, 45 million dry tons of forest residues, and 150.7 million dry tons of corn stover and wheat straw. [58] Moreover, even land marginal for agriculture could be planted with cellulose-producing crops, such as switchgrass, resulting in enough production to substitute for all the current oil imports into the United States. [59]

Paper, cardboard, and packaging comprise around 17% of global household waste [57] although some of this is recycled. As these products contain cellulose, they are transformable into cellulosic ethanol, [58] which would avoid the production of methane, a potent greenhouse gas, during decomposition. [60]

General disadvantages Edit

The main overall drawback of ethanol fuel is its lower fuel economy compared to gasoline. [49]

Disadvantages of cellulosic ethanol over corn or sugar-based ethanol Edit

The main disadvantage of cellulosic ethanol is its high cost and complexity of production, which has been the main impediment to its commercialization (see below). [61] [62]

Although the global bioethanol market is sizable (around 110 billion liters in 2019), the vast majority is made from corn or sugarcane, not cellulose. [63] In 2007, the cost of producing ethanol from cellulosic sources was estimated ca. USD 2.65 per gallon (€0.58 per liter), which is around 2–3 times more expensive than ethanol made from corn. [64] However, the cellulosic ethanol market remains relatively small and reliant on government subsidies. [62] The US government originally set cellulosic ethanol targets gradually ramping up from 1 billion liters in 2011 to 60 billion liters in 2022. [65] However, these annual goals have almost always been waived after it became clear there was no chance of meeting them. [61] Most of the plants to produce cellulosic ethanol were canceled or abandoned in the early 2010s. [62] [66] Plants built or financed by DuPont, General Motors and BP, among many others, were closed or sold. [67] As of 2018, only one major plant remains in the US. [62]

In order for it to be grown on a large-scale production, cellulose biomass must compete with existing uses of agricultural land, mainly for the production of crop commodities. Of the United States' 2.26 billion acres (9.1 million km 2 ) of unsubmerged land, [68] 33% are forestland, 26% pastureland and grassland, and 20% crop land. A study by the U.S. Departments of Energy and Agriculture in 2005 suggested that 1.3 billion dry tons of biomass is theoretically available for ethanol use while maintaining an acceptable impact on forestry, agriculture. [69]

Comparison with corn-based ethanol Edit

Currently, cellulose is more difficult and more expensive to process into ethanol than corn or sugarcane. The US Department of Energy estimated in 2007 that it costs about $2.20 per gallon to produce cellulosic ethanol, which is 2–3 times much as ethanol from corn. Enzymes that destroy plant cell wall tissue cost USD 0.40 per gallon of ethanol compared to USD 0.03 for corn. [64] However, cellulosic biomass is cheaper to produce than corn, because it requires fewer inputs, such as energy, fertilizer, herbicide, and is accompanied by less soil erosion and improved soil fertility. Additionally, nonfermentable and unconverted solids left after making ethanol can be burned to provide the fuel needed to operate the conversion plant and produce electricity. Energy used to run corn-based ethanol plants is derived from coal and natural gas. The Institute for Local Self-Reliance estimates the cost of cellulosic ethanol from the first generation of commercial plants will be in the $1.90–$2.25 per gallon range, excluding incentives. This compares to the current cost of $1.20–$1.50 per gallon for ethanol from corn and the current retail price of over $4.00 per gallon for regular gasoline (which is subsidized and taxed). [70]

Enzyme-cost barrier Edit

Cellulases and hemicellulases used in the production of cellulosic ethanol are more expensive compared to their first generation counterparts. Enzymes required for maize grain ethanol production cost 2.64-5.28 US dollars per cubic meter of ethanol produced. Enzymes for cellulosic ethanol production are projected to cost 79.25 US dollars, meaning they are 20-40 times more expensive. [71] The cost differences are attributed to quantity required. The cellulase family of enzymes have a one to two order smaller magnitude of efficiency. Therefore, it requires 40 to 100 times more of the enzyme to be present in its production. For each ton of biomass it requires 15-25 kilograms of enzyme. [72] More recent estimates [73] are lower, suggesting 1 kg of enzyme per dry tonne of biomass feedstock. There is also relatively high capital costs associated with the long incubation times for the vessel that perform enzymatic hydrolysis. Altogether, enzymes comprise a significant portion of 20-40% for cellulosic ethanol production. A recent paper [73] estimates the range at 13-36% of cash costs, with a key factor being how the cellulase enzyme is produced. For cellulase produced offsite, enzyme production amounts to 36% of cash cost. For enzyme produced onsite in a separate plant, the fraction is 29% for integrated enzyme production, the faction is 13%. One of the key benefits of integrated production is that biomass instead of glucose is the enzyme growth medium. Biomass costs less, and it makes the resulting cellulosic ethanol a 100% second-generation biofuel, i.e., it uses no ‘food for fuel’. [ citation needed ]

In general there are two types of feedstocks: forest (woody) Biomass and agricultural biomass. In the US, about 1.4 billion dry tons of biomass can be sustainably produced annually. About 370 million tons or 30% are forest biomass. [74] Forest biomass has higher cellulose and lignin content and lower hemicellulose and ash content than agricultural biomass. Because of the difficulties and low ethanol yield in fermenting pretreatment hydrolysate, especially those with very high 5 carbon hemicellulose sugars such as xylose, forest biomass has significant advantages over agricultural biomass. Forest biomass also has high density which significantly reduces transportation cost. It can be harvested year around which eliminates long term storage. The close to zero ash content of forest biomass significantly reduces dead load in transportation and processing. To meet the needs for biodiversity, forest biomass will be an important biomass feedstock supply mix in the future biobased economy. However, forest biomass is much more recalcitrant than agricultural biomass. Recently, the USDA Forest Products Laboratory together with the University of Wisconsin–Madison developed efficient technologies [15] [75] that can overcome the strong recalcitrance of forest (woody) biomass including those of softwood species that have low xylan content. Short-rotation intensive culture or tree farming can offer an almost unlimited opportunity for forest biomass production. [76]

Woodchips from slashes and tree tops and saw dust from saw mills, and waste paper pulp are forest biomass feedstocks for cellulosic ethanol production. [77]

Switchgrass (Panicum virgatum) is a native tallgrass prairie grass. Known for its hardiness and rapid growth, this perennial grows during the warm months to heights of 2–6 feet. Switchgrass can be grown in most parts of the United States, including swamplands, plains, streams, and along the shores & interstate highways. It is self-seeding (no tractor for sowing, only for mowing), resistant to many diseases and pests, & can produce high yields with low applications of fertilizer and other chemicals. It is also tolerant to poor soils, flooding, & drought improves soil quality and prevents erosion due its type of root system. [78]

Switchgrass is an approved cover crop for land protected under the federal Conservation Reserve Program (CRP). CRP is a government program that pays producers a fee for not growing crops on land on which crops recently grew. This program reduces soil erosion, enhances water quality, and increases wildlife habitat. CRP land serves as a habitat for upland game, such as pheasants and ducks, and a number of insects. Switchgrass for biofuel production has been considered for use on Conservation Reserve Program (CRP) land, which could increase ecological sustainability and lower the cost of the CRP program. However, CRP rules would have to be modified to allow this economic use of the CRP land. [78]

Miscanthus × giganteus is another viable feedstock for cellulosic ethanol production. This species of grass is native to Asia and is a sterile hybrid of Miscanthus sinensis and Miscanthus sacchariflorus. It has high crop yields, is cheap to grow, and thrives in a variety of climates. However, because it is sterile, it also requires vegetative propagation, making it more expensive. [79]

It has been suggested that Kudzu may become a valuable source of biomass. [80]

Fueled by subsidies and grants, a boom in cellulosic ethanol research and pilot plants occurred in the early 2000s. Companies such as Iogen, POET, and Abengoa built refineries that can process biomass and turn it into ethanol, while companies such as DuPont, Diversa, Novozymes, and Dyadic invested in enzyme research. However, most of these plants were canceled or closed in the early 2010s as technical obstacles proved too difficult to overcome. As of 2018, only one cellulosic ethanol plant remained operational. [62]

In the later 2010s, various companies occasionally attempted smaller-scale efforts at commercializing cellulosic ethanol, although such ventures generally remain at experimental scales and often dependent on subsidies. The companies Granbio, Raízen and the Centro de Tecnologia Canavieira each run a pilot-scale facility operate in Brazil, which together produce around 30 million liters in 2019. [81] Iogen, which started as an enzyme maker in 1991 and re-oriented itself to focus primarily on cellulosic ethanol in 2013, owns many patents for cellulosic ethanol production [82] and provided the technology for the Raízen plant. [83] Other companies developing cellulosic ethanol technology as of 2021 are Inbicon (Denmark) companies operating or planning pilot production plants include New Energy Blue (US), [84] Sekab (Sweden) [85] and Clariant (in Romania). [86] Abengoa, a Spanish company with cellulosic ethanol assets, became insolvent in 2021. [87]

The Australian Renewable Energy Agency, along with state and local governments, partially funded a pilot plant in 2017 and 2020 in New South Wales as part of efforts to diversify the regional economy away from coal mining. [88]

US Government support Edit

From 2006, the US Federal government began promoting the development of ethanol from cellulosic feedstocks. In May 2008, Congress passed a new farm bill that contained funding for the commercialization of second-generation biofuels, including cellulosic ethanol. The Food, Conservation, and Energy Act of 2008 provided for grants covering up to 30% of the cost of developing and building demonstration-scale biorefineries for producing "advanced biofuels," which effectively included all fuels not produced from corn kernel starch. It also allowed for loan guarantees of up to $250 million for building commercial-scale biorefineries. [89]

In January 2011, the USDA approved $405 million in loan guarantees through the 2008 Farm Bill to support the commercialization of cellulosic ethanol at three facilities owned by Coskata, Enerkem and INEOS New Planet BioEnergy. The projects represent a combined 73 million US gallons (280,000 m 3 ) per year production capacity and will begin producing cellulosic ethanol in 2012. The USDA also released a list of advanced biofuel producers who will receive payments to expand the production of advanced biofuels. [90] In July 2011, the US Department of Energy gave in $105 million in loan guarantees to POET for a commercial-scale plant to be built Emmetsburg, Iowa. [91]

The Different Parts of the Urinary Tract, and Those More Prone to Infection

The urinary system is well designed and can often keep E. coli and other types of microscopic invaders at bay. For instance, urinating usually does an excellent job of flushing out lingering bacteria from the urethra before it causes any issues. (3) But when this defense fails, bacteria such as E. coli enters the urinary tract (which is made up of the kidneys, ureters, bladder , and urethra), multiplies, and then a urinary tract infection can develop. (5)

While any part of the urinary tract can be impacted, most E. coli–caused UTIs occur in the lower urinary tract, which includes the bladder (where urine is stored) and the urethra (the tube urine passes through to leave the body). A UTI that resides in the bladder is called cystitis one that resides in the urethra is called urethritis. (5)


Approaches Based on Algae

While nonphotosynthetic processes for carbon waste gas utilization can produce high biomass yields, these systems suffer from poor life-cycle analyses when compared to photosynthetic methodologies (Vieira et al., 2013). Among photosynthetic processes, algal biomass provides high yields, with more than 30- to 50-fold improvement in oil yield in comparison to common agricultural crops (Singh and Gu, 2010). The term algae broadly refers to any photosynthetic prokaryotic microorganism (cyanobacteria) or eukaryotic microorganism (microalgae) that can be cultivated. Algae cultivation was introduced in the 1950s and has been practiced both academically and commercially for decades (Fisher and Burlew, 1953 Golueke and Oswald, 1959). Algae can be viewed as self-replicating machines that convert sunlight and CO2 into value-added products. Products of algae cultivation include a wide range of biofuels, dietary protein and food additives, commodities, and specialized chemicals.

Algal biofuels have long been seen as a potentially viable strategy for addressing dwindling petroleum reserves, and investments in algal biofuel technologies have tended to rise when crude oil prices rise. For example, the U.S. Department of Energy (DOE) Office of

Fuels Development funded the Aquatic Species Program, focused on the production of biodiesel from microalgae, from 1978 to 1996 (Sheehan et al., 1998), and in 2008 DOE renewed its support for algal biofuel research after crude oil prices topped $100/barrel. This most recent spike in research investment coincided with large investments in algal biofuel technologies by startups and established companies however, a subsequent decrease in crude oil prices to below $30/barrel in 2016 created a major setback to industrial efforts. There are a wide range of estimates for the cost of production for algal biofuels (NRC, 2012) and it is difficult to predict how future policy and other factors may influence their competitiveness with traditional fossil resources however, one challenge of algal cultivation stems from the cost and development of infrastructure for physical processing, such as dewatering and extraction, that continue to pose challenges for energy demand (Sander and Murthy, 2010). One obstacle to the growth of these methods will be adoption of support and infrastructure commonly available to other forms of domestic agriculture (Trentacoste et al., 2015). Multiple reports describe the state of the science and research challenges for algal biofuels (DOE BETO, 2016, 2017 DOE EERE, 2010, 2017 NRC, 2012 Sheehan et al., 1998).

Algae cultivation offers significant productivity improvements as compared to contemporary agriculture. Multiple studies have demonstrated up to 30-fold productivity improvements in oil production (Abishek et al., 2014 Wen and Johnson, 2009) and 50-fold improvements in protein production from algae when compared to soybean, canola, or corn (Bleakley and Hayes, 2017) per acre of land. Although not fully evaluated, protein derived from algal biomass has been proposed for both animal and human consumption, and some companies, for example, Qualitas, 1 already produce specialized algae to enrich animal feeds. The competitive landscape for algal cultivation would shift should these commercial applications prove competitive with current agricultural products such as soy or corn.

Despite these advantages, there are also significant challenges to algae cultivation. Photosynthesis is inherently inefficient, as only 3-6 percent of total solar radiation energy is captured (Zelitch, 1975). In addition, a limitation of algae is the efficiency of the carbon dioxide conversion that limits the annual flux (Wilcox, 2012). To compensate for this low efficiency, cultivation ponds or bioreactors are often designed to maximize light exposure through large volume and surface area as a result, algal cultivation can be land and water intensive. Theoretically, to capture all CO2 from a 10 kiloton/day power plant would require 25-37 acres of cultivation (Hazelbeck, D., personal communication, 2018). In addition, biomass cultivation presents capital and operations costs that do not have close parallels in existing large-scale industries.

On the other hand, a key benefit to biological systems is their inherent flexibility in terms of feedstocks and environments. There are opportunities to utilize nonarable land and saline water or wastewater for algae cultivation, thereby minimizing competition for natural

resources. In addition, biological systems can tolerate low CO2 concentrations and impurities in the carbon sources common to industrial power generation. These variables have a considerable impact on capital and operations costs for biological conversion technologies, and it is important to account for resource use and environmental impact when comparing algae cultivation to conventional agriculture or other activities.

Approaches Based on Green Algae

Green algae, a common term for eukaryotic single cellular photosynthetic organisms deriving from several phyla, are a highly diverse group of algae representing many thousands of identified species. Multiple strains have been validated in biomass applications, and their selection is commonly determined by culture conditions. For instance, algae from cold environments have adapted to these environments through modification of primary metabolism and physiology (Morgan-Kiss et al., 2006).

The majority of biomass cultivation efforts using green algae have focused on biofuel production. In addition, several co-products have been identified that can help make biofuel production more economically feasible. The following sections describe opportunities for using green algae to produce various biofuels and co-products, as well as opportunities and limitations involved in advancing such applications through genetic manipulation.

Biodiesel Production

Biodiesel, or fatty acid methyl ester, production from plant- and animal-based fats and oils (including those from green algae) is now considered part of a mature industry. While biodiesel can be used as drop-in diesel replacement in many diesel engines, its hygroscopicity, cloud point, and fouling properties have limited widespread adoption. Once considered an ultimate product of biomass production, biodiesel has more recently become relegated to small fuel markets and as an additive for ultra-low-sulfur petroleum diesel to provide added lubricity properties (Hazrat et al., 2015).

After biomass harvesting, lipids are extracted by one of several methods to prepare isolated neutral lipids, or triacylglyceride (TAG), or total lipids containing both polar and neutral lipids. The traditional process for biodiesel production from plant- and animal-based fats and oils typically requires a high-purity source of TAG however, methods have been developed to produce biodiesels from total lipid extracts (Asikainen et al., 2015 Mubarak et al., 2015).

Renewable Diesel/Gasoline Production

The most broadly adoptable advances in algae biomass conversion to biofuels lies in the catalytic hydrogenation of lipid extracts, known as hydrotreatment, for the production of true diesel and gasoline components. Although the methods for these transformations have been

known for decades, the capital infrastructure required to carry out hydrotreatment on a commercial scale has limited its implementation to a few refining companies, such as Neste Oil and Chevron (Al-Sabawi and Chen, 2012 No, 2014).

One particular benefit of renewable diesel and gasolines is the flexibility with regard to the source of feedstock. Currently, industrial hydrotreatment processing can utilize fats and oils from any bioderived triacylglyceride source, including vegetable oils and animal fats. However, while algal lipids do contain triacylglycerides, extracts from many species also contain significant portions of polar lipids and hydrophobic pigments. New technology may be required for conversion of these species into renewable fuels by hydrotreatment facilities (Davis et al., 2013). The requirement of hydrogen gas for the catalytic hydrogenation of lipid extracts must also be considered.

In addition, liquefaction, pyrolysis, and gasification offer routes to produce a variety of fuels, including methane, ethanol, and fuel oils (Demirbas, 2010). Many of these have been proposed to be routed to more complex chemicals through reforming processes (Bhujade et al., 2017).

Valorization of Co-Products

In order to compete with fossil fuels, biofuels must provide attractive economics. This can be achieved by increasing the cost of petroleum products through pricing nonrenewable carbon (or other mechanisms), or the value of algae cultivation can be further increased by production of additional products. Biomass conversion to biofuels inherently creates waste products from unused biomass, primarily composed of protein and carbohydrates. These can be consumed by anaerobic digestion to produce biogas, a low-value end product. Or, they can be converted into a variety of more valuable products. Combined approaches that utilize photosynthetic, fermentative, and chemical methods can be employed to produce products with high value. For instance, a combined algal processing method capable of producing multiple products from algae biomass was recently demonstrated (Miara et al., 2014). In such systems, algal carbohydrates can be diverted to fermentative production of ethanol or other commodity chemicals, such as succinic acid (Raab et al., 2010) from metabolically engineered yeast, while the proteins may be applied to adhesive manufacturing (Roy et al., 2014). Examples of co-products that may be valorized to improve the economics of biofuel production from algae include dietary protein, polyunsaturated fatty acids, and pigments.

Dietary protein. Algae have long been viewed as an attractive source of dietary protein, both as animal feeds and for human consumption (Becker, 2007). Green algae contain between 40 and 70 percent of their dry weight as protein, and the amino acid profile of most algae compares favorably with common food proteins. A few studies have been performed to determine algae&rsquos protein efficiency ratio (PER), which reflects an animal&rsquos weight gain per unit

of protein consumed one study found algae offered up to 80 percent PER compared to milk proteins (Becker, 2004) while incurring a substantially smaller footprint than milk production in terms of freshwater resources and arable land (Bleakley and Hayes, 2017). Protein productivity from algae has been estimated at up to 50 times that of soybeans per acre of land. This suggests green algae has the potential to supplement or replace crops for animal feed with vastly reduced demands for arable land, but studies have not been validated at scale.

Polyunsaturated fatty acids. Many green algae naturally produce polyunsaturated fatty acids (PUFAs) that are valuable for humans and animals as food additives. While they are typically harvested from fish, PUFAs from algae represent a viable and more sustainable set of target molecules (Adarme-Vega et al., 2012). In particular the omega-3 varieties eicosapentaenoic acid and docosahexaenoic acid are valuable for the supplementation of many farmed animals, particularly carnivorous fish such as salmon and tuna (Ahmed et al., 2012 Bimbo, 2007). Omega-3 feeding studies have also been performed on cattle and poultry (Nitsan et al., 1999 Ponnampalam et al., 2006). In addition, omega-3 fatty acids demonstrate a consumer market as neutraceutical products that is currently dominated by fish oils.

Pigments. Colorants are manufactured as additives to foods, drinks, cosmetics, and a host of other products in order to increase the appeal of the product to the consumer. Some pigments are manufactured utilizing fossil fuel sources such as coal. Algae are well known for their ability to produce a variety of pigments and could provide a more sustainable alternative to the utilization of fossil fuels (Kaur et al., 2009). One well-known example of the successful application of algae for the commercial production of pigments is astaxanthin (see Box 5-1). The manufacture of other pigments with algae remains a potential opportunity for further exploration.

Genetic Manipulation of Green Algae

The tools to facilitate genetic manipulation of green algae are far less advanced than tools for genetic manipulation of bacteria, yeast, and vascular plants. Key challenges include poor genome insertion, gene silencing, and unoptimized promoter systems. Nevertheless, some academic and industrial groups have attempted genetic modification of green algae for the purposes of improving photosynthetic efficiency, decreasing photodamage of the light harvesting complex, and optimizing the efficiency of carbon uptake and incorporation. Several products have also been sought from transgenic expression, including high-value therapeutic proteins, biohydrogen, lipids, and terpenoids (Gimpel, 2013).

Many species of green algae are genetically tractable, but Chlamydomonas reinhardtii has become the primary model system, with the majority of tools and techniques developed for this species (Rasala et al., 2013). Green algae have three separate genomes: nuclear,

Box 5-1Astaxanthin

Astaxanthin is a carotenoid pigment produced by a variety of microalgae. A polyunsaturated tetraterpenoid, astaxanthin presents a red coloration and is naturally responsible for pink or red pigmentation in fish, crustaceans, and shellfish through their diet. As a result, astaxanthin has become an important component in aquaculture feed to provide red/pink pigmentation to a variety of farmed seafood, particularly shrimp and salmon, with a market value approaching $250 million in 2010 (Borowitzka, 2013). It is also marketed as a nutraceutical, and a variety of vitamin supplements contain astaxanthin for its antioxidant properties. Cyanotech (Kona, Hawaii) produces astaxanthin from Haematococcus pluvialis in 25 hectares of open ponds, while Algatechnologies Ltd. (Ketura, Israel) uses photobioreactors with the same organism. BASF (Hutt Lagoon, Australia) produces the pigment from Dunaliella salina in 740 hectares of open ponds (Maeda et al., 2018). While commercial operations have shown profitability using open-air CO2 absorption, CO2 fixation by these commercial strains could benefit from increasing CO2 concentration up to a 5 percent level can provide significant improvements in biomass and astaxanthin levels (Chekanov et al., 2017).

chloroplast, and mitochondrial. In C. reinhardtii, each of these genomes has been sequenced, and tools and protocols for genetic manipulation have been developed for each. Commonly, nuclear and chloroplast genomes have been primarily targeted for transgene expression, and each location involves special tools, advantages, and challenges. Engineering the nuclear genome for protein production offers benefits similar to other eukaryotic expression systems, including regulated expression, cytoplasmic protein folding machinery, access to posttranslational modification (such as glycosylation), and protein secretion. Furthermore, like many green algae, C. reinhardtii is a haploid organism. As a result, nuclear genome modifications can be used in combination with sexual breeding to provide additional genetic control. However, nuclear transformation currently suffers from several drawbacks, including random gene insertion, low protein expression, and gene silencing (Cerutti et al., 1997 De Wilde et al., 2000 Wu-Scharf, 2000). These complications are being actively researched, and some limited solutions have been recently developed. For instance, gene silencing has been addressed by directly coupling transgene expression with antibiotic resistance gene expression, therefore imparting a strict selection method for gene retention (Rasala, 2012, 2013). In addition, experiments employing new technologies for gene integration and knockout, including CRISPR methodologies, are currently under way (Ferenczi et al., 2017 Shin et al., 2016). Nevertheless, nuclear expression remains limited in terms of gene size, number, and complexity.

Tools and methodologies for manipulating gene expression in the chloroplast have achieved significant advances in recent years. Unlike nuclear expression, chloroplast expression allows homologous recombination, and multiple vectors, selection, and transformation methods have been developed. The green algae chloroplast offers an environment for protein expression, including chaperones and posttranslational modification enzymes that can facilitate proper protein and expression folding, and heterologous protein expression has achieved remarkable levels, sometimes as high as 10 percent of total soluble protein (Mayfield, 2007). Despite these benefits, chloroplast expression remains limited in terms of gene size and number, challenging the needs for engineering large operons and metabolic pathway design.

In the broader scheme of genetic engineering tools, photosynthetic organisms have historically received a low level of research support, and algae represent a small subset of this group (Gao et al., 2012). However, tools developed in recent decades underscore the potential for green algae as an ideal photosynthetic host organism. New and improved genetic tools for green algae genetic manipulation remain a key need for the advancement of algae biomass applications. This includes development of genetic insertion (homologous recombination) technologies, identification of robust promoters for gene expression, development of synthetic operons for multiple gene incorporation, tools for engineering large genes and pathways, and novel selection methods.

Approaches Based on Cyanobacteria

Cyanobacteria are being engineered to directly convert solar energy, carbon dioxide, and water to biofuels and other products. Cyanobacteria-based approaches possess advantages over traditional biological production systems based on plants, green algae, or heterotrophic organisms. For example, cyanobacteria are more amenable to genetic manipulation than are algae and can therefore be adapted for the production of a wider range of products (see Figure 5-1). The photosynthetic efficiency of cyanobacteria is two to four times higher than that of plants (Melis, 2009), and their cultivation does not compete with food crops for land usage. Cyanobacteria also do not have the sugar requirements of heterotrophic organisms such as Escherichia coli and yeast, although they have a reduced growth rate and fewer available synthetic biology tools compared to these hosts. While heterotrophic hosts may be adapted to utilize cellulosic feedstocks these technologies are still not well developed and as such efficiency can be low. Cyanobacteria allow us to skip this carbon-harvesting step and fix CO2.

There is a plethora of cyanobacterial species but only a few have been adapted to chemical production. The three predominant strains utilized for chemical production are Synechococcus elongatus PCC 7942 (7942), Synechocystis sp. PCC 6803 (6803), and Synechococcus sp. PCC 7002 (7002). These strains all have sequenced genomes, established culturing methods, and basic metabolic engineering tools (Berla et al., 2013 Markley et al., 2015 Nozzi et al., 2017 Yu et al., 2015), yet each strain presents its own unique advantages and challenges.

FIGURE 5-1 Example pathways for products from cyanobacteria. In the figure 23BD = 2,3-butanediol FPP = farnesyl pyrophosphate GPP = gross primary production DMAPP = dimethylallyl diphosphate IPP = isopentenyl diphosphate 3HB CoA = 3-hydroxybutyryl-CoA 4HB CoA = 4-hydroxybutyryl-CoA FAEEs = fatty acid ethyl esters 3HP = 3-hydroxypropionic acid Poly 3HB = poly(3-hydroxybutyrate) and Poly (3-HB-Co-4-HB) = poly(3-hydroxybutyrate-co-4-hydroxybutyrate). 1
1 Reprinted from Carroll, Austin L., Anna E. Case, Angela Zhang, and Shota Atsumi. 2018. &ldquoMetabolic engineering tools in model cyanobacteria.&rdquo Metabolic Engineering. doi:10.1016/j.ymben.2018.03.014 with permission from Elsevier.

7942 is a freshwater unicellular cyanobacterium and studied for photosynthesis and genetic manipulation (Golden et al., 1987). 7002 is a marine species capable of growth in a variety of conditions (salt, temperature, and light) (Batterton and Van Baalen, 1971), making it more apt to utilize natural saltwater resources. However, metabolic engineering has not been as well studied in 7002 as in freshwater species and synthetic biology capabilities for 7002 lag behind that of 7942 and 6803. 6803 has a good selection of genetic tools and can grow photomixotrophically.

Cyanobacteria have been engineered to produce a wide range of fuels, fuel precursors, and commodity chemicals. The productivities and titers (mostly on the order of milligrams per liter) are generally too low to make commercialization of the technology appealing, however, and these technologies have been demonstrated primarily on small, academic research scales. To move these technologies toward commercialization, it will be critical to reduce the cost of production and further advance tools for genetic manipulation and metabolic engineering of cyanobacteria.

Fuel Production

Fuels and fuel precursors that can be produced with cyanobacteria include ethanol, butanol, fatty acids, heptadecane, limonene, and bisabolene.

Ethanol. Ethanol, a common biofuel candidate, has been produced in cyanobacteria by the transfer of two ethanol pathway genes from yeast. This production has been enhanced to form 5.5 g/L ethanol in Synechocystis sp. PCC 6803 (Dexter and Fu, 2009 Gao et al., 2012).

Butanol (n-butanol and isobutanol). Isobutanol has applications as a drop-in biofuel that could be integrated into current energy infrastructure. Production can be achieved by expressing heterologous 2-ketoisovalerate decarboxylase and aldehyde reductase that redirect carbon flux from the L-valine biosynthetic pathway to isobutanol resulting in a titer of 0.5 g/L in 7942 (Atsumi et al., 2009). n-butanol, which also has uses as a drop-in biofuel and is currently used in a variety of consumer products, can be produced in 7942 (Lan and Liao, 2011). The n-butanol biosynthetic pathway was constructed based on the natural n-butanol biosynthetic pathway found in Clostridium acetobutylicum. Examination and characterization of Coenzyme A-acylating (CoA) aldehyde dehydrogenases that are oxygen tolerant enabled production of butyraldehyde from butyryl-CoA. Final titers of 404 mg/L were achieved with productivity of 2 mg/L/h (Lan et al., 2013).

Fatty acids. Fatty acids can be used in the synthesis of biodiesels. Cyanobacteria natively produce free fatty acids in trace amounts. This production has been improved in 7942, 6803, and 7002. Free fatty acid overproduction leads to reduced cell fitness. Free fatty acid production in 7002 results in reduced negative impacts on the cell, and production can be further improved by the nonnative expression of RuBisCO leading to >130 mg/L free fatty acids being produced (Ruffing, 2014). When coupled with native free fatty acid production, the expression of a heterologous diacylglycerol acyltransferase in tandem with an ethanol production pathway results in a variety of fatty acid ethyl esters (FAEEs) in 7942. This platform leads to the accumulation of up to 7 mg/L of FAEEs (Lee et al., 2017a).

Heptadecane. Long-chain hydrocarbons such as heptadecane (see Table 5-1) can be readily used in fuel production for the synthesis of biodiesel. Heptadecane production can be achieved by capitalizing on the natural fatty acid production in the cell. Titers can be improved in strains that natively produce alkanes by overexpressing an acyl-ACP reductase/aldehyde-deformylating oxygenase (AAR/ADO) pathway. Expression of the AAR/ADO pathway from 7942 in a marine cyanobacterium results in up to 4.2 µg/g dry cell weight heptadecane (Yoshino et al., 2015).

TABLE 5-1 Summary of commodity chemicals and fuels that are currently synthesized from CO2 from cyanobacteria and their reported production.

Compound Strain Reported Production
Ethanol Synechocystis sp. PCC 6803 5.5 g/L
Isobutanol Synechococcus elongatus PCC 7942 0.5 g/L
n-Butanol Clostridium acetobutylicum 2 mg/L/h
Fatty acids Synechococcus elongatus PCC 7942
Synechocystis sp. PCC 6803
Synechococcus sp. PCC 7002
>130 mg/L
Synechococcus elongatus PCC 7942 4.2 µg/g dry cell weight
Synechococcus elongatus PCC 7942
Synechococcus sp. PCC 7002
Synechocystis sp. PCC 6803
1 mg/L/OD730/day
4 mg/L
7 mg/L
Synechococcus sp. PCC 7002 0.6 mg/L
Synechococcus elongatus PCC 7942 2.4 g/L (after 21 days)
3 g/L (after 10 days, photomixotrophic conditions)
12.6 g/L (continuous lighting with glucose and CO2)
5.7 g/L (23-hour light cycling)
Synechococcus elongatus PCC 7942 0.3 g/L

Compound Strain Reported Production
Ethylene Synechocystis sp. PCC 6803 2.5 mL/h/OD730
Glycogen Synechococcus sp. PCC 7002 3.5 g/L (after 7 days)
Lactate Synechocystis sp. PCC 6803
Synechococcus elongatus PCC 7942
0.8 g/L (after 2 weeks)
1.4 g/L (after 10 days)
Synechococcus elongatus PCC 7942
Synechocystis sp. PCC 6803
0.8 g/L (after 6 days)
Isoprene Synechococcus elongatus PCC 7942 1.3 g/L (after 21 days)
Synechococcus elongatus PCC 7942 50 mg/L
Synechococcus elongatus PCC 7942 5 mg/L

Limonene. Limonene has applications as both a biofuel and a solvent. Limonene can be produced with the expression of a single heterologous gene for limonene synthase (LS) selected from spearmint for its high selectivity for limonene production. The expression of LS capitalizes on carbon flow through the native methyl-D-erythritol 4-phosphate (MEP) pathway, commonly found in plants, and funnel flow to limonene (Wang et al., 2016a). Establishing high LS expression is critical in improving titers. Limonene production has been established in 7942 and 7002 at 1 mg/L/OD730/day (Wang et al., 2016a) and 4 mg/L (Davies et al., 2014), respectively. In 6803, carbon flux through the oxidative pentose phosphate (OPP) pathway is used to drive limonene production. Overexpression of two native enzymes

(ribose-5-phosphate isomerase and ribulose 5-phosphate 3-epimerase) and expression of a heterologous geranyl diphosphate synthase gene establishes limonene production at 7 mg/L (Lin et al., 2017).

Bisabolene. Bisabolene has applications as a biodiesel candidate and production relies on carbon flow through the MEP pathway and the expression of a single heterologous gene, (E)-&alpha-bisabolene synthase. Production has been established in 7002 resulting in 0.6 mg/L (Davies et al., 2014).

Commodity Chemicals Production

Commodity chemicals that can be produced with cyanobacteria include 2,3-butanediol, 1,3-propanediol, ethylene, glycogen, lactate, 3-hydroxypropanoic acid, 3-hydroxybutanoic acid, 4-hydroxybutanoic acid, isoprene, and farnesene.

2,3-Butanediol. 2,3-Butanediol is a commodity chemical used to make synthetic polymers (van Haveren et al., 2008) and can readily be converted to methyl ethyl ketone, a fuel additive and solvent (Tran and Chambers, 1987). 2,3-Butanediol relies on carbon syphoned from central metabolism in the form of pyruvate. The condensation of two pyruvate molecules, followed by decarboxylation and reduction through heterologously expressed genes, results in 2.4 g/L 2,3-butanediol after 21 days of production (Oliver et al., 2013). 2,3-Butanediol was similarly produced in 6803 (Savakis et al., 2013) and 7002 (Nozzi et al., 2017). In 7942 the addition of glucose and expression of galP, encoding a hexose symporter, allows growth using both CO2 and glucose (McEwen et al., 2013). This modification permits continued 2,3-butanediol synthesis in both light and dark conditions, a potentially useful strategy for 24-hour metabolite production. Under photomixotrophic conditions 3 g/L 2,3-butanediol can be produced over 10 days (McEwen et al., 2016). Global metabolic rewiring of these strains for improved CO2 fixation increases 2,3-butanediol titers, glucose utilization efficiency, and carbon fixation in both lighted and diurnal conditions (Kanno et al., 2017). Under continuous lighting, rewired 7942 produces 12.6 g/L 2,3-butanediol (Kanno et al., 2017) when fed glucose and CO2. Under natural 24-hour (diurnal) light cycling in the rewired strain, 5.7 g/L 2,3-butanediol is produced.

1,3-Propanediol. 1,3-Propanediol has a variety of uses including in polymers, paints, solvents, and antifreeze. Production in cyanobacteria is dependent on removal of carbon from the Calvin-Benson cycle in the form of dihydoxyacetone phosphate (DHAP). DHAP can be converted to 1,3-propanediol via a four-step heterologously expressed metabolic pathway. After the disruption of production bottlenecks 0.3 g/L 1,3-propanediol was produced in 7942 (Hirokawa et al., 2016).

Ethylene. Ethylene can be used to generate polyethylene, a widely used polymer, and in its gaseous state can be used to speed the ripening of produce. Production has been established in 6803 and relies on the expression of a single heterologous gene, efe, encoding ethylene forming enzyme (Efe). Efe converts 2-oxoglutarate from the tricarboxylic acid (TCA) cycle to ethylene. Partial deletion of ntcA, encoding the global transcription factor known for regulating carbon metabolism (Mo et al., 2017), exhibits a 23 percent increase in ethylene production and 1.5-fold increase in Efe activity in 6803 (Mo et al., 2017). Ethylene production in this strain reached 2.5 mL/h/OD730 (Mo et al., 2017).

Glycogen. Glycogen is a common form of energy storage and a potential carbon source for chemical production. It can also be converted into ethanol for use as a biofuel. 7002 naturally amasses glycogen under nitrogen-depleted conditions. Glycogen production can be improved by modifying culture conditions, including light intensity, CO2 concentration, and salinity (Aikawa et al., 2014). Improved production conditions result in 3.5 g/L glycogen after 7 days (Aikawa et al., 2014).

Lactate. Biological synthesis of lactic acid for biodegradable polymers has been established in 6803. Lactic acid production requires the heterologous expression of a single enzyme, lactate dehydrogenase, which pulls pyruvate from central metabolism to catalyze the conversion to lactic acid at a titer of 0.8 g/L after 2 weeks of cultivation (Angermayr et al., 2014). Further optimization of co-factor requirements leads to approximately 1.4 g/L lactate produced after 10 days in 7942 (Li et al., 2015).

3-Hydroxypropionic acid, 3-hydroxybuterate, and 4-hydroxybuterate. 3-Hydroxypropionic acid (3-HP), 3-hydroxybuterate (3-HB), and 4-hydroxybuterate (4-HB) (Table 5-1) all have applications for the synthesis of polymers and plastics used in daily life. 3-HB was produced in 6803 and 7942 by the expression of a nonnative malonyl-CoA reductase, which converts malonyl-CoA to 3-HP. Malonyl-CoA natively feeds into fatty acid biosynthesis. This production can be aided by streamlining carbon flux to malonyl-CoA, expressing malonyl-CoA reductase, and optimizing nicotinamide adenine dinucleotide phophaste (NADPH) levels, resulting in 0.8 g/L after 6 days (Wang et al., 2016b). 7002 has been demonstrated to produce poly-3-hydroxybutyrate and poly-3-hydroxybutyrate-co-4-hydroxybutyrate through the introduction of a gene cluster from Chlorogloea fritschii PCC 9212 (Zhang et al., 2015). 3-HB production relies on native acetyl-CoA production, which can be converted to 3-HB in two consecutive steps. Alternatively, 4-HB can be produced by pulling carbon from the TCA cycle in the form of succinic semialdehyde. 3-HB can be produced by itself or in combination with 4-HB to create a copolymer. Production can reach 0.05 g/L of 3-HB or 4.5 percent total cell dry weight of the copolymer, with 4-HB accounting for 12 percent of the copolymer (Zhang et al., 2015).

Isoprene. Isoprene is commonly used for the production of synthetic rubbers. Overexpression of the native MEP pathway and the expression of plant isoprene synthase gene (ispS) in 7942 results in the production of isoprene. With this production platform 1.3 g/L isoprene was produced after 21 days (Gao et al., 2016).

Squalene. Squalene is widely used in the food, personal care, and medical industries but its commercial production is unreliable and nonideal. Squalene production has been previously established in heterotrophic hosts, such as yeast. Similarly, squalene was produced in 7942 by the overexpression of the native MEP pathway in addition to the expression of a heterologous squalene synthase from yeast (Choi et al., 2017). Following optimization in carbon flux, the expression of this pathway results in 50 mg/L squalene.

Farnesene. Farnesene has been used as a precursor for high-performance polymers and as a jet fuel candidate. Production in 7942 was achieved by the heterologous expression of a single gene encoding farnesyl synthase, which funnels carbon out of the MEP pathway in the form of farnesyl diphosphate. The engineered strain produced 5 mg/L of &alpha-farnesene (Lee et al., 2017b).

Genetic Manipulation of Cyanobacteria

While some target fuels and chemicals can be produced through exploitation of naturally occurring cyanobacteria traits, manipulation of organisms&rsquo genetic material can enable production of many other target molecules. An understanding of areas where desired genetic material may be inserted without interrupting essential functions (gene integration and plasmids), ways to control production of the desired chemical target (promoters and riboswitches), and ways to tell the cell where to start and stop generating the target molecule (ribosomal binding sites) are necessary in order to conduct genetic manipulations. CRISPR is another tool for reliable insertion of desired genetic material into host cells.

In addition to an enhanced ability to introduce and control novel pathways in cyanobacteria, a thorough understanding of the flow of carbon through the cell&rsquos metabolism is key to creating an industrially relevant production system. Techniques for improving carbon flux have included the use of carbon sinks, disrupting side pathways, removing inhibitors, and protein fusions to improve rate. While these techniques have pushed cyanobacteria toward industrial relevance there is still much that is unknown and needs further study.

Metabolic Engineering of Cyanobacteria

One of the primary challenges of chemical production in cyanobacteria is the low titers that result from most pathways. Production is usually limited to the milligram per liter (mg/L) order. A great deal of effort has gone into engineering RuBisCO, the primary carbon

capture mechanism in cyanobacteria. While some improvements have been made (Whitney et al., 2011), RuBisCO has proven very difficult to improve, and alterations still remain below the required threshold for wide-scale chemical production.

An alternative to RuBisCO engineering is the supplementation of cyanobacteria with alternate carbon sources including glucose, glycogen, acetate, and xylose. While these carbon sources effectively improve titers, this additional carbon is not always directed toward chemical production and is often lost as biomass. Careful rewiring of carbon integration into metabolism can help improve this carbon utilization and act as a carbon sink to improve CO2 fixation (Oliver and Atsumi, 2015 van der Woude et al., 2014). However, this approach requires a detailed understanding of carbon metabolism in the cell and how an alternate carbon source is being incorporated. In addition, these alternate carbon sources can increase the risk of contamination. As a slow-growing organism, cyanobacteria in culture can quickly become overcome by competing organisms that can utilize sources such as glucose. The necessity of using stricter techniques to prevent contamination, combined with the added cost of the alternate carbon source, can significantly raise production costs and eliminate some of the advantages of using cyanobacteria as a host strain.

In order to optimize carbon flow, researchers must fully understand it. Genome-scale models (GSMs) are important tools for assessing and engineering metabolic systems. The models may be used to describe an organism&rsquos entire metabolism utilizing genomic information (Broddrick et al., 2016 Kim et al., 2017 Shirai et al., 2016 Triana et al., 2014). It would be challenging to optimize production of a target chemical, identify bottlenecks, and target the best production system for the host without GSM. GSM-directed engineering has been successfully used to improve a variety of production platforms in E. coli, including 1,4-butanediol (Yim et al., 2011), lycopene (Alper et al., 2005), lactic acid (Fong et al., 2005), and succinate (Lee et al., 2005).

The transfer of GSMs from heterotrophs to photoautotrophs is a difficult process one reason is that photosynthetic growth complicates simple factors, like energy input, making them complex and difficult to measure. The recent development of two GSMs (Broddrick et al., 2016 Triana et al., 2014) for 7942 allows for greater predictive power when making modifications to metabolism. New insights have been gained using a GSM developed for 7002 (Vu et al., 2013), which could help engineer the strain (Hendry et al., 2016). Based on this model it has been predicted that up to 10 percent of fixed carbon could be rerouted to production (Vu et al., 2013). These predictions have not been confirmed experimentally.

Construction of a comprehensive GSM for 6803 has attempted to resolve many of the problems that are seen in 7942 and 7002 with GSM development in cyanobacteria (Hucka et al., 2003 Knoop et al., 2010 Stanford et al., 2015). However, automated assembly of key GSM components is prevented by incongruent gene annotations and database nomenclature. GSM analysis uses 3,167 genes to study gene function, carbon metabolism, photosynthesis, and chemical production (Shirai et al., 2016 Yoshikawa et al., 2017). These models can iden-

tify deletions of native metabolic reactions that compete for reductive power required in targeted chemical production successfully (Yoshikawa et al., 2017). While traditional GSMs are limited to the native genes found in 6803, recent iterations incorporate nonnative metabolic reactions to construct hybrid phototrophic and heterotrophic cyanobacteria models (Saha et al., 2016 Shirai et al., 2016). These expanded models can predict new strategies for metabolic pathway construction and increase yields in targeted chemical production.

Synthetic biology tools are essential to utilize a strain as a production host. These tools can range from sites for the expression of nonnative genes to systems for controlling gene expression or tools for genetic manipulation (Albers et al., 2015 Immethun et al., 2017 Ueno et al., 2017). Heterotrophic hosts such as yeast and E. coli have well-developed synthetic biology toolboxes however, these tools are often incompatible with photoautotrophic hosts such as cyanobacteria. In fact, tools developed for cyanobacteria are often difficult to transfer between strains, necessitating the development of a unique set of tools for gene integration and controlling gene expression on a transcriptional and translational level for each strain. Many of these tools have been developed for cyanobacteria (Berla et al., 2013 Golden et al., 1987 Markley et al., 2015). In addition to the difficulty of transferring tools between hosts, it can also be difficult to transfer pathways between host organisms. Gene expression, intermediates, and final products have varying toxicity across strains. Each strain can have a unique codon preference, altered enzyme activities, and constitutive promoters. One of the greatest challenges to metabolic engineering efforts for cyanobacteria is the careful tailoring of tools to each strain.

Key Considerations for Photosynthetic Approaches

Whether they involve the cultivation of green algae or cyanobacteria, photosynthetic approaches to carbon dioxide utilization raise a unique set of considerations relevant to their costs, benefits, environmental impacts, and social acceptability. Key considerations include the impact of the cultivation method used, restrictions in the use of genetically modified organisms, the degree of CO2 solvation, nutrient requirements and downstream burdens, impacts on water and land use, and availability and suitability of CO2 waste streams.

Impact of Cultivation Method

A key consideration for algae biomass production is the selection of cultivation method. Both open ponds and closed photobioreactor (PBR) designs have been implemented on academic and industrial scales (Figure 5-2). Open pond designs commonly employ oblong, &ldquoraceway&rdquo designs (Benemann et al., 1978) in which a high rate of water movement can be achieved with paddlewheels or air lift pumps. Raceway ponds have proven to be the dominant design, and their design has changed little in three decades, as their construction is believed to

FIGURE 5-2 Examples of algal cultivation technologies: (a) 1 and (b) 2 are different types of closed photobioreactors and (c) 3 is an example of an open raceway pond.
1 Photo by IGV Biotech.
2 Huang et al., 2017.
3 See (accessed October 10, 2018).

be among the least expensive of reactor designs. Raceway ponds can vary in size from several square feet to a hectare or more, and typically employ polymer bed liners and paddlewheel-style impellers. Bed liners are a requirement in some jurisdictions to meet water treatment codes, and the use of any genetically modified species may also mandate such barriers. The cost of bed liners commonly dominates the capital costs of a raceway pond, particularly with increasing size (Rogers et al., 2014).

Closed PBRs can involve a variety of scales, materials, and designs (Gupta et al., 2015), many of which serve to optimize solar radiation upon the biomass culture. The simplest and most common PBR implements a &ldquohanging bag&rdquo approach, where polyethylene tubes are suspended vertically with air and CO2 diffusers to provide agitation and carbon addition (Martínez-Jerónimo and Espinosa-Chávez, 1994). Multiple larger system designs have been introduced in recent years using a variety of materials. Inherent in the viability of any PBR system is temperature control, a common challenge of closed systems, and the ability to agitate the culture, typically with pumped liquids.

The capital and operational costs of PBRs are considerably higher than those for open ponds, and the adoption of each technology is dependent on multiple factors (Richardson et al., 2012). Closed PBRs have the benefit of tight control over cultivation conditions, including temperature, CO2 and O2 levels, sun exposure, contaminants, and evaporation. As a result, culture density can reach levels not obtainable in open ponds (Schoepp et al., 2014). The ability to exclude contaminants and exogenous species, including predators, offers additional benefits. As a result, closed PBRs have been favored for the production of high-value products, where product purity is favored over low-cost cultivation. Similarly, open ponds have been favored for the production of biofuels given the need to minimize production costs.

Use of Genetically Modified Organisms

Current U.S. law and regulations tightly constrain the use of genetically modified algae in outdoor open ponds due to concerns regarding containment and spread of genetically modified organisms (GMOs). A limited study was recently performed to evaluate the dispersal, colonization, and impact of GMO algae into native water bodies (Szyjka et al., 2017). The particular study found that while broad dispersal occurs, impact on indigenous species was viewed as negligible. Additional work in this area is needed.

Degree of CO2 Solvation

A major challenge to waste CO2 utilization in all aquatic systems is CO2 solvation. Simple sparging of gases in aqueous solutions results in costly and inefficient solubilization. Two methods have been developed: one utilizes an amine-based CO2 concentrator, followed by thermal stripping, and the second uses carbonate salts to deprotonate solvated CO2 into soluble bicarbonate. The latter method has also been shown to be improved with the use of exogenous

carbonic anhydrase, an enzyme that catalyzes CO2 solvation (Hernandez-Mireles et al., 2014). Each of these methods contains drawbacks, such as thermal input or high pH values. While some strains of algae can adapt to these conditions, more research is required to improve their integration into large-scale biomass cultivation (Könst et al., 2017). Some algal strains produce extracellular carbonic anhydrase to facilitate CO2 solvation (Huertas et al., 2000).

Nutrient Requirements and Downstream Burdens

As with all photosynthetic crops, cultivating algae requires the application of fertilizers such as phosphorus and nitrogen. This has raised concerns that algae cultivation could contribute to eutrophication (excessive algal growth due to the influx of nutrients) in freshwater and coastal zones. Eutrophication is already a serious environmental concern in many areas due to agricultural runoff and municipal wastewater. To prevent algae cultivation from further compounding this problem and even partially address eutrophication, some have advocated the use of municipal and agricultural wastewater as sources of nutrients for algal cultivation (Woertz et al., 2009), thus capturing these nutrients with biomass cultivation before they reach downstream water bodies (Abdel-Raouf et al., 2012 Benemann et al., 2003 Brune et al., 2003). These scenarios would require co-localization of algae cultivation and wastewater treatment, and model pilot studies show significant promise (Bohutskyi et al., 2016 Chekroun et al., 2014). Nutrient recycling is also an important area of research for algal cultivation both as a means to minimize waste and as part of efforts to conserve water use (Rösch et al., 2012).

Impacts on Water and Land Use

Algae biomass cultivation depends upon plentiful sources of water. This poses concerns related to competition for scarce freshwater resources. However, algae are naturally abundant in a variety of environmental conditions, including freshwater, saltwater, brackish water, and a number of extreme environments. Both freshwater and saltwater algal strains have been used for demonstration and pilot-scale programs. Saline groundwater can be sourced throughout the United States. Other viable sources are ocean water and municipal wastewater (Farooq et al., 2015). As a result, freshwater resources need not be threatened by algae cultivation. The recycling, treatment, and disposal of water used for algal biomass has also been a topic of research interest because the life-cycle implications of the type of water used and whether it is recycled are significant (Guieysse et al., 2013 Yang et al., 2011).

Land use is also an important consideration, given that converting CO2 waste from a single power plant would require dozens of hectares of biomass cultivation. Since algae cultivation does not require arable land, it will not compete with agriculture and can valorize regions with marginal or saline soils. Like traditional agriculture, the species selection will depend upon local climate. Endemic algae species have been studied for a variety of climates and environmental conditions.

Availability and Suitability of CO2 Waste Streams

To achieve maximal biomass production, CO2 must be supplied to algae cultivation. For many pilot-scale installations, sourcing CO2 has presented a challenge that has been overcome through on-site compressed CO2 storage and delivery. Economic studies have indicated that co-localization of power generation with biomass production can provide significant advantages (Kadam, 1997 Zeiler et al., 1995). Common CO2 concentrations of flue gas from power generation ranges from 12 to 15 mol%, and these concentrations can be efficiently assimilated by algae biomass. Several pilot-scale facilities have implemented this strategy, whereby flue gas is utilized at the site to provide CO2 to algal cultures (Chen et al., 2012 de Morais and Costa, 2007 Wang et al., 2008). Such co-localization strategies dominate technoeconomic analyses, as transportation of CO2 through pipelines or vehicles would require purification and concentration of CO2 from the source and fail to take advantage of the inherent flexibility of photosynthetic microorganisms to utilize a range of CO2 concentrations.

Biomass utilization of CO2 from industrial power plants provides the potential opportunity for large-scale carbon capture. Natural gas&ndashfired power plants offer a waste stream with low sulfur and nitrogen content. The low levels of NOx and SOx present in the waste stream ultimately can be metabolized by most strains of algae and, as such, require considerations primarily for transport, heat exchange, and time of use (Radmann et al., 2011). Coal-fired power plants, on the other hand, present the challenge of additional contaminants. If not adequately purified, these waste streams can include contaminants known to be algecedic such as arsenic, cadmium, mercury, and selenium (Vocke et al., 1980). In addition to posing challenges for biomass cultivation, trace metals and other contaminants that derive from flue gas may limit potential applications of biomass products. For example, products derived from coal-fired flue gas are likely to contain more contaminants, making these sources better suited for biofuel applications than for the production of protein for animal feed.

Examples of a Facultative Anaerobe


A common facultative anaerobe is yeast, used in various cooking applications such as making bread or beer. In either case, this facultative anaerobe must function without oxygen. Yet, the yeast can still survive, and must for these products to come out right.

In bread, yeast is responsible for making the bubbles in the dough. These pockets of air make the bread light and fluffy. Otherwise, the bread would bake into a solid mass more like a cake or brownie. Yeast creates these air pockets through the release of carbon dioxide, a byproduct of converting the glucose in the dough into energy. For a lighter, more airy dough chefs often let the dough “rise”. This term simply means setting the yeast-laden dough in a warm place, and allowing the facultative anaerobe to do its work. Over the course of an hour or so, the yeast will create large amounts of carbon dioxide within the dough, expanding it and making it lighter.

In beer, wine, and other alcoholic beverages, yeast is the key ingredient. The process of fermentation, or the creation of alcohol, occur in yeast when they have plenty of sugar but little oxygen. Brewers and wine-makers use this aspect of the facultative anaerobe to generate the alcohol within their products. Aerobic respiration completely reduces glucose to a few recyclable molecules and carbon dioxide. Fermentation, on the other hand, leaves a final product: ethanol. Beer and wine makers create the ethanol (an alcohol) in their products by strictly controlling the amount of sugar and oxygen in their fermentation tanks. In these conditions any facultative anaerobe will resort to fermentation, and put off ethanol as a byproduct. When the alcohol reaches the proper level in the mixture, the yeast are filtered out and the drink is bottled.


To solve their conundrum, mussels like those in the image above have evolved the abilities of a facultative anaerobe. Instead of relying on their normal aerobic respiration when the tide goes out, the mussels switch to a form of energy which breaks down amino acids. This allows the mussel to survive hours, or even days, without getting a fresh source of oxygen.

1. Humans muscles rely on aerobic respiration to produce the ATP necessary to work them. However, in times of stress and intense exercise, these muscles often run out of oxygen. In this case, the muscles must resort to a form of fermentation which produces lactic acid. Lactic acid can damage cells when it builds up, so the cells must quickly revert to aerobic respiration if they are to survive. Are humans facultative anaerobes?
A. No
B. Yes
C. Maybe

2. What is the difference between a facultative anaerobe and an obligate anaerobe?
A. A facultative anaerobe only has anaerobic pathways.
B. An obligate anaerobe can survive the presence of oxygen.
C. A facultative anaerobe can survive and use oxygen.

3. While scientists used to believe that facultative anaerobe organisms were typically single-celled remnants of an earlier time, evidence has showed that many gut parasites are often facultative anaerobes. Which of the following provides an explanation of this fact?
A. These organisms have a constant access to oxygen.
B. Often, areas of the gut are anaerobic, forcing these organisms to use an anaerobic pathway.
C. These organisms do not represent a facultative anaerobe.

How long does it take for E. coli to shift feedstocks? - Biology

Some prokaryotes and eukaryotes use anaerobic respiration in which they can create energy for use in the absence of oxygen.

Learning Objectives

Describe the process of anaerobic cellular respiration.

Key Takeaways

Key Points

  • Anaerobic respiration is a type of respiration where oxygen is not used instead, organic or inorganic molecules are used as final electron acceptors.
  • Fermentation includes processes that use an organic molecule to regenerate NAD + from NADH.
  • Types of fermentation include lactic acid fermentation and alcohol fermentation, in which ethanol is produced.
  • All forms of fermentation except lactic acid fermentation produce gas, which plays a role in the laboratory identification of bacteria.
  • Some types of prokaryotes are facultatively anaerobic, which means that they can switch between aerobic respiration and fermentation, depending on the availability of oxygen.

Key Terms

  • archaea: A group of single-celled microorganisms. They have no cell nucleus or any other membrane-bound organelles within their cells.
  • anaerobic respiration: A form of respiration using electron acceptors other than oxygen.
  • fermentation: An anaerobic biochemical reaction. When this reaction occurs in yeast, enzymes catalyze the conversion of sugars to alcohol or acetic acid with the evolution of carbon dioxide.

Anaerobic Cellular Respiration

The production of energy requires oxygen. The electron transport chain, where the majority of ATP is formed, requires a large input of oxygen. However, many organisms have developed strategies to carry out metabolism without oxygen, or can switch from aerobic to anaerobic cell respiration when oxygen is scarce.

During cellular respiration, some living systems use an organic molecule as the final electron acceptor. Processes that use an organic molecule to regenerate NAD + from NADH are collectively referred to as fermentation. In contrast, some living systems use an inorganic molecule as a final electron acceptor. Both methods are called anaerobic cellular respiration, where organisms convert energy for their use in the absence of oxygen.

Certain prokaryotes, including some species of bacteria and archaea, use anaerobic respiration. For example, the group of archaea called methanogens reduces carbon dioxide to methane to oxidize NADH. These microorganisms are found in soil and in the digestive tracts of ruminants, such as cows and sheep. Similarly, sulfate-reducing bacteria and archaea, most of which are anaerobic, reduce sulfate to hydrogen sulfide to regenerate NAD + from NADH.

Anaerobic bacteria: The green color seen in these coastal waters is from an eruption of hydrogen sulfide-producing bacteria. These anaerobic, sulfate-reducing bacteria release hydrogen sulfide gas as they decompose algae in the water.

Eukaryotes can also undergo anaerobic respiration. Some examples include alcohol fermentation in yeast and lactic acid fermentation in mammals.

Lactic Acid Fermentation

The fermentation method used by animals and certain bacteria (like those in yogurt) is called lactic acid fermentation. This type of fermentation is used routinely in mammalian red blood cells and in skeletal muscle that has an insufficient oxygen supply to allow aerobic respiration to continue (that is, in muscles used to the point of fatigue). The excess amount of lactate in those muscles is what causes the burning sensation in your legs while running. This pain is a signal to rest the overworked muscles so they can recover. In these muscles, lactic acid accumulation must be removed by the blood circulation and the lactate brought to the liver for further metabolism. The chemical reactions of lactic acid fermentation are the following:

Pyruvic acid + NADH ↔ lactic acid + NAD +

Lactic acid fermentation: Lactic acid fermentation is common in muscle cells that have run out of oxygen.

The enzyme used in this reaction is lactate dehydrogenase (LDH). The reaction can proceed in either direction, but the reaction from left to right is inhibited by acidic conditions. Such lactic acid accumulation was once believed to cause muscle stiffness, fatigue, and soreness, although more recent research disputes this hypothesis. Once the lactic acid has been removed from the muscle and circulated to the liver, it can be reconverted into pyruvic acid and further catabolized for energy.

Alcohol Fermentation

Another familiar fermentation process is alcohol fermentation, which produces ethanol, an alcohol. The use of alcohol fermentation can be traced back in history for thousands of years. The chemical reactions of alcoholic fermentation are the following (Note: CO2 does not participate in the second reaction):

Pyruvic acid → CO2 + acetaldehyde + NADH → ethanol + NAD +

Alcohol Fermentation: Fermentation of grape juice into wine produces CO2 as a byproduct. Fermentation tanks have valves so that the pressure inside the tanks created by the carbon dioxide produced can be released.

The first reaction is catalyzed by pyruvate decarboxylase, a cytoplasmic enzyme, with a coenzyme of thiamine pyrophosphate (TPP, derived from vitamin B1 and also called thiamine). A carboxyl group is removed from pyruvic acid, releasing carbon dioxide as a gas. The loss of carbon dioxide reduces the size of the molecule by one carbon, making acetaldehyde. The second reaction is catalyzed by alcohol dehydrogenase to oxidize NADH to NAD + and reduce acetaldehyde to ethanol.

The fermentation of pyruvic acid by yeast produces the ethanol found in alcoholic beverages. Ethanol tolerance of yeast is variable, ranging from about 5 percent to 21 percent, depending on the yeast strain and environmental conditions.

Other Types of Fermentation

Various methods of fermentation are used by assorted organisms to ensure an adequate supply of NAD + for the sixth step in glycolysis. Without these pathways, that step would not occur and no ATP would be harvested from the breakdown of glucose.Other fermentation methods also occur in bacteria. Many prokaryotes are facultatively anaerobic. This means that they can switch between aerobic respiration and fermentation, depending on the availability of oxygen. Certain prokaryotes, like Clostridia, are obligate anaerobes. Obligate anaerobes live and grow in the absence of molecular oxygen. Oxygen is a poison to these microorganisms, killing them on exposure.

It should be noted that all forms of fermentation, except lactic acid fermentation, produce gas. The production of particular types of gas is used as an indicator of the fermentation of specific carbohydrates, which plays a role in the laboratory identification of the bacteria.