Category Archives: Biofuels from Agricultural Wastes and Byproducts

The Role of Water in HTL

Water plays an essential role in HTL. It is therefore critical to understand the fundamentals of water chemistry when subjected to high temperature conditions. Water is rather benign and will not likely react with organic molecules under standard environmental conditions (20°C and 101,325kPa). However, when the temperature increases, two properties of water mole­cules change substantially. First, the relative permittivity (dielectric constant), er, of water decreases quickly when the temperature increases. When the thermal energy increases, the shared electron by oxygen and hydrogen atoms tends to circulate more evenly and the electro­negativity of the oxygen molecule is reduced (less polar). For example, when temperature increases from 25°C to 300°C, the relative permittivity decreases from 78.85 to 19.66, result­ing in water molecules from very polar to fairly nonpolar, in a relative term. This polarity
change makes water more affinitive to the organic hydrocarbons, most of which are nonpolar molecules.

image068 Подпись: R Подпись: 1 T Подпись: (1)

Second, the dissociation of water dramatically increases with the increase of temperature. Water, like any other aqueous solutions, split into H+ and OH+ ions in hydrolysis or dis­sociation. This process is reversible and the rate is sufficiently rapid so it can be considered to be in equilibrium at any instant. Based on Arrhenius reaction rate, the equilibrium con­stant (or the dissociation constant), Kw, affected by the temperature change, can be written as (Benjamin 2002):

where Kw1 and Kw2 are equilibrium constant at temperatures T1 and T2 , respectively; AEAr is the net change in heat content of the molecules in the overall reaction, also called the molar enthalpy of reaction; R is the universal gas constant, and T is the absolute temperature in Kelvin. AEAr is an empirical constant specific to particular reaction and in units of energy per mole.

Подпись: Temperature dependence of water ionization at 25 MPa (absolute) Figure 10.1. Effect of temperature on water dissociation constant at 25 MPa. The dissociation constant Kw is expressed as pKw, where pKw = -log10(Kw). (IAPWS 2004).

The effect of temperature on water dissociation is illustrated in Figure 10.1, where the left side vertical axis is in pKw = — log10(Kw), and the right side vertical axis is the ratio of Kw to the Kw0. Kw0 is the water dissociation constant at a temperature of 25oC at atmospheric pressure, and is 10+14, From Figure 10.1, water molecules dissocia­tion constant at 300oC is about 500 times higher than that of 25oC at atmospheric pressure. The increase in the dissociation constant will increase the rate of both acid — and base-catalyzed reactions in water far beyond the natural acceleration due to increased temperature.

Pressure dependence of water ionization at 25 deg C

image073

Figure 10.2. Effect of pressure temperature on water dissociation constant Kw at temperature of 25oC. The dissociation constant Kw is expressed as pKw, where pKw = — log10(Kw).

There is also a (usually small) dependence on pressure (ionization increases with increasing pressure). The dependence of the water ionization on temperature and pressure has been fully investigated (Figure 10.2) and a standard formulation exists (IAPWS 2004).

For the above two reasons, water at high temperatures becomes a good solvent for hydro­carbons that are typically nonpolar hydrophobic under standard environmental conditions. These changes in physical properties make the solvent properties of water at 300°C roughly equivalent to those of acetone at 25°C. Ionic reactions of organics should be favored by increased solubility in water. The enhancement of this solubility of hydrocarbons in water will further enhance the possibilities of contact of dissociated H+ with hydrocarbons, hence accelerate the activities of hydrolysis.

The dramatic changes in the physical and chemical properties of water as temperature increases suggest the possibility of organic chemical reactions to take place (Siskin and Katritzky 1991; Kruse and Gawlik 2003). In addition, water has the ability to carry out con­densation, cleavage, and hydrolysis reactions, and to affect selective ionic chemistry, which is not accessible thermally, largely due to changes in its chemical and physical properties, which become more compatible with the reactions of organics as the temperature is increased.

Some classes of organic molecules in biomass proved very susceptible to water’s influence. Hot water as a reactant and catalyst likely creates a second pathway for the cascade of molecular transformations that leads to oil. In this pathway, water causes organic material to disintegrate and reform (by adding H+ to open carbon bond) into fragments that then transform into hydrocarbons. This implies that hot water becomes a catalyst for a series of ionic reac­tions. The acidic and basic nature of hot water—rather than heat—drives this cascade. For example, water may function first as a base, nibbling away at certain linkages in the organic material. As new molecular fragments build up and modify the reaction environment, water can change its catalytic nature. It can then act as an acid, accelerating different reactions. The resulting products attack parts of the remaining molecules, further speeding the breakdown (Siskin and Katritzky 1991). The above analysis may also help to explain why HTL will more likely directly convert biomass into oil than pyrolysis in which water is not involved.

Conclusions

In conclusion, numerous advances have been made in the conversion of biomass to biofuels as biomass is the only source that is renewable and economically available. Problems associated with inhibitor generation and detoxification, fermentation of both hexoses and pentoses to ethanol, and the development of efficient microbial strains have partially been addressed. Simultaneous product recovery and process consolidation and integration will further improve the economics of production of biofuels from biomass. It is emphasized that numerous domestic and international companies have initiated their programs to commercialize conversion of biomass to biofuels. Separation and use of coproducts as additional sources for generating additional revenue can strengthen the approach further. 1

Acknowledgements

Nasib Qureshi would like to thank Adam Wallenfang for his help on finding information on published data. Nasib Qureshi and Stephen Hughes would like to thank Michael A. Cotta (U. S. Department of Agriculture, Peoria, IL) for reading this manuscript and providing critical comments.

Endnote

1. Mention of trade names or commercial products in this article is solely for the purpose of providing scientific information and does not imply recommendation or endorsement by the U. S. Department of Agriculture.

References

Dien, B. S., M. A. Cotta, and T. W. Jeffries. 2003. Bacteria engineered for fuel ethanol production: Current status. Appl. Microbiol. Biotechnol. 63:258-266.

Ezeji, T. C., N. Qureshiand, and H. P. Blaschek. 2007a. Butanol production from agricultural residues: Impact of degradation products on Clostridium beijerinckii growth and butanol fermentation. Biotechnol. Bioeng. 97:1460-1469.

Ezeji, T. C., N. Qureshi, and H. P. Blaschek. 2007b. Bioproduction of butanol from biomass: From genes to bioreactors. Curr. Opin. Biotechnol. 18:220-227.

Grohmann, K. and R. J. Bothast. 1997. Saccharification of corn fibre by combined treatment with dilute sulfuric acid and enzymes. Proc. Biochem. 32:405-415.

Hahn-Hagerdal, B. and N. Pamment. 2004. Microbial pentose metabolism. Appl. Biochem. Biotechnol. 113-116:1207-1209.

Hahn-Hagerdal, B., M. Galbe, M. F. Gorwa-Grauslund, G. Liden, and G. Zacchi (2006) Bio-ethanol—The fuel of tomorrow from the residues of today. Trends Biotechnol. 24: 549-556.

Hughes, S. R., R. E. Hector, J. O. Rich, N. Qureshi, K. M. Bischoff, B. S. Dien, B. C. Saha, S. Liu, E. J. Cox, J. S. Jackson, Jr., D. E. Sterner, T. R. Butt, J. LaBaer, and M. A. Cotta. 2009a. Automated yeast mating protocol using open reading frames from Saccharomyces cerevi — siae genome to improve yeast strains for cellulosic ethanol production. J. Assoc. Lab. Autom. 14(4): 190-199.

Hughes, S. R., J. O. Rich, K. M. Bischoff, R. E. Hector, N. Qureshi, B. C. Saha, B. S. Dien, S. Liu, J. S. Jackson, D. E. Sterner, T. R. Butt, J. LaBaer, and M. A. Cotta. 2009b. Automated yeast transformation protocol to engineer Saccharomyces cerevisiae strains for cellulosic ethanol production with open reading frames that express proteins binding to xylose isom — erase identified using robotic two-hybrid screen. J. Assoc. Lab. Autom. 14(4):200-212.

Ingram, L. O., T. Convey, D. P. Clark, G. W. Sewell, and J. F. Preston. 1987. Genetic engineer­ing for ethanol production in Escherichia coli. Appl. Environ. Microbiol. 53:2420-2425.

Lynd, L., W. H.V Zyl, J. E. McBride, and M. Laser. 2005. Consolidated bioprocessing of cellulosic biomass: An update. Curr. Opin. Biotechnol. 16:577-583.

Mohaghegi, A., N. Dowe, D. Schell, Y. C. Chou, C. Eddy, and M. Zhang. 2004. Performance of a newly developed integrant of Zymomonas mobilis for ethanol production on corn stover hydrolysate. Biotechnol. Lett. 26:321-325.

Nichols, N. N., L. N. Sharma, R. A. Mowery, C. K. Chambliss, P. G. van Walsum, B. S. Dien, and L. B. Iten. 2008 . Fungal metabolism of fermentation inhibitors present in corn stover dilute acid hydrolysate. Enzyme Microb. Technol. 42:624-630.

Nigam, J. N. 2001. Ethanol production from wheat straw hemicelluloses hydrolysate by Pichia stipitis. J. Biotechnol. 87:17-27.

Qureshi, N. 2009 . Solvent production. In: Encyclopedia of Microbiology, edited by M. Schaechter, pp. 512-528. Oxford: Elsevier Ltd.

Qureshi, N. and T. C. Ezeji. 2008. Butanol “A superior biofuel” production from agri­cultural residues (renewable biomass): Recent progress in technology. Biofuels, Bioprod. & Biorefining 2(4):319-330.

Qureshi, N. and G. J. Manderson. 1995. Bioconversion of renewable resources into ethanol: An economic evaluation of selected hydrolysis, fermentation and membrane technologies.

Energy Sources 17:241-265

Qureshi, N., B. Dien, N. N. Nichols, B. C. Saha, and M. A. Cotta. 2006. Genetically engineered Escherichia coli for ethanol production from xylose: Substrate and product inhibition and kinetic parameters. Trans IChemE (Chem. Eng. Res. & Design) 84(C2): 114-122.

Sedlak, M. and N. W. Ho. 2004. Production of ethanol from cellulosic biomass hydrolysates using genetically engineered Saccharomyces cerevisiae yeast capable of cofermenting glucose and xylose. Appl. Biochem. Biotechnol. 113-116:403-416.

Slininger, P. J., S. W. Gorsich, and Z. L. Liu. 2009. Culture nutrition and physiology impact the inhibitor tolerance of the yeast Pichia stipitis NRRL Y-7124. Biotechnol. Bioeng. 102:778-790.

Tran, A. V and R. P. Chambers. 1985. Red oak wood derived inhibitors in the ethanol fermentation of xylose by Pichia stipitis CBS 5776. Biotechnol. Lett. 7:841-846.

Wang, B., T. C. Ezeji, Z. Shi, H. Feng, and H. P. Blaschek. 2009. Pretreament and conversion of distiller’s dried grains with soluble for acetone-butanol-ethanol (ABE) production.

Trans. Am. Soc. Agri. Biol. Eng. 52:885-892.

Anaerobic Digestion in Denmark

Biogas generation from agricultural wastes was first introduced in Denmark in the late 1970s during the first energy crisis. Although not as widespread as on farms in Germany, where many small farms have digesters, Denmark has three categories of digester: (1) 20 community (centralized) biogas plants ranging from 540 to 7500 m3 in volume and fed with animal and industrial wastes, which provide electricity to the grid and heat for houses (Figure 4.8); (2) eight large farm biogas plants with CHP, but without co-digestion; and (3) 18 farm biogas digesters operated at thermophilic conditions in the capacity range of 150-800 m3 with co­digestion and CHP (Al Seadi and Holm-Nielsen 2000; Sannaa 2004).

Nearly all community biogas plants are performing co-digestion, partly for increased biogas production, but also for “tipping fees” from suppliers of organic wastes. The average community digester obtains its income from three sources: electricity, heat, and tipping fees from industries. In general, such plants get -20% to over 50% of their wastes from nonagri­cultural suppliers (e. g., slaughter houses, fish processing, pharmaceutical industry, hospital kitchens, hotels). Drivers for anaerobic digesters in Denmark have been investment grants to help capital costs (-20%), long-term loans at low interest rates, a requirement for a 9-month storage for untreated manure slurry, favorable prices for biogas-produced electricity, oppor­tunities for district heating, and demonstration and research programs.

Biosurfactants, Fatty Acids, and Lipids

Biosurfactants are microbial-derived surface-active agents that can be used as emulsifiers, de — emulsifiers, wetting agents, foaming agents, functional food ingredients, and detergents in various industrial sectors such as petrochemicals, food and beverages, cosmetics and pharmaceuticals, agrochemicals, and fertilizers. They are preferred to chemical surfactants due to their biodegradability, surface properties, and low toxicity. Rhamnolipid is a type of biosurfactants synthesized by Pseudomonas aeruginosa. This microorganism produced 15.4 g/L of rhamnolipid during growth in a basal medium supplemented with crude glycerol (Zhang et al. 2005).

Another type of biosurfactant, called sophorolipid (SL), is produced by yeast Candida bombicola as an extracellular glycolipid. This glycolipid is composed of a disaccharide (i. e., sophorose) attached to a hydroxy fatty acyl moiety at the omega minus one or omega carbon atom (Asmer et al. 1988). The fatty acid (saturated or unsaturated) component of the glyco­lipid is composed of 16-18 carbons. The carboxyl group is either lactonized to the disac­charide ring or free as in the open chain form. Since the hydroxyl groups attached to the disaccharide ring are capable of being acetylated, the whole molecule is amphiphilic with surfactant properties. C. bombicola produced 9 g/L of SL in a pure glycerol medium, but when grown in glycerol waste stream obtained from a biodiesel plant, up to 60 g/L SLs was produced. This dramatic increase in SL production was attributed to low osmotic stress and presence of fatty acids in the crude glycerol (Ashby et al. 2005).

The conversion of glycerol to single cell oil (SCO) by microbes is another commercially important process due to the use of SCO as nutraceuticals, pharmaceuticals, and feed ingre­dients for aquaculture. Microbial lipids have similar properties to vegetable fats and oils in terms of structure and composition and hence have potential to replace them. Y. lipolytica, when grown in a medium containing crude glycerol, produced 8.1 g/L of SCO at the dilution rate of 0.03/h with a maximum productivity of 1.2 g/L/h (Papanikolaou et al. 2002).

Docosahexaenoic acid is an omega-3 polyunsaturated fatty acid (ю — 3 PUFA) that has a protective role against cancers, Alzheimer’s disease, cardiovascular diseases, schizophrenia, and so on. This compound is currently obtained from fish oil but can also be extracted from ю — 3 PUFA-enriched marine algae. The extracted PUFA can be used as a food additive, pharmaceutical, and aquaculture feed ingredient. The marine alga Schizochytrium limacinum produced large amounts of docosahexaenoic acid when grown on either a glucose or a glyc­erol medium. When crude glycerol was used as a carbon source, S. limacinum produced approximately 4.91 g/L docosahexaenoic acid (Chi et al. 2007).

Glycerol can be chemically converted to a variety of commercially important chemicals such as propylene glycol, propanediols, and acrolein. Traditionally, glycerol is esterified to fatty acids to produce polyglycerol esters that are used either as emulsifiers in the food indus­try or as detergents. The price of glycerol has fallen recently, and the production of other high-value chemicals from glycerol has been explored. These include the oxidation of glycerol to glyceric acid and tartronic acid, and the production of glycerol carbonate.

Comparison between Herbaceous Fiber and Grain Systems

Harvesting

There are many different models and sizes of combines for harvesting grain and several dif­ferent models and sizes of cotton harvesters. In like manner, there are a number of different machines for harvesting sugarcane. The manufacturers of harvest machines all state that the capacity of their machine (t/hour), averaged over a day’s operation, is dependent on the infield hauling operation, and, in the case of grain and sugarcane, it is also dependent on the cycling of the highway hauling trucks. This is a key point when the performance of a system of equipment is simulated. Capacity of a “middle” unit in the system depends on the “upstream” unit and the “downstream” unit. Often an advantage can be obtained by uncoupling the harvest operation from the infield hauling operation (sometimes not possible) and by uncou­pling the highway hauling from the infield hauling (sometimes not possible). Let us see how this works when we compare the grain, cotton, and sugarcane systems.

A grain combine has an onboard grain tank. When this tank is full, an infield hauling unit (truck, trailer, or wagon) must come alongside so the grain can be unloaded or the combine must stop and wait. An obvious point must be emphasized. Capacity of the most sophisticated (and expensive) combine is 0 t/hour during the period when it is not moving.

The cotton harvester also has some onboard storage capacity. When this chamber is full, a side-dump trailer must come alongside to receive the seed cotton, or the harvester must stop and wait, thus reducing its average capacity (t/hour).

The sugarcane harvester is an even more dramatic example of the linkage between harvest­ing and infield hauling. The sugarcane harvester, like a forage harvester, has no onboard storage. An infield hauling unit must be alongside to catch the harvested material or the harvest operation stops. Cycling of the infield hauling units is the key to sugarcane harvest­ing. Remember that the infield hauling units must be able to dump at the truck loading station in order to cycle back to the harvester. Now there is a direct coupling of the infield hauling and the highway hauling. When all operations are under central control, like with the sugar mill, an inter-connected logistic system like this can be the most efficient option. Unfortunately, there are few locations, other than the sugar industry (Florida, Louisiana, Texas), where it is practical.

It is appropriate to define a “harvesting unit.” This unit is defined as one (or more) harvester(s) and the pieces of equipment needed to haul material at a rate that keeps the harvester(s) moving. Excess infield hauling equipment and over-the-road equipment (each unit spends time waiting for the harvester) is generally not the most cost-effective organiza­tion of the unit. Trade-offs are always used in organizing a unit. In a commercial operation, each piece of equipment will spend some time waiting—scheduling is never perfect. Because the harvester costs more to operate per hour, it is generally best to minimize harvester wait time.

The harvest units described below are given to provide a reference point. They are not offered as an optimum solution.

Supercritical Fluid Extraction (SFE)

Any substance at a temperature and pressure above its thermodynamic critical point will become supercritical fluid, which can diffuse through solids like a gas and dissolve materials like a liquid (Hawthorne 1990). Additionally, close to the critical point, small changes in pressure or temperature result in large changes in density, allowing many properties to be adjusted. Supercritical fluids may be suitable as a substitute for organic solvents in a range of industrial and laboratory processes.

Persson et al. (2002b) performed countercurrent flow supercritical CO2 (200 bar and 40oC) extraction of hydrolysates. A reduction in the concentration of a variety of inhibi­tors, such as furan derivatives, aliphatic acid, and phenolic compounds, were observed. The effect of the SFE treatment was examined with respect to alcoholic fermentation by Saccharomyces cerevisiae. The ethanol yield increased from 0.30 to 0.43 g/g glucose, and its productivity from 0.14 to 0.46g/Lh. SFE has several advantages such as clean­ness, biocompatibility, and high concentration factor. But the capital cost for SFE is usually high.

Encapsulation

With cell encapsulation, fermenting yeasts are protected by an artificial membrane, and successful fermentation with toxic hydrolysates has been reported (Talebnia et al. 2005) . Talebnia et al. (2005) used encapsulated S. cerevisiae CBS 8066 to ferment two different types of dilute-acid hydrolysates in the presence of furfural (0.39 and 0.31 g/L) and hydroxy — methylfurfural (HMF; 0.74 g/L and 1.58g/L). While the free cells were not able to ferment the hydrolysates in 24 hours, the encapsulated yeast successfully converted glucose and mannose in the hydrolysates in less than 10 hours with minimal lag phase. Talebnia and Taherzadeh (2006) further demonstrated that encapsulation is a promising method to keep the cells viable in a toxic environment and help the process to run continuously at high dilution rates and high productivities. The physiological and morphological characteristics of the encapsulated S. cerevisiae CBS 8066 were studied by Talebnia and Taherzadeh et al. (2007). After 20 consecutive batch cultivations in a defined synthetic medium, the ethanol yield increased from 0.43 to 0.46 g/g, while the biomass and glycerol yields decreased by 58% and 23%, respectively. The growth rate of the encapsulated cells in the first batch was 0.13/hour, but decreased gradually to 0.01/hour. After long-term application, most of the encapsulated yeast existed in the form of single and non-budding cells. Total RNA content of these yeast cells decreased by 39%, while the total protein content decreased by 24%. On the other hand, the stored carbohydrates (glycogen and trehalose) content increased. Because of the higher biomass concentrations inside capsules, the glucose dif­fusion rate through the membrane drastically decreased to 1/5 of that seen in cell-free capsules.

Molecular Sieve

Molecular sieves are used as adsorbents for gases and liquids. Molecular sieves have tiny pores of a precise and uniform size. Molecules that are small enough to pass through these pores will be adsorbed, while larger molecules are not. For instance, Tran and Chambers (1986) treated unfermentable red oak hydrolysates with a molecular sieve. The treatment with the molecular sieve decreased the concentration of acetic acid by 40% and furfural, by 82%. Treatment of hydrolysates with molecular sieve, however, resulted in a 10% loss in xylose concentration.