Technological advances in the major process components — fermentation, primary purification, and secondary purification and polymerization/chemical conversion of lactic acid and its derivatives — have recently occurred. These and other advances would enable low-cost, large-volume, and environmentally sound production of lactic acid and its derivative products.

In fermentation, high (> 90%) yield from carbohydrate, such as starch, is feasible, together with high product concentration (90 g/L, 1 M). Stable strains with good productivity (> 2 g/L h) that utilize low-cost nutrients (such as com steep liquor) are available. Furthermore, the fermentation is anaerobic and thus has low power and cooling needs. All of these make the fermentation step very facile and inexpensive.

Recent advances in membrane-based separation and purification technologies, particularly in micro — and ultrafiltration and electrodialysis, have led to the inception of new processes for lactic acid production. These processes would, when developed and commercialized, lead to low-cost production of lactic acid, with a reduction of nutrient needs and without creating the problem of by-product gypsum (18-21). Desalting electrodialysis has been shown to need low amounts of energy to recover, purify, and concentrate lactate salts from crude fermentation broths (19). The recent advances in water-splitting electrodialysis membranes enable the efficient production of protons and hydroxyl ions from water and can thus produce acid and base from a salt solution (20-21). These advances have led to the development of proprietary technologies for lactic acid production from carbohydrates without producing salt or gypsum by-products (19-20). In recently issued patents to Datta and Glassner (19-20), an efficient and potentially economical process for lactic acid production and purification is described. By using an osmotolerant strain of lactic acid bacteria and a configuration of desalting electrodialysis, water-splitting electrodialysis, and ion-exchange purification steps, a concentrated lactic acid product containing less than 0.1% of proteinaceous impurities could be produced from a carbohydrate fermentation. The electric power requirement for the electrodialysis steps was approximately 0.5 kWh/lb (~1 kWh/kg) lactic acid. The process produces no by-product gypsum, only a small amount of by-product salt from the ion-exchange regeneration. Such a process can be operated in a continuous manner, can be scaled up for large-volume production, and forms the basis for commercial developments for several companies that have announced intention to be commercial producers of lactic acid and its derivative products (3-4).

Another recent entrant, Ecochem, a DuPont-ConAgra partnership, had developed a recovery and purification process that produces a by-product ammonium salt instead of insoluble gypsum cake and attempted to commercialize

the process and sell this by product as a low-cost fertilizer. A 10 — ton/yr demonstration-scale plant was recently completed to prove the process and develop products and markets for polymers and derivatives. However, due to poor choice of feedstock (whey) and purification process technology, this attempt failed. The polymer patent portfolio of DuPont was acquired by Chronopol, Inc. which is building a demonstration-scale plant for lactic acid polymer production.

The utilization of the purified lactic acid to produce polymers and other chemical intermediates requires the development of secondary purification and integration of catalytic chemical conversion process steps with the lactic acid production processes. Examples of such process steps would be dilactide production for polymerization to make high-molecular-weight polymers or copolymers and hydrogenolysis to make propylene glycol — a large-volume intermediate chemical. In the past, very little effort was devoted to develop efficient and potentially economical processes for such integrations, because only small-volume, high-margin specialty polymers for biomedical applications or specialty chemicals were the target products.

Recently, several advances in catalysts and process improvements have occurred and proprietary technologies have been developed that may enable the commercialization of integrated processes for large-scale production in the future. In a recent patent issued to Gruber et al. of Cargill, Inc. (22), the development of a continuous process for manufacture of lactide polymers with controlled optical purity from purified lactic acid is described. The process uses a configuration of multistage evaporation followed by polymerization to a low-molecular-weight prepolymer, which is then catalytically converted to dilactide, and the purified dilactide is recovered in a distillation system with partial condensation and recycle. The dilactide can be used to make high-molecular-weight polymers and copolymers. The process has been able to use fermentation-derived lactic acid, and the claimed ability to recycle and reuse the acid and prepolymers could make such a process very efficient and economical (22). In recent patents issued to Bhatia et al. of DuPont, Inc. (23-30), processes to make cyclic esters, dilactide, and glycolide from their corresponding acid or prepolymer are described. This process uses an inert gas, such as nitrogen, to sweep away the cyclic esters from the reaction mass and then recovers and purifies the volatilized cyclic ester by scrubbing with an appropriate organic liquid and finally separates the cyclic ester from the liquid by precipitation or crystallization and filtration of the solids. Very high purity lactide with minimal losses due to racemization have been claimed to be produced by this process. Recycle and reuse of the lactic moiety in the various process streams have been claimed to be feasible (30). DuPont’s patents have been acquired by Chronopol and both Cargill and Chronopol are developing their processes to commercial scale; their goal is large-scale production of biodegradable polymers in the future.

Hydrogenolysis reaction technology to produce alcohol from organic acids or esters has also advanced recently — new catalysts and processes yield high selectivity and rates and operate at moderate pressures (31-33). This technology has been commercialized to produce 1,4 butanediol, tetrahydrofuran, and other four-carbon chemical intermediates from maleic anhydride. In the future, such technologies could be integrated with low-cost processes for the production of lactic acid to make propylene glycol and other intermediate chemicals (34).

Technical Accomplishments and Future Directions at Argonne

In the past two years, under a U. S. Department of Energy-sponsored project at ANL, several important technical advances have been achieved and demonstrated at the laboratory scale. Notably, these advances have occurred in fermentation, primary purification, and polymer synthesis. In fermentation, high product yield (95%) from starch by means of an enzymatic saccharification/fermentation process with high lactate concentration (100 g/L) and good productivity (3 g/L*h) have been achieved. The ED-based primary purification process has been operated in the laboratory in short-term feasibility experiments to obtain flux and power data for design and economics. A proprietary method to produce a high-molecular-weight copolymer of polylactic acid with other copolymers has been developed at the laboratory scale. Methods to modify and test the degradability of polylactic acid have been developed. Furthermore, the development of secondary purification processes and specialty products derived from lactic acid with targeted properties have been initiated.

The ANL program of oxychemicals and polymer feedstock production from carbohydrate-derived lactic acid is schematically shown in Figure 1. The fermentation and primary purification process to make purified lactic acid has been developed and demonstrated at ANL and elsewhere. The program is now focusing on developing efficient and economical secondary purification processes to make esters that can serve as the key intermediate for the production of a host of other chemicals, polymers, and specialty derivatives. The products and the processes to be developed or integrated are shown in Figure 1. This matches several of the target products listed in Table I that can be derived from lactic acid.

Thus, a rational program targeted at development of economical processes for key intermediates of lactic acid and its derivative products has been the primary focus at Argonne.

Conclusions

9

A wide range of products with U. S. market size exceeding 6×10 lb/yr and product

# 9

values exceeding $4×10 /yr could be potentially manufactured from lactic acid.

Degradable and environmentally sound products will provide the initial impetus for development and deployment of new lactic acid technologies and products.

Several major U. S. agriprocessing/chemical companies have built demonstration-scale plants and have identified the trends in the environmentally sound products and processes; consequently, they have plans for major large-scale plants in the future.

Novel separations processes that have recently emerged can enable large-scale and economical production of purified lactic acid without waste gypsum or salt by-product.

В

Figure 1. ANL Program of Oxychemicals and Polymer Feedstock Production from Carbohydrate Derived Lactic Acid

Several novel processes are being deployed for facile production of lactic polymer feedstock from lactic acid.

A wide variety of polymers/copolymers with many potential consumer uses could be derived as these products and processes are brought on-stream.

With the new technologies, the manufacturing costs and economics of lactic acid and its derivatives have an attractive potential in large-scale systems.

The lactic acid program at ANL has achieved several important milestones mainly in fermentation and methods of copolymer development.

The technical strategy of the program is to develop novel and economical technologies for key intermediates and products (beyond the degradable polymers) that have a wide range of potential applications.

Acknowledgments

Work supported by the U. S. Department of Energy, Assistant Secretary for Energy Efficiency and Renewable Energy, under Contract W-31-109-Eng-38..

The industrial enzyme market approaches approximately one billion US dollars annually. Several enzymes have already become commodity chemicals for many industrial application purposes such as in the production of various com syrups and sweeteners and fuel ethanol from starch. Right now, the market for the enzymes involved in various lignocellulosic biomass conversion is limited and depends entirely on their use in the conversion of various lignocellulosic feedstocks to fermentable sugars for the subsequent production of fuel alcohols and value-added chemicals. Currently, cellulolytic enzymes are expensive and their hydrolysis rates are very slow. The development of an environmentally compatible highly efficient enzyme system free from product and substrate inhibitions for conversion of various pretreated agricultural residues to glucose is very important for use of these materials for production of fuel alcohol. The market for these enzymes will expand rapidly if certain properties of them can be improved and if these enzymes are made available for biomass conversion at a competitive price like starch degrading enzymes. On the other hand, the development of a very efficient substrate pretreatment that increases the susceptibility of crystalline cellulose to enzymatic hydrolysis significantly will lower the cost of producing ethanol from lignocellulosic biomass.

Economic reasons have been one of the major obstacles in the use of biodiesel. Diesel fuel (DF) derived from vegetable oils is more expensive than petroleum-based DF. The feedstock for biodiesel is already more expensive than conventional DF. For example, in the United States, a gallon of soybean oil costs approximately two to three times as much as a gallon of conventional DF. However, in the case of conversion of vegetable oils or fats to their esters, the resulting glycerol co-product, which has a potential market of its own, may offset some of the costs.

In most European countries, however, transportation fuels are so heavily taxed that tax incentives can be applied to encourage the use of biodiesel in the form of lower or no taxes on the biofuel and higher taxes on the petroleum-based fuel (3,4). This subsidy artificially cheapens the biodiesel to make it competitive. In many developing countries, the overriding concern is to become independent of the imported commodity petroleum. In the United States, the tax mechanism is inapplicable because of the comparatively low taxes on transportation fuels. Artificially regulating the demand for fuels from specific sources by means of taxation is currently politically not feasible.

Nevertheless, biodiesel is attractive for other reasons. Besides being a renewable resource and therefore creating independence from the imported commodity petroleum and not depleting natural resources, health and environmental concerns are the driving forces overriding the economic aspects in some cases. These concerns are manifested in various regulatory mandates of pollutants, particularly CAAA (Clean Air Act Amendments of 1990) and EPACT (Energy Policy Act of 1992) in the United States, which present opportunities for alternative fuels such as biodiesel. A life-cyle analysis of biodiesel (5) has shown that it is competitive with other alternative fuels such as compressed natural gas (CNG) and methanol in the urban transit bus market.

It is generally recognized that biodiesel has lower emissions, with the exception of nitrogen oxides (NOJ, than conventional petroleum-based DF. For example, due to its lack of sulfur, biodiesel does not cause S02 emissions. The lower emissions have caused biodiesel to be used in urban bus fleets and to make it especially suitable for other niche markets such as mining and marine engines. Besides environmental and health reasons with accompanying Government regulations, focusing on the use of biodiesel in niche markets is rendered additionally attractive because not enough vegetable oil is produced to supply the whole diesel market with biodiesel.

Numerous reports exist showing that fuel economies of certain biodiesel blends and conventional DF are virtually identical. In numerous on-the-road tests, primarily with urban bus fleets, vehicles running on blends of biodiesel with conventional DF (usually 80% conventional DF and 20% biodiesel; for a list of most biodiesel demonstration programs in the United States, see Ref. 6) required only about 2-5% more of the blended fuel than of the conventional fuel. No significant engine problems were reported as discussed later.

C. S. Gong, Ningjun Cao, and G. T. Tsao

Laboratory of Renewable Resources Engineering, Purdue University,
West Lafayette, IN 47907

A simple and effective method of treatment of lignocellulosic material was used for the preparation of poplar wood chip and com cob for the production of 2,3-butanediol by Klebsiella oxytoca ATCC 8724 in a simultaneous saccharification and fermentation (SSF) process. During the treatments, lignin and hemicellulose fractions of lignocellulosic materials were sequentially removed by aqueous ammonia (10-20%) steeping at 24°C for 24 h followed by, dilute hydrochloric acid (1%, w/v) hydrolysis at 100-108°C for 1 h. The cellulose fractions (80 g/L) were then converted to butanediol by K. oxytoca in the presence of a fungal cellulase (8.5 g IFPU per g cellulose). The butanediol concentrations of 24 and 25.5 g/L, and ethanol concentrations of 6 and 7 g/L were produced by K. oxytoca from wood chip and com cob, respectively. The average butanediol volumetric productivity was 0.26 g/L/h from wood chip and 0.35 g/L/h from com cob.

2,3- Butanediol (2,3-butylene glycol) is a metabolic product of simple carbohydrates produced by many species of enterobacteria through a fermentative metabolic pathway (/). It is a colorless and odorless liquid that has a high boiling point of 180­184° C and a low freezing point of -60°C. The heating value of butanediol (27,198 J/g) is very similar to that of ethanol (29,055 J/g) and methanol (22,081 J/g) which makes it a potentially valuable liquid fuel and fuel additive (2). Butanediol can be dehydrated to methyl ethyl ketone (MEK) and used as an octane booster for gasoline or as high-grade aviation fuel (2). MEK can also be further dehydrated to 1,3- butadiene and dimerized to styrene (5). Therefore, butanediol has a diverse industrial usage, particularly as a polymeric feedstock, in addition to its use for manufacturing butadiene or antifreeze. Currently, butanediol is enjoying an annual growth rate of 4 to 7 percent, buoyed by the increased demand for polybutylene terephthalate resins, y — butyrolactone, Spandex, and its precursors (4).

© 1997 American Chemical Society

2,3- Butanediol is the only isomer, among many, that can be produced by microorganisms. Bacterial species, particularly those belonging to Klebsielleae, are known to metabolize carbohydrates to produce neutral compounds such as 2,3- butanediol, acetoin, and ethanol as metabolic products. Other groups of enterobacteria, such as Erwinia, produced

channeled into butanediol production. The enzymes of the butanediol pathway can constitute as much as 2.5% of the total protein in E. aerogenes (18).

Zeng et al. (19) investigated the effect of pH on growth and product formation of glucose by E. aerogenes in a continuous culture operation and found the optimal pH range of 5.5-6.5 for butanediol and acetoin production. Similar pH optimum was also observed in K. oxytoca (7). In general, the biomass concentration increases steadily with increased pH. At high pH, butanediol concentration decreases with the increase of acetic acid production. Acetic acid has a dual role in the regulation of butanediol formation. It serves as the activator for butanediol accumulation at low concentrations, and at a concentration of 10 g/L or higher, it inhibits butanediol production (19-21). The strength of acetic acid inhibition depends on the concentration of its undissociated form, HAc. The concentration of HAc was in turn determined by the pHs.

The production of butanediol from lignocellulosic materials has been considered as an alternative approach in the conversion of biomass substrates to liquid fuels and chemical feedstocks (3,22). Over the years, there have been many studies conducted utilizing agricultural residues and wastes for butanediol production. The materials studied include: citrus waste (23), water hyacinth (24), wheat and barley straws (25), com stover (25), and hard wood hemicellulose fraction (25-30).

For the biological production of 2,3-butanediol to be economically competitive with petrochemical-based processes, the substrate source must be inexpensive, while reactor yields and productivity should be high. Lignocellulosic materials from forestry and agriculture residues, such as wood chips and com crop residues, are inexpensive and abundant and can be used in many bioprocesses for the production of products of high economical value. An effective utilization of xylose, arabinose, and other minor sugars in addition to glucose is important in the process economics. K. oxytoca ATCC 8724 is capable of producing butanediol from both hexoses and pentoses with good yield (7). In this study, we used K. oxytoca to produce butanediol from pretreated poplar wood chip and ground com cob cellulose fraction in the presence of a fungal cellulase.

Normal Fermentation. Figure 1 shows a normal industrial batch fermentation as described above. Similar patterns were seen in other runs, with some differences of rate. The free glucose is seen to decrease smoothly from an initial level of about 60 g/1. This level reflects the balance between glucose production by the glucoamylase and its consumption by the yeast. Ammonia and FAN decrease rapidly; both are largely consumed by 8 h and exhausted by 12 h (Figure la). A portion of the FAN is not utilizable. The increase in viable cell count is essentially complete by 9h (Figure lb), the rate of glucose utilization slows at his point, and the general pattern is consistent with nitrogen limitation. The initial cell viability is low, typically 60­70%; this increases to the 88-93% range by 4 h and remains nearly constant within that range through 40 h (viability data are not plotted on the graph). Fermentation, as reflected by ethanol production, was 82% complete at 20 h and 94% complete at 30h. Essentially all of the starch was converted to glucose and fermented by 37 h. Glycerol production in the industrial fermentation appears to be largely growth-

Figure la. Substrate Levels, Control Run

log viable —♦— glycerol —0— ethanol/10 —■— C02, mM count

Figure lb. Product Levels, Control Run

и g/l. h —■ — g/vc. h

Figure lc. Glycerol Productivity, Control Run

associated; the productivity on a per-cell basis drops abruptly after 8 hours (Figure lc), at the same time the N sources are exhausted. The high starting glycerol level is due to the recycle of thin stillage and glycerol produced in the yeast propagator; there is a further increase in glycerol concentration in the beer well and still (not shown), possibly due to breakdown of cell components. The dissolved carbon dioxide concentration in a control run is shown in figure lb; it increased from 27 mM at the start to 35 mM at the time of peak fermentation, then settled back to about 29 mM. These concentrations correspond to partial pressures of 1.5, 1.7, and 1.3 atm (solubility calculated separately for the estimated medium composition at each point), confirming that supersaturation is significant in this system.

Laboratory Fermentations. We set up a series of laboratory fermentations to test the effects of CO2. The conditions of inoculum, pH (4.0), temperature (32.5 °С), ammonium and FAN and the initial glucose concentration were initially set to simulate the industrial process, and the glucose feed rate was set to simulate the rate of release of free glucose by glucoamylase in the industrial process. In one run, a low level of oxygen was added as air along with 1.5 atm CO2. The results of this series of experiments are summarized in Table II. Peak viable cell counts (5×10? to 10^ cells/ml) were lower than the levels seen in the plant but peak viability was typically 95-96%. The limiting factor or factors were not identified. Ammonia was not depleted and FAN was not reduced to the levels seen in the plant, so usable N was not limiting. Unlike the industrial fermentation, the lab runs showed decreasing viability after the peak cell count was reached.

Carbon Dioxide Effects. Carbon dioxide at 3.5 atm was somewhat inhibitory to cell growth (Table II). The peak cell count was decreased relative to the lower CO2 levels, and the rate of cell death was slightly increased compared with the runs at 0.5 and 2.5 atm (cell death in the 1.5 atm run was biphasic and cannot be readily compared with the others). Peak viable count was also somewhat reduced at 2.5 atm compared with the lower CO2 levels. An increase in cell size was noticed at

2.5 and 3.5 atm, but no unusual budding was observed. The fermentation rate decreased with increasing carbon dioxide concentrations (Table III); this effect was confined to roughly the last half of the fermentation. There were substantial differences between treatments at 30 and 45 h, but by 65 h ethanol had reached comparable levels in all the runs except possibly the run at 3.5. The ethanol production measured for the 3.5 atm run does not account for all the glucose apparently consumed; we have no ready explanation for this discrepancy but the fermentation rate is still depressed when measured as glucose consumption.

Percentage completion was calculated as the increase in alcohol concentration through the indicated time divided by the total increase in alcohol concentration when all glucose was consumed. *glucose feed was started late, at 20h, in the 2.5 atm experiment, contributing to the low completion at 30 h.

Glycerol and Ethanol Production Kinetics. Both glycerol and ethanol accumulated most rapidly while the cell number was still increasing; volumetric productivity declined after the peak cell number was reached (Figure 2). Although this would suggest that the production of both glycerol and ethanol was largely growth associated, a more detailed analysis disputes that conclusion. When the glycerol productivity is calculated on a per-viable-cell basis, most of the runs show a similar pattern: Initially high per-cell productivity, about 7xl0"^g/(cell. hour) declines within the first 6 hours to a plateau which persists through the remainder of the growth phase, stationary phase, and death phase (Figure 2a-c). Since the cell number is still increasing at 6 h, the higher glycerol productivity in the early periods is more specifically related to rapid growth than to growth per se. The bulk of the glycerol accumulation occurs later when the per-cell accumulation rate is constant, and cell growth has ceased; thus it is not growth-associated. The glycerol accumulation rate in this period is greater than that in the comparable period of the industrial fermentation. Overall glycerol yield was greatest at 0.5 atm CO2 and least at 3.5 atm CO2 among the non-aerated runs. An exception to this pattern was the air-supplemented run. Besides eliminating the initial spike air supplementation decreased the per-cell productivity during the plateau phase. However, the cell count in this run was enough higher to counteract this decrease so that the overall glycerol yield was little different than the control. The total oxygen uptake

time, h
g/l-hr	g/vohr
Figure 2a. Glycerol Productivity, 3.5 Atm C02. g/l-hr —■— g/vc-hr
Figure 2b. Glycerol Productivity, 1.5 Atm C02.

g/l. h, 3.5 atm —0— g/lh, 0.5 atm —■— g/vc. h, 3.5 —°— g/vc. h, 0.5 atm atm

Figure 2d. Ethanol Productivity

amounted to 5.8 mmol/1 based on the flow rate and composition of the gas streams entering and leaving the fermentor.

Ethanol productivity did not follow as simple a pattern and declined more slowly (Figure 2d), but also showed more rapid accumulation in the early part of the run and a plateau of decreased but continuing per-cell productivity as the run progressed (Figure 2d). The changes in per-cell productivity during the run were influenced by the CO2 level. Ethanol production was judged not to be strongly growth-associated.

In-plant Experiments. We attempted to manipulate the fermentation in the plant, first by carrying out a fermentation under partial vacuum. Due to the limited capacity of the Roots blower used as vacuum pump, we could only achieve appreciable vacuum after the rate of CO2 release had slowed. We ultimately reduced the headspace pressure to about 0.6 atm absolute, but had no impact on the level of dissolved carbon dioxide during the critical middle part of the run. The fermentation timecourse and the glycerol yield were similar to the control run (data not shown).

We also carried out an air-supplemented run in the plant. The fermentor was aerated at 20 cfm through sintered metal spargers, and the headspace gas was recirculated at about 40 cfm through separate perforated-pipe spargers. Figure 3c shows that there was a small impact of CO2 levels, as shown by the reversal of the CO2 trace when the sparge was shut off. However, the lowered CO2 concentration was still at the level seen in the control run and there was no improvement in the fermentation kinetics. As in the lab run, aeration led to a slightly higher cell count and altered the pattern of glycerol accumulation. However this time the early growth-associated glycerol production was not eliminated, though it was reduced on a per-cell basis (Figure 3c). Later in the run, second wave of glycerol production occurred which brought the overall glycerol yield up to a level comparable to the control run.

While most aspects of biodiesel discussed above have received considerable attention, relatively few papers {165-167) deal with the aspect of (storage) stability of biodiesel or fatty alkyl esters. The use of biodiesel is advantageous compared to conventional diesel fuel from the aspect of handling and storage safety because of the higher flash point of both vegetable oils and their methyl esters.

Generally, the stability of fatty compounds is influenced by factors such as presence of air, heat, traces of metal, peroxides, light, or structural features of the compounds themselves, mainly the presence of double bonds. The more conjugated or methylene-interrupted double bonds in a fatty molecule, the more susceptible the material is to oxidation and degradation.

Early storage tests gave the following decreasing order of stability for different refinement grades of various vegetable oils {165): soybean oil» degummed soybean oil > refined soybean oil = refined sunflower oil > degummed sunflower oil = crude sunflower oil. The stability of the crude and degummed oils was significantly improved by the addition of diesel fuel (in 1:1 mixtures) but this did not improve the stability of refined oils. The storage stability of 1:1 mixtures were in the decreasing order of crude soybean oil £ crude sunflower oil > degummed soybean oil > degummed sunflower oil » refined soybean oil > refined sunflower oil. A degummed oil / diesel blend with better stability characteristics than that of a refined oil / diesel blend could be prepared. Additionally, the purity of the degummed oils was sufficiently improved by the addition of diesel fuel to meet the required fuel specification.

A study on the stability of the methyl and ethyl esters of sunflower oil reports that ester fuels (biodiesel) should be stored in airtight containers, the storage temperature should be < 30°С, that mild steel (rust-free) containers could be used, and that tert.- butylhydroquinone (TBHQ), an oxidation inhibitor, has a beneficial effect on oxidation stability (766). Methyl esters were slightly more stable than ethyl esters. Light caused only a small increase in the oxidation parameters of esters stored at the high temperature level. The changes in the samples were reflected by increasing acid and peroxide values in storage at 50 °С and increases in ultraviolet (UV) absorption.

Two parameters, namely temperature and the nature of the storage container, were claimed to have the greatest influence on the storage stability (167). Samples stored in the presence of iron behaved differently than those stored in glass. Higher temperature favored degradation of the hydroperoxide at a faster rate than when it was stored at room temperature. Secondary oxidation products were formed in greater amounts in the presence of iron (from the primary peroxides) while in glass the concentration of primary oxidation products is higher. Acidity values were also monitored in this work. Even for samples stored at 40°С, the increase in free acids was within the limits of technical specifications. The free acids need to be controlled because they are mainly responsible for corrosion.

The enzymatic production of xylitol from xylose using xylose reductase of C. pelliculosa

coupling with the oxidoreductase system of Methanobacterium sp. capable of recycling NADP (H) has been demonstrated by Kitpreechavanich et al. (59). A sulfonated polysulfone membrane reactor for in situ regeneration and retention of coenzymes NADP (H) using the xylose reductase of C. pelliculosa coupled with oxidoreductase system of Methanobacterium sp. in the reduction of xylose to xylitol with hydrogen gas was also used (60). The membrane rejected the permeation of NADP (H) (92 and 97%) F420 (97%) and the required enzymes (100%) almost completely, but did not reject for the permeation of xylitol. Nishio et al. (67) reported the enzymatic conversion of xylose into xylitol by the immobilized cells of C. pelliculosa (NADP+ dependent xylose reductase) coupled with the immobilized cells of Methanobacterium sp. HU (hydrogenase and F420-NADP+ oxidoreductase) using hydrogen as an electron donor. The continuous production of xylitol in a column reactor packed with the co­immobilized cells could operate stably for 2 weeks. Xylitol was produced from xylose using commercial immobilized xylose isomerase from Bacillus coagulans and immobilized cells ofM smegmatis (30). From 10 g xylose, 4 g of xylitol was produced and 5 g xylose remained in the reaction mixture; no xylulose was detected. The washed cells of M smegmatis converted xylulose to xylitol under aerobic and anaerobic conditions. The washed cells of a gluconate-utilizing Corynebacterium strain grown in a gluconate-xylose medium produced xylitol from xylose in the presence of gluconate (29). Xylose was reduced to xylitol by coupling the xylose reductase activity to the 6- phosphogluconate dehydrogenase activity with NADP as a cofactor using cell-free extract and the fractionated enzymes of Corynebacterium strain.

The promise of highly efficient conversion processes coupled to a "green" technology is now universally appealing to industry and governmental policymakers. In general, hydrolytic enzymes offer depolymerization of naturally occurring polymers in high yield, with few, if any, by-product disposal problems, unlike acid-based hydrolysis processes. More than a decade ago, advances in the production of fungal cellulase preparations rekindled interest in enzyme-based biomass conversion processes. However, there is some evidence that this technology has now reached its zenith. Also, cellulase biochemistry has reached an enabling phase of development, in which combined efforts in biochemistry and molecular biology may be able to deliver improved cellulase systems for industrial application. Many factors that govern cellulase component action on crystalline cellulose (cellulase synergism, crystallographic structures, cellulase active site structure/fiinction relationships) have been established (or at least preliminarily elucidated), so design of engineered recombinant cellulase systems by means of advanced concepts, such as enzyme component selection and site-directed mutagenesis (SDM), is now at hand. To be commercially viable, engineered cellulase systems must ultimately produce highly active cellulases with improved specific activities, protein yield, and/or production cost relative to current submerged culture fimgal preparations. Once these goals are achieved, the ability to "tune" recombinant systems should permit access to new process feedstocks and markets.

Cellulosic Biomass and Cellulase Action. Biomass feedstocks most commonly considered for conversion to bioethanol in the near term are waste wood, agricultural

wastes, and the paper fraction of municipal solid waste. The fermentable fractions of these feedstocks include cellulose (p-l,4-linked glucose) and hemicellulose. Although it is an abundant biopolymer, cellulose is unique because it is highly crystalline, water insoluble, and highly resistant to depolymerization. Three features of the cellulose in pretreated biomass make it extremely resistant to enzymatic hydrolysis.

First, the p-l,4-glycosidic linkage renders cellulose oligomers (cellodextrins) with a chain length in excess of six glucose residues extremely insoluble in aqueous systems. Cellulose chain lengths in plant cell walls are very much longer, and therefore, completely insoluble in water.

Second, in plant tissues, cellulose is organized by extensive hydrogen bonding and hydrophobic interaction into semicrystalline bundles of parallel cellulose chains, called microfibrils. This structural feature contributes to the rigidity and strength exhibited by wood and simultaneously protects most of the glycosidic bonds from attack by cellulases.

Third, acid pretreatment alters the chemistry of biomass to yield a substrate totally unlike materials encountered in nature. Specifically, the acidic cooking of biomass at temperatures above the phase transition of lignin (about 160°C) leads to redistribution of lignin during cool-down, perhaps with the effect of "coating" residual polysaccharide fibers. It is reasonable to assume that this redistribution of lignin inpedes enzyme action through strong, nonproductive binding. This hypothesis follows, because several studies have shown a strong tendency for lignin in wood to interfere with cellulase action (58-59). Pretreated hardwood pulp also harbors a weak net negatively charged surface not associated with native wood, because regions of hemicellulose adjacent to 4-<9-Me — glucuronate branches are protected during dilute acid hydrolysis and probably remain in the fiber {60-61) and glucuronate content in cellulose increases as a function of normal wood oxidation (62). The result of the alteration of wood surface chemistry following pretreatment most certainly alters interactions between cellulases and the biomass surface.

As a consequence of these properties, converting cellulose to glucose requires substantially larger ratios of enzyme to substrate than for otherwise similar processes which convert starch to glucose. Despite this fact, it is remarkable that an amount of cellulose approximately equivalent to that synthesized during the yearly growing season is recycled on a global scale during the same period of time in the environment.

Cellulose is enzymatically degraded to glucose by the synergistic action of three distinct classes of enzymes: the "endo-l,4-p-glucanases" or 1,4-p-D-glucan 4-glucanohydrolases (EC 3.2.1.4), which act randomly on soluble and insoluble 1,4- p — glucan substrates, the "exo-l,4-p-D-glucanases," including both the 1,4-p-D-glucan glucohydrolases (EC 3.2.1.74), which liberate D-glucose from 1,4-p-D-glucans and hydrolyze D-cellobiose slowly, and 1,4-p-D-glucan cellobiohydrolase (EC 3.2.1.91), which liberates D-cellobiose from 1,4-p-glucans, and the "p-D-glucosidases" or p-D — glucoside glucohydrolases (EC 3.2.1.21), which act to release D-glucose units from cellobiose and soluble cellodextrins, as well as an array of glycosides. The concepts of exo-endo and exo-exo synergism are shown diagrammatically in Figure 1.

Synergism between exo — and endoglucanases is best explained in terms of providing new sites of attack for the exoglucanases. These enzymes normally find available chain ends at the reducing and nonreducing termini of cellulose microfibrils. Each random internal cleavage of surface cellulose chains by an endoglucanase provides two additional sites for attack by cellobiohydrolases. Therefore, each hydrolytic event by an endoglucanase yields both a new reducing and a new nonreducing chain end. Thus, logical consideration of catalyst efficiency dictates the presence of exoglucanases specific for reducing and nonreducing termini, which has now been confirmed for Trichoderma reesei CBH I and CBH II (63). Exo-exo synergism may be explained by considering the tendency of cellulose chains to "reanneal" or return to the crystallite surface following hydrolysis if unimpeded by the presence of an exoglucanase. It is possible that maximal initiation of the processive hydrolytic process catalyzed by exoglucanases occurs only if both exoglucanases (reducing and nonreducing specific) are present at the site of internal bond hydrolysis immediately following endoglucanase action. In order to reduce the effects of steric hinderance at this site, both exoglucanases must bind to their respective substrates with high precision.

As of early 1996, glycosyl hydrolases have been grouped into 56 families of related proteins, based on amino acid sequence homology (64-66) and hydrophobic cluster analysis (67) of catalytic domains. Cellulases are included in 11 of the 56 glycosyl hydrolase families so far elucidated by these methods. Some cellulase families include endo — and exoglucanases from either fungal or bacterial systems. The mechanism of cellulase action is a major feature common to all members of a family (e. g., single displacement, leading to inversion of configuration at the anomeric carbon, or double displacement, leading to retention of configuration at the anomeric carbon (68, 69). Subsets of these glycosyl hydrolase families have been grouped into super-families, or clans, on the basis of conservation of enzyme mechanism and tertiary structure of the molecule (70, 71).

Applications and Economics of Cellulase Production. The ultimate goal of advanced cellulase development research is to develop a robust and easily integratable cellulase production system that can produce adequate amounts of highly effective enzyme to satisfy process requirements at a cost compatible with overall ethanol process economics-probably in the range of $0.05 to $0.10/gallon ethanol produced. For comparison, starch-based ethanol processes currently consume approximately $0.04 to 0.05 in enzyme per gallon of ethanol produced (Miller, C, personal communication, 1995). Current assumptions dictate that the enzyme production system must be able to produce vast quantities of enzyme. For example, at an enzyme loading of 25 FPU/g cellulose, a bioethanol process will require about 11 million FPU (19 kg, 42 lbs) of cellulase to process 1 ton of biomass (1000 lb cellulose) to 84 gallons of ethanol. This amount of enzyme is equivalent to about 143 L of a commercial preparation that contains

80.0 FPU/L. Therefore, a single 2,000 ton/day bioethanol plant would require a staggering 15,000 ton/year cellulase, which is conservatively one-fourth the entire 1994 U. S. market for all industrial enzymes (72). Put another way, at 25 FPU/g cellulose,

136.0 FPU (approximately 1.8 L of commercial enzyme) of cellulase will be required to produce 1 gallon of ethanol.

Most industrially useful enzymes are produced using fermentative processes, which involve capital — and labor-intensive production plants that use large stainless steel tanks, huge volumes of media, expensive energy, and material inputs (agitation, oxygenation, media, steam sterilization, etc.). Commercially available cellulase preparations are currently employed on a large scale almost entirely in non-biomass conversion applications, such as textile processing ("biostone"-washed jeans), detergents, and food processing. Each of these markets commands a much higher price for cellulase than can be afforded by any projected bioethanol process in the United States today, which is due, in large part, to the low cost of gasoline and the high cost of feedstock materials.

Given that cellulases are a critical component of lignocellulose conversion technology and that commercially available cellulase preparations are currently far too expensive for use in bioethanol processes, the alternatives are many and can be summarized briefly.

• Develop an on-site liquid or solid-state fermentative process for producing cellulase with optimized microbial sources (e. g., filamentous fungi, such as T. reesei, Aspergillus nigerf or Humicola insolens).

♦ Develop a genetically engineered cellulase production system that uses a bacterial or fimgal host to express and secrete effective cellulase preparation, or some other genetically engineered cellulase production system that uses a non-microbial host organism, such as insect cells, crop plants, or lactating mammalian systems.

Developing cost-effective cellulase production methods, designed for complete and efficient hydrolysis of cellulose in relevant feedstocks is therefore essential. Besides the cost that results from losses in overall ethanol yield when enzyme is produced from biomass, high cellulase production costs are also due to the intrinsic costs of fermentative processes, including capital-intensive tankage, agitation, and sterilization equipment, and costly media and contamination control chemicals.

Prospective for T. reesei Cellulase Use in Near-Term Bioethanol Plants. T. reesei mutants are generally recognized to be the best strains currently available for the industrial production of cellulases (73). Yet, the cost of bulk quantities of cellulase to the ethanol from biomass process has remained an area of uncertainty. Although few detailed economic studies are available, an estimate for the cost of cellulase production from lactose for an advanced bioethanol plant based on Iogen technology was proposed as $0.53/gallon ethanol (74). More recently, an estimate of $0.30 to $0.81 per pound cellulase protein was proposed for on-site cellulase production based on Army-Natick data (75). (These values may be cautiously converted to a cellulase cost of approximately $0.11 to $0.30/gallon ethanol assuming a specific activity of 600 FPU/g protein and that

100,0 FPU are required to produce 1 gallon ethanol). Cellulase cost data from the 1993 study by Hayn (76) proved to be in agreement with the earlier Iogen data and were based on actual separate hydrolysis and fermentation (SHF) pilot plant results, which provided a cellulase cost estimate of approximately $0.68/gallon ethanol. Because no detailed pilot — scale studies of cellulase production from pretreated woody biomass are readily available, production parameters based on small-scale T. reesei growth and induction studies are critical.

Enzyme Technology for Next-Generation Bioethanol Plants and Beyond. Because the cost of producing the enzymatic catalysts for the SSF process is a critical issue, the available enzymatic activity must be maximized. This requirement can be met by ensuring that the enzymes used are obtainable at minimal cost and have the highest specific activity and the highest possible stability at the pH and temperature of the intended application. Esterbauer and co-workers (77) caution that, "In retrospective, we and others feel that cellulase production by Trichoderma has its limitations and a significant further improvement cannot be expected. In the future, efforts should also be focused on other cellulolytic microorganisms, both bacteria and fungi."

We feel that cellulase systems capable of greater productivities and carbon conversion efficiencies than those possible from fungi are required for the success of bioethanol plants targeting objectives for the next decade (i. e., $0.67/gallon ethanol) (78,3). Preliminary technoeconomic analyses of the bioethanol process at NREL show that the cost of on-site cellulase production is keenly sensitive to delivered feedstock cost ($/ton feedstock), moderately sensitive to the carbon conversion efficiency of cellulase production [FPU produced/gram carbon consumed from feedstock = (gram cellulase/gram carbon) x (FPU/gram cellulase)], and less sensitive to enzyme loading (FPU/gram cellulose content in SSCF) (Glassner, D., personal communication, 1996). Thus, the key to this strategy is to increase the specific activity, thus deriving more FPU activity per gram of protein produced and to increase both carbon conversion efficiency and effective enzyme loading. The degree to which the specific activity of the cellulase system can be increased is not known. A related issue of equal importance is the development of an expression system that can produce large quantities of recombinant enzymes at low cost, preferably from low-value processing plant streams.

Strategies for Improving Cellulase Systems. The first major research goal for advanced cellulase technology is to increase the specific activity of cellulase systems. We propose approaching this challenge through the use of at least four distinct strategies. First, the component enzymes constituting known cellulase systems can be isolated and recombined to create new, non-natural systems to be evaluated for possible improvements in activity-to a large degree, work initiated at NREL (79) and elsewhere (80-83) has shown that significant, but probably limited, increases in system efficiency can be expected from this approach. One important discovery from two-component enzyme mixing studies at NREL was that a significant improvement in degree of synergism (DSE) and reducing sugar (RS) release (i. e., as much as 40% DSE and 30% RS release) can be found when mixing enzymes produced from diverse organisms, in this case a hot-spring bacterium and a filamentous fungus (79). Our progress in demonstrating the potential for new cellulase system engineering by assessing the efficacy of mixed origin and native binary and ternary systems is shown in Table 1. Although admittedly not as active on Sigmacell as the native T. reesei ternary system, the new ternary system based on Acidothermus cellulolyticus El, Thermomonospora fusca E3, and T. reesei CBH I is competitive and offers the potential for another important advantage for bioethanol process cellulases, thermal tolerance and increased process half-life.

Engineered Cellulase Systems at NREL. A. cellulolyticus is a thermo tolerant, cellulolytic bacterium which was originally isolated from a Yellowstone National Park hot spring (84). One component enzyme from this cellulase system, El, has been purified, characterized, crystallized, and subjected to x-ray crystallographic analysis (85). El is a 60 kD endoglucanase that is highly thermostable, demonstrates high specific activity on carboxymethylcellulose (86, 87), and is a member of the glycosyl hydrolase Family 5, Subfamily 1 (88). Because of its membership in this family, El is expected to show retention of the stereochemistry at the anomeric hydroxyl following catalysis.

Cellobiohydrolase I (CBH I), a 52 kD exoglucanase produced by the filamentous fungus, T. reesei, is the most abundant component of that organism’s cellulase system. T. reesei also produces another exoglucanase, CBH II, and at least three endoglucanases, referred to as EG I, EG П and EGV. CBH I is produced in extremely large amounts from a single genomic gene and may represent up to 60% of the protein secreted by strains of T. reeseitbax can produce extracellular protein at concentrations up to 40 grams/L (89).

Table 1

Non-Native Endoglucanase/Exoglucanase Mixtures Tested at
10% Total Saccharification for Degree of Synergistic Effect (DSE)
and Reducing Sugar (RS) Release

CBH I is not thermotolerant, acts processively from the reducing end of a cellulose substrate, demonstrates retaining-type product stereochemistry (63), is a member of glycosyl hydrolase Family 7 (90), and exhibits a synergistic activity in combination with every endoglucanase that has so far been tested, including El (79).

E3 is a nonthermotolerant exoglucanase secreted by the actinomycete T. fusca (80). This enzyme is about 60 kD in molecular weight, demonstrates an inverting-type product stereochemistry, and is a member of glycosyl hydrolase Family 6 (Wilson, D., personal communication, 1996).

We feel that when acting on pretreated biomass, engineered enzyme mixtures can be formulated that are more effective than the most active native mixtures, at least at the binary enzyme level. Based on the notion that an engineered mixture of activities is desirable, and that an effective cellulase system must have at least one highly active endoglucanase, one cellulose reducing terminus-specific exoglucanase, and one cellulose nonreducing terminus-specific exoglucanase, we have proposed the selection of the

A. cellulolyticus El, T. reesei CBH I, and T. fusca E3 as components for our basic system.

Second, it should be possible to improve the kinetic efficiency of cellulases that work on pretreated biomass by using known principles of enzyme engineering (new strategies may also be required and are proposed for future work). We have proposed, for example, a two-phase approach to this problem by first modifying the native structure of El by targeted amino acid replacement to ensure optimal enzyme-cellulose (biomass) surface interaction and by improving the catalytic efficiency of the active site, also by SDM. A similar approach will be used to improve the action of the exoglucanases on pretreated biomass, with the additional goal of improving thermal tolerance.

Third, the usefulness of "accessory" glycosyl hydrolases for the enhanced saccharification of pretreated biomass substrates should be investigated. For example, the new countercurrent pretreatment methodologies described earlier produce somewhat higher levels of cellulose hydrolysis; however, the material left insoluble is more resistant to conventional cellulase action. Also, the liquid waste streams from complete hydrolysis tend to harbor enhanced levels of xylooligodextrins, as well as other, as yet unidentified, oligosaccharides. An important example of the effectiveness of removing substituent chemical inpediments to cellulases action was shown by Kong and co-workers (97) when they demonstrated that the cellulase digestion of aspen wood was substantially accelerated by prior chemical deactylation. This result could presumably also be achieved by the action of xylan acetyl esterases (92).

The overall strategy for developing engineered cellulase systems is depicted in Figure 2. The obvious benefit of routine reassessment of the specific enzyme candidates chosen for system membership is apparent, but resource intensive. A fundamental dilemma is encountered when limited resources dictate that choices be made between attempting to improve the operational characteristics of competitive enzymes (by SDM or mutation/selection) or simply returning to new rounds of screening from known cellulase producing organisms. A secondary benefit for the enzyme engineering approach is the acquisition of fundamental knowledge concerning cellulase action and structure relationships, which should improve chances of future success.

Relevant Cellulase Assays for Bioethanol Process Applications. The activities described above will be supported by the critical determination of the effectiveness of new enzymes using a novel assay method. Although cellulase enzymes are widely sold, and

Improve by SDM using biomass

Commercial scale expression system

their industrial utilization estimated, on the basis of the FPU of activity, the traditional "filter-paper assay" is severely limited as a predictor of cellulase performance in the extensive saccharification (80%-90%-plus) of actual industrial lignocellulosic substrates. These limitations are traceable both to the chemical and physical differences between filter paper and industrial substrates, and to the nonhomogeneous nature of most cellulosic substrates (filter paper included), which means that assays run to very limited extents of conversion (such as the 4% conversion target in the filter-paper assay) measure the digestibility of only the most easily digestible fraction of the substrate, and reveal little about the convertibility of the bulk of the substrate. Actual performance of cellulases is estimated better by assays that utilize the actual application substrate, and are run to the extents of conversion required in the process.

Because of the inhibitory nature of the products of cellulase action (primarily glucose and cellobiose), such high-conversion assays encounter the problem of significant product inhibition, if run in "closed" systems as simple saccharifications (93, 94). A new saccharification assay has been devised at NREL in which a continuously buffer-swept membrane reactor is used to remove the solubilized saccharification products. The diafiltration saccharification assay (DSA) serves as a reliable predictor of the performance of combinations of cellulase and substrate under simulated SSF conditions but retains the analytically more direct and accurate nature of a saccharification reaction. This assay will be used to compare the effectiveness of commercial T. reesei and specially engineered (cloned) cellulase preparations in the saccharification of standard and novel dilute acid pretreated substrates.

For decades, cellulase biochemists have proposed that enzyme-secreting microorganisms should produce somewhat different hydrolytic enzyme systems when presented complex biomass substrates, than when grown on simple, soluble sugars. We have recently shown that a T. reesei mutant archived at NREL produces a cellulase system considerably more effective at hydrolyzing pretreated hardwood sawdust when grown in the presence of the same substrate (95).

Expression of Engineered Cellulase Genes. Our initial goal was to identify the cellulase genes that will be incorporated into the first-generation cellulase production system. This has been done (at least provisionally). The next goal is to select a host organism most likely to be able to produce a high specific activity enzyme preparation in a cost-effective manner. Initially, single-gene expression strains will be constructed to maximize expression of active gene product. Strains shown to express different target genes at desired levels will be crossed to combine two or more cellulase genes in the same strain. It is essential to maximize the specific activity (activity/g protein) of the genetically engineered cellulase preparation, which requires specific molar ratios of each particular combination of endoglucanases, exoglucanases, and p-glucosidases. Multi­gene expression strains will eventually be constructed to maximize expression of balanced cellulase expression systems.

Once a host system has been selected, the cellulase coding sequences must be incorporated into genetically engineered artificial gene constructs, which will be recognized and readily expressed in that organism. Because the cellulase genes targeted for expression in the desired host may have originated from either a bacterium or a fungus, the standard approach will be to use the coding sequence from the selected cellulase gene and place it downstream from a host-derived promoter known to express its native product at very high levels. Downstream from the coding sequence will be spliced appropriate transcription termination signals (and polyadenylation signals, if required) known to function effectively in the chosen host organism, possibly from the same gene as the promoter was derived. The exact DNA sequence context and proximity of the host promoter sequence and the foreign coding sequence is critical in any chimeric gene construction, because a single nucleotide change in sequence or distance can make a huge difference in the behavior of the gene. How changes in promoter sequence context will affect expression of chimeric genes cannot be accurately predicted for any organism at this time (except perhaps E. coli).

The cellulase coding sequences that have been selected for incorporation into the initial cellulase production system include:

• A. cellulolyticus El endoglucanase (gene of bacterial origin; 61% G+C content)

• T. reesei CBHI (cDNA clone of fungal origin; ~50% G+C content)

• T. fusca E3 (gene of bacterial origin; high G+C content)

• A. cellulolyticus p-glucosidase (gene not yet cloned)

Initial work in the area of gene cloning took place in the early 1970s using the gram­negative bacterium E. coli as the host organism. E. coli is still by far the most well — characterized organism in terms of its molecular genetics and the biochemistry of its genetic machinery. Within a short period it became possible to clone and express foreign DNA in other bacteria, such as Bacillus subtilis, Streptomyces lividans, and others, as well as brewer’s yeast (Saccharomyces cerevisiae). During the past 10 years, cloning and expression of foreign DNA in fungi has become fairly routine (Neurospora, Aspergillus, and Trichoderma, for example). The ability to introduce foreign DNA and control its expression in a wide variety of higher organisms has also been achieved only recently (including many plant species, insects, and mammalian lactation system).

The most technically approachable systems for expressing foreign DNA at high levels include bacteria (E. coli, S. lividans, B. subtilis, B. brevis, B. stearothermophilis, and B. licheniformis), yeasts (S. cerevisiae, Pichia pastor is, and P. stipidis), filamentous fungi (A. niger, A. oryzae, A. nidulans, and A. awamori), higher plants (tobacco, alfalfa, and Arabidopsis thaliana), insect cells and larvae (Baculovirus), and mammalian milk. Two of the three bacterial systems listed have already been explored at NREL for their potential to express and secrete functional foreign cellulase gene products at high levels. Although perfectly adequate for producing reagent quantities of functional cellulases, both E. coli and S. lividans are inadequate as cellulase production systems on a scale required by the bioethanol process.

(1) . Escherichia coli. E. coli is still the workhorse of modem molecular biology. As a genetic system, it is extremely well characterized. All cellulase genes obtained from various microorganisms under the sponsorship of the ethanol project during the past few years were first cloned and expressed in E. coli. Efforts to improve recombinant bacterial expression of cloned cellulases quickly gave way to other hosts, including S. lividans and

B. brevis after it became clear that heterologous products were being degraded by proteases and incorporated into insoluble inclusion bodies in E. coli, and that E. coli has a limited capacity for secreting foreign proteins into the medium (87).

Although E. coli can synthesize a foreign protein to levels of 20%-30% of total cell protein, because of limitations of cell density in batch liquid culture, this amounts to only modest volumetric yields of protein (e. g., 1-2 g/L) (96). Fed-batch and continuous culture methods can achieve significantly higher specific volumetric yields (e. g., g/L/h). We employ E. coli only as a host for cloning and expressing new cellulase genes, for constructing expression plasmids which will be used in other host systems, and for producing reagent quantities of individual cellulases.

(2) . Streptomyces lividans. Reputedly "strong" promoters isolated from various Streptomycetes were used at NREL to construct expression vectors for use in S. lividans, including tipA (a thiostrepton inducible promoter) and STI-II (S. longisporus soybean trypsin inhibitor II). Using these promoters, we have constructed various expression vectors in hopes of achieving g/L quantities of secreted, functional recombinant El in S. lividans. We have successfully secreted fully active El from most of these constructs, but none exceeds the production level of that produced by the native gene expressed in S. lividans. On the other hand, efforts to increase the expression level of a p-glucosidase cloned from Microbispora bispora successfully increased the expression level by a factor of 3-4, to more than 200 mg/L (Xiong, X., unpublished results, 1996). Additionally, Wilson reports that he has successfully increased the level of expression of one T. fusca exoglucanase gene, E3, approximately five fold using the STI-II promoter (i. e., 150-200 mg/L) (Wilson, D., personal communication, 1996). Despite these successes, it is unlikely that the S. lividans system will be capable of much more than about 1 g/L production levels in batch culture.

(3) . Saccharomyces cerevisiae. Despite being genetically well characterized, which provides the genetic engineer with numerous potential host strains, selectable markers, well-characterized promoters, and several vector alternatives, baker’s yeast is not a good choice as a host for a cellulase expression system (97). Because of its notorious capacity for hyperglycosylation of foreign proteins and its limited capability for synthesizing and secreting foreign proteins, S. cerevisiae has not been considered a serious candidate for a cellulase production system at NREL.

(4) . Pichia pastoris. P. pastoris can generate very high densities of protein-rich biomass in simple defined media, using methanol as a carbon source. For this reason, P. pastoris was first exploited commercially in a methanol-based process to produce single-cell protein (SCP) for use as a protein supplement in animal feeds. After characterization of the biochemical pathways for the metabolism of methanol in this and similar organisms, the two alcohol oxidase genes from P. pastoris were cloned and sequenced. Efficient transformation and expression systems were also developed for this organism, and are based on the methanol-inducible promoter from the alcohol oxidase 1 (AOX1) gene. The vectors used for P. pastoris transformation integrate into the yeast genome by homologous crossover at the AOX1 locus. Because P. pastoris does not heavily glycosylate secreted proteins, as is common in S. cerevisiae, it is useful in the production of human pharmaceuticals that require glycosylation for biological activity. The state of the field of heterologous gene expression in P. pastoris has been very recently reviewed (98).

Several examples show that Pichia can routinely achieve percentage yields (5%-40% of total cell protein) much higher than baker’s yeast, and often equivalent to E. coli or baculovirus (99,100). Because Pichia is able to grow to much higher densities in liquid culture than any of the aforementioned systems, it can produce much higher volumetric yields (g/L). Scale-up of Pichia culture to extremely high cell density is simple and has resulted in enormous volumetric yields (e. g., 12 g/L for tetanus toxin fragment C [101] and >3 g/L secreted human serum albumin [102]).

We have tested a strain of P. pastoris designed to express and secrete of the A. cellulolyticus El endoglucanase. In this construction, the mature El coding sequence was joined in the same translational reading frame to the yeast alpha factor signal sequence present in pPIC9. Transformants have not yet been analyzed for gene copy number, but trial fermentations have already yielded 1.5 g/L of EI. Approximately 50% of the EI produced in P. pastoris is secreted into the medium and the remainder is found intracellularly (Thomas, S., unpublished results, 1995).

(5) . Extracellular Production of Heterologous Fungal Proteins in Fungal Hosts. Several investigations concerned with the efficiency of secretion of heterologous proteins have focused on fungal enzymes. The results of published experiments to express foreign genes in filamentous fungi have recently been summarized by van den Hondel {103). Most of these enzymes are important for the industrial production of foodstuffs, animal feeds, and detergents. A large number of studies have been carried out using Aspergillus awamori, A. niger and A. oryzae, for which extensive experience in fermentation and downstream processing has been established.

The initial level of production of fungal proteins in heterologous hosts is usually in the range of 10 to 50 mg/L. Under nonoptimized conditions, similar production levels are observed for efficiently secreted homologous proteins. However, after optimization of the production process, levels of at least 3 g/L have be obtained. Clearly production yields can usually be improved considerably through the use of modified hosts, media optimization, use of appropriate large-scale fermentation conditions, and classical strain improvement procedures.

(6) . Extracellular Production of Bacterial Proteins in Fungal Hosts. The

literature reveals only a few reports of studies that deal with the expression and secretion of bacterial proteins in filamentous fungi are available to date. The Cellulomonas fimi endoglucanase, being a cellulase, is particularly relevant to the matter at hand. The 5′ region of the inducible A. nidulans ale A gene was employed to direct expression of the

C. fimi endoglucanase in an A. nidulans host strain previously modified to overproduce the alcR gene product, which positively regulates alcA transcription. This promoter is repressed in media that contains glucose, but can be induced when the carbon source is switched to ethanol. This work demonstrated production of approximately 20 mg/L of functional, secreted endoglucanase in shake flask cultures growing sub-optimally in minimal media at 37°C for 48 h {104).

Alternatively, Turnbull and co-workers {105) produced E. coli enterotoxin subunit В (LTB) at low levels (2 ng/|ig soluble protein, or 24 ig/g wet weight of mycelia) in

A. nidulans using an "up-regulated" inducible amdS promoter. This promoter can be induced by either acetate or acetamide. The fact that the рге-LTB product was properly processed to remove the bacterial signal peptide but not secreted into the medium indicates some sort of incompatibility with the fungal secretory apparatus, or that secreted protein was rapidly degraded by extracellular proteases.

(7) . Trichoderma reeseu T. reesei can produce remarkable amounts of extracellular protein (20 to 50 g/L). Even so, until now, the use of filamentous fungi as general production hosts has been restricted mostly to research and industrial laboratories with special interest in these organisms. Compared with E. coli and S. cerevisiae, which serve as model organisms for basic research and are widely used in molecular biology, the efforts so far invested in the development of filamentous fungi as production hosts have been very limited. The major cellulase, cellobiohydrolase I (CBH I), which is produced from a single copy gene, represents -50% of the total protein secreted. Thus, the cbhl promoter is extremely strong. The excellent synthesis and secretion capacity of the organism, together with established fermentation conditions, prompted development of

T. reesei as a host for production of heterologous proteins. T. reesei has the advantage of possessing a eukaryotic secretory machinery, and, most likely, similar protein modification properties (e. g., high mannose type N-glycosylation; 106) to mammalian systems.

Recently, in a continued search for powerful promoters that are active in the presence of glucose-containing media, the group at VTT (Espoo, Finland) has screened a cDNA library for sequences that are highly abundant (707,108), It has then used those cDNAs to isolate the corresponding genomic clones and the promoters for those genes. This level of expression proved to be 20 to 50-fold higher than that of the pgkl gene. Promoters for two of these genes have been isolated and used to drive expression of the homologous EG I coding sequence in T. reesei strain QM9414 growing in glucose — containing medium.

(8) . Higher Plants. There is sound reasoning behind the approach to produce bulk industrial proteins in crop plants. According to Pen and co-workers, "In terms of cost — effectiveness for producing biomass, the growing of crops in the field can generally compete with any other system. It is inexpensive, it can be done in bulk quantities, and it requires limited infrastructure. These observations suggest that the exploitation of arable crops for the production of food, feed, or processing materials would be very attractive" (109), Aside from various academic laboratories around the world, at least three plant genetic engineering companies are actively pursuing the expression of relatively low value bulk industrial proteins in crop plants of one sort or another: MOGEN International (Leiden, The Netherlands), human serum albumin (HSA) and a-amylase in tobacco and potato; Ciba-Geigy (Research Triangle Park, NC), cellulases in maize; Calgene, Inc. (Davis, CA), cellulases in tobacco.

The idea for production of industrial enzymes and other bulk proteins in plants is not unique to NREL. For example, others have already demonstrated expression of Bacillus licheniformis a-amylase in tobacco (109, 110), Clostridium thermocellum xylanase (XynZ) in tobacco (111), and HSA in potato (109). These examples involve the stable transformation of plant nuclear DNA via Agrobacterium-mediated gene transfer. In all three cases, no effect on cell growth and development was observed, indicating the foreign DNA expression did not adversely affect the transformed plants. A patent suggests that a variety of proteins from many sources can be used as target proteins for expression in barley endosperm (772).

B. licheniformis a-amylase was expressed at 0.5% of total protein in tobacco. Alpha- amylase purified from tobacco exhibited a slightly higher molecular weight than the native enzyme (64 versus 55 kDa), and was entirely due to glycosylation in the plant system. Nevertheless, despite glycosylation, the a-amylase expressed in tobacco was active, secreted, stable at 95-100°C, and otherwise completely indistinguishable from the native enzyme. Perhaps just as important is that the starch content of transformed leaves (in chloroplasts) was unaffected by the cytoplasmically expressed a-amylase. This is an important point because it clearly demonstrates the ability to isolate a transgenic protein from its potential substrate by compartmentalization, thus avoiding potentially detrimental effects on the plant.

Herbers and co-workers (111) recently expressed a truncated version of a thermostable C. thermocellum xylanase in transgenic tobacco plants. The authors speculate that these plants might be useful for producing xylanase, which has numerous applications in the paper industry and agriculture. The xylanase was synthesized as a 37 kDa polypeptide and correctly targeted to the intercellular space by means of a proteinase inhibitor П signal peptide. The xylanase was one of the most abundant proteins in total extracts (4.1 +/- 1.6%) and represented more than 50% of protein present in the intercellular fluids. The transgenic plants, grown under greenhouse conditions, were not affected by the foreign enzyme, possibly because of the high temperature optimum of the xylanase and the low levels of xylan in tobacco cell walls.

In 1993, the highest published level of expression of a foreign protein by nuclear transformation in a plant system was 1.5% of soluble protein {113). Differences in the level of foreign gene expression seem largely gene dependent and may be due to efficiencies in transcription/translation or stability of the gene products. Further improvements in expression levels seem likely, given the modest effort expended in this area so far.

Another new approach to plant transgenic expression systems involves integrating the target gene into the tobacco mosaic virus RNA genome, followed by infecting tobacco and other solanaceous species (e. g., tomato). As the infection spreads systemically, the plant becomes a dedicated bioreactor for expressing the foreign gene (114-116). TMV — based vectors can produce heterologous proteins in tobacco at levels between 5% and 40% of total cell protein (della-Cioppa, G., personal communication, 1995).

Despite progress in the development of gene expression technology, significant problems remain in the manufacture of many complex proteins. Many post-translational modifications performed by animal cells can be performed by green plants which, like animals, are complex, eukaryotic organisms. Green plants are a promising, underexploited system for expressing new proteins, including cellulases. In addition, green plants are photoautotrophic, requiring only carbon dioxide, water, nitrogen, sulfur, phosphorus, and trace amounts of other elements for growth.

Rapier (10) has determined that a mixture of 80% municipal solid waste (MSW) and 20% sewage sludge (SS) provides the optimal combination of energy and nutrients for a mixed culture of acid-forming microorganisms; therefore, this ratio was used in this study.

A series of semi-solid fermentations were operated using custom fermentors. The fermentors were horizontal, stainless-steel cylinders of 17.5-cm length and 10-cm diameter. A center shaft had finger-like projections that extended nearly to the cylinder wall. As the shaft rotated, it "kneaded" the fermentor contents through finger-like projections located on the cylinder wall. The 1.5-L fermentor was filled with about 0.5 L of fermenting MSW and SS.

determined as the weight loss from 2-g biomass samples placed in fine-mesh nylon bags located in the rumen of a fistulated steer.

Treatment conditions: lime loading = 0.1 g Ca(OHyg dry biomass, temperature = 100°C, water loading = 9 g HjO/g dry biomass, time = 1 h (bagasse) or 2 h (African millet straw, sorghum straw, tobacco stalks).

Two or four fermentors were operated in series with solids flowing countercurrently to the liquid. Solid/liquid separation was achieved by centrifuging the fermentor contents and decanting the liquid in an anaerobic hood. This countercurrent operation allows high VFA concentrations to be generated in the fermentor receiving fresh, highly reactive solids. Because inhibition is low, it also allows high conversion in the fermentor receiving fresh liquid.

Table ІП shows the results from four countercurrent fermentations. Compared to rumen fermentations which typically require only a couple of days, the fermentor residence times are significantly longer due to the inhibition from the high VFA concentration (20 — 30 g/L versus 8-10 g/L). Fortunately, because the fermentors can be very inexpensive, the long residence time does not impose a severe economic penalty. The process time scales are similar to those for composting; thus, the process may be viewed as an anaerobic composting operation. The fermentor volume is proportional to the liquid residence time whereas the conversion is proportional to the solids residence time.

Fermentor A used two countercurrent stages and Fermentor В used four countercurrent stages. Similar VFA concentrations, conversions, yields, and selectivities were obtained; however, Fermentor A required 55% longer liquid residence time. This comparison shows the beneficial effect of increasing the number countercurrent stages.

Fermentor C also employed four countercurrent stages. Compared to Fermentor B, the ratio of solid:liquid residence times increased allowing the conversion to increase while holding the product concentration constant.

Fermentor D used essentially the same liquid and solid residence times as Fermentor B, but increased the nutrient content allowing both the VFA concentration and conversion to increase substantially. The conversion of Fermentor D represents 84% of the maximum possible. (Considering ash and lignin content, the maximum digestibility of the MSW/SS mixture is 77.5%.) Further research is required to determine the optimal nutrient package.

R. R. Gokarn, M. A. Eiteman, and J. Sridhar

Department of Biological and Agricultural Engineering,
Driftmier Engineering Center, University of Georgia, Athens, GA 30602

Numerous anaerobic microorganisms synthesize succinic acid as a fermentation product. This chapter reviews the literature for succinate producing organisms and compares the growth and succinate production of two widely differing anaerobic bacteria, Fibrobacter succinogenes and Clostridium coccoides. F. succinogenes degrades simple sugars such as glucose as well as cellulosic materials such as pulped shredded office paper. The principal products after 90 hours from 10 g/L pulped paper are succinate (3.2 g/L) and acetate (0.58 g/L), with lower concentrations of formate (0.070 g/L). C. coccoides degrades simple sugars only, and after 24 hours the principal products from 5 g/L glucose are acetate (3.0 g/L), succinate (0.57 g/L) and lactate (0.58 g/L).

Succinic acid is a four-carbon aliphatic dicarboxylic acid having pKat = 4.2 and pKaj = 5.6. The depronated form succinate can be produced by many anaerobic microorganisms at their operating conditions, usually near neutral pH. Succinic acid can be used to manufacture specialty chemicals including tetrahydrofuran, 1,4- butanediol, maleic anhydride, adipic acid, and dimethyl succinate. Its derivatives are used in the food, pharmaceutical, cosmetics and polymer industries. Anaerobic processes from renewable resources are particularly appealing for the synthesis of succinate becuase of their high yields and straght-forward scale-up requirements. This chapter reviews the literature on anaerobic processes for succinate production as well as comparing the succinate production by two widely different microorganisms.

Rumen Bacteria

The rumen is a highly competitive microbial ecosystem of primarily anaerobic bacteria, fungi and protozoa. These microorganisms ferment cellulose, starch and various other carbohydrates into numerous low molecular weight products. The

© 1997 American Chemical Society

principal acid products from rumen fermentations are acetate, propionate and butyrate. The production of propionate in the rumen involves cross feeding between succinate — producing microorganisms and species that decarboxylate succinate to propionate and carbon dioxide (1,2). Therefore, even though succinate itself is not a product of the entire rumen ecosystem, numerous anaerobes have been isolated which synthesize succinate as a primary end product.

Fibrobacter succinogenes. Fibrobacter succinogenes (previously named Bacteroides succinogenes) is the predominant cellulolytic bacterial species found in the rumen. Hungate (3) isolated these organisms from bovine rumen and characterized them as obligately anaerobic, gram negative, cellulolytic, non-motile, non-sporeforming, rod­shaped bacteria whose morphology may change to lemon-shaped when cultivated in the laboratory. Strains of this organism have also been isolated from mice caeca (4), pig caeca (5), gut of horses (6), mice (7), langur monkeys (8) and several African ruminants (9). A more recent study (10) comparing the 16S ribosomal ribonucleic acid sequence demonstrated that this organism differs from other Bacteroides species, and hence the organism was renamed Fibrobacter.

Most F. succinogenes strains utilize glucose, cellobiose, maltose, dextrins, lactose, pectin or cellulose, while some strains also use starch as a carbon source (3). F. succinogenes requires one branched (i. e., isobutyrate or a-methyl butyrate) and one linear (i. e., valeric, caproic, heptaonic or caprylic) volatile fatty acid for the synthesis of long chain fatty acids and aldehydes incorporated into phospholipids (11,12). F. succinogenes has an absolute requirement for biotin (13) and for ions such as Na+, K+, Ca2+, Mg2+ and P043* (2). F. succinogenes synthesizes almost all of its cellular nitrogenous compounds from exogenous ammonia even when large amounts of amino acids and nucleotide precursors are present in the media. Although glutamine or asparagine may substitute for ammonia, ammonia is preferred when multiple nitrogen sources are present (14). Like all rumen bacteria which produce succinate as a major end product (15,16), F. succinogenes has an absolute requirement for carbon dioxide. Initiation of the growth of F. succinogenes is achieved at 0.02 to 0.05% of carbon dioxide, while optimal growth is observed when the carbon dioxide concentration is above 0.1% (15). F. succinogenes also fixes carbon dioxide during succinate production, with carbon dioxide incorporated in the carboxyl group of succinate (16).

F. succinogenes can degrade highly structured, crystalline cellulose such as cotton fibers (17). The cellulose degrading enzymes and mode of cellulose degradation by this organism have been extensively studied (18-20). In order to carry out cellulose degradation, cells must have intimate contact with cellulose fibers, since the cellulase enzyme endo-P-l,4-glucanase is membrane bound (18). This requirement for contact makes the available gross surface area of the substrate a major determinant factor of hydrolytic rate (21). Endoglucanase activity is about seven times greater when the organisms are grown on cellulose than when grown on cellobiose or glucose, suggesting that the enzyme system may be regulated by a catabolite repression mechanism (18). However, Hiltner and Dehority (22) found that the presence of glucose or cellobiose does affect cellulose digestion when pH is controlled, an observation which seems to contradict the catabolic repression hypothesis. In addition to cellulose degradation, F. succinogenes also degrades hemicellulose (18,23). However, the organism cannot utilize as a substrate for growth the pentoses which are released during hemicellulose degradation (24,25). The inability of F. succinogenes to utilize pentoses is attributed to the lack of key enzymes such as xylose permease, xylose isomerase and xylulokinase (26).

F. succinogenes has restricted ranges of redox potential and pH. The redox potential range for cell viability is -290 to +175 mV, and the most prevalent morphology at highest redox potential is greatly elongated cells (27). F. succinogenes has a pH range for growth of 6.1 to 6.9 (28), and cell wash out occurs at a pH of 6.0 when the organisms are grown on cellobiose in a chemostat (29). The highest cell yield on cellobiose and cellulose occurs at the lower pH limit (28,29). The inability of the organisms to grow at lower pH may be due to inhibition of the glucose transport system or due to low substrate affinity (28,30). When grown on microcrystalline cellulose, F. succinogenes has a maximum specific growth rate of 0.076 h’1 and a maintenance requirement of 0.04-0.06 g cellulose/g cells (28), while on cellobiose or cellodextrins F. succinogenes has a maximum specific growth rate of about 0.44 to 0.48 h*1 (31).

Figure 1 summaries the biochemical pathway for succinate production by F. succinogenes (34). The degradation products of cellulose, glucose and cellobiose, are transported into the cell by a highly specific active transport system. The glucose transport system is energized by a proton gradient, while the cellobiose transport system is energized by a sodium ion gradient (32). F. succinogenes possesses fructose 1,6-biphosphate aldolase (33) and glyceraldehyde-3-phosphate dehydrogenase (34). Oxaloacetate formation from phosphoenolpyruvate is accompanied by carbon dioxide fixation and is catalyzed by GDP-dependent phosphoenolpyruvate- carboxykinase. Reduction of oxaloacetate results in malate formation with NADPH or NADH acting as the electron donor. Even though fumarase activity has not been demonstrated, Miller (34) proposed that conversion of malate to fumarate is catalyzed by fumarase. Fumarate is then reduced to succinate by a flavin-dependent membrane — bound fumarate reductase (34). During the fumarate reduction, cytochrome b acts as an electron carrier, and the step may result in ATP generation via electron transport linked phosphorylation. This hypothesis is supported by observations of higher growth yields than can be explained solely by substrate level phosphorylation (27), and of decreased growth rates in the presence of electron uncouplers (35). Conversion of pyruvate to acetyl-CoA is accompanied by the reduction of FMN with carbon dioxide evolution. The formation of acetyl phosphate from acetyl-CoA is catalyzed by phosphotransacetylase, and acetate production from acetyl phosphate yields ATP. Studies with partially isolated phosphoenolpyruvate-carboxykinase indicate that this enzyme is active only in presence of bicarbonate, GDP and the Mn2+ ion (34).

Ruminococcus flavefaciens. Sijpesteijn described Ruminococcus flavefaciens as a gram positive, non-motile, anaerobic, cellulolytic, streptococci of 0.8-0.9 pm diameter (36). The cells can exist singly, in pairs or may form a chain. R. flavefaciens ferments xylans, cellobiose or cellulose, while the fermentations of glucose, xylose or other simple carbohydrates is restricted to only a few strains (36,37). R. flavefaciens is active on amorphous cellulose (17) and also breaks down hemicellulose, but most strains cannot utilize pentoses as an energy source (38). A distinct feature of R. flavefaciens is the production of a yellow pigment when grown on cellulose. The temperature range for growth is 30 to 45°C, with 39°C being the optimum. R.

Figure 1 Biochemical pathway for succinate production by F. succinogenes.

flavefaciens produces reducing sugars when grown in excess cellulose, although usual fermentation products include succinate, acetate and at least trace formate and lactate (36).

R. flavefaciens has a requirement for branched fatty acids, which are incorporated into lipids and amino acids (39). R. flavefaciens can utilize ammonia as a sole nitrogen source (14). Biotin is required by all strains, while vitamin B12 is required by some strains. p-Aminobenzoic acid has been shown to stimulate growth (13). R. flavefaciens has an absolute requirement for carbon dioxide: 0.05 to 0.1% carbon dioxide for growth initiation and above 0.1% for optimal growth (15).

R. flavefaciens possesses enzymes which are active on cellulose, hemicellulose and pectin. Most of these enzymes are believed to be cell wall associated, since cell attachment to cellulose fibers is necessary for plant wall degradation. R. flavefaciens has active exo-l,4-P-glycosidase enzymes which generate cellobiose and cellotriose from cellulose and xylobiose and xylotriose from xylan. R. flavefaciens also exhibits low levels of aryl p-glucosidase and aryl p-xylosidase activity (40). Presence of soluble sugars such as cellobiose seems not to affect cellulose digestion, although cellulose digestion may be influenced by a pH decrease when soluble sugars are rapidly fermented (22). R. flavefaciens has a poor affinity for cellobiose which may result in its poor utilization. With cellobiose in a continuous culture, cell wash out occurs at a pH of about 6.1, and cell yield decreases abruptly (29). An increase in growth rate results in a shift toward more acetate and formate with less succinate. On cellulose the organism has a low maintenance requirement of 0.07 g cellulose/g cell per hour (41).

For one R. flavefaciens strain isolated from sheep, succinate is the major product of glucose fermentation in the presence of carbon dioxide, but in the absence of carbon dioxide the fermentation shifts to a homolactic pattern (42). The pathway for succinate and acetate production by R. flavefaciens is believed to be identical to that of F. succinogenes shown in Figure 1 (42-44). Succinate formation is accompanied by fixation of carbon dioxide, which is incorporated in the carboxyl group. Formate can form from free carbon dioxide or from pyruvate (43). PEP — carboxykinase requires GDP and the bicarbonate ion as cosubstrates, and this enzyme is most effective in converting phosphoenolpyruvate to oxaloacetate in the presence of Mn2+ (44).

Ruminobacter amylophilus. Hamlin and Hungate (45) first isolated Ruminobacter amylophilus (Bacteroides amylophilus) from bovine rumen and characterized the species as an obligately anaerobic, gram negative, non-sporeforming, non-motile bacterium (45). A more recent study of the 16S ribosomal ribonucleic acid sequence showed that this organism differs from other Bacteroides species and hence was renamed Ruminobacter (46). R. amylophilus cells are rod-shaped, 0.9-1.6 pm, but may also exhibit larger irregular shapes. The organism has subsequently been isolated by Blackman and Hobson (47), Bryant and Hobson (48), Caldwell et al. (16), and Blackman (49). R. amylophilus uses only starch and maltose as substrates, and the organism’s population in the rumen is increased when the animal is fed a high starch diet (36). Fermentation products include acetate, succinate, formate and trace ethanol and lactate (45). The organisms grow at a pH of 6.5-7.8 and temperature range of 35 to 45°C (45). R. amylophilus growth occurs within a wide redox potential range of -320 mV to +250 mV, although the specific growth rate decreases above 0 mV (27). Above +200 mV more lactate is produced at the expense of succinate (27).

R. amylophilus has an absolute requirement for Na+, and this requirement cannot be replaced by K+, Li+, Cs+ or Rb+ (50). In addition to the Na+ ion, R. amylophilus requires K+, P043 and trace Mg2+ (50). The Na+ and K+ ions affect the growth rate and growth yield, while P043’ affects only the growth yield (50). R. amylophilus also has an absolute requirement of carbon dioxide for growth; however, bicarbonate is a suitable substitute (16). Growth is initiated at a carbon dioxide concentration between 4.5 x 10*3 M and 9 x 10’3 M, while optimal growth is achieved at 1.2 x 10‘3 M (16).

R. amylophilus possesses starch-degrading enzymes such as amylase and amylopectinase. The attachment between the cell and starch molecule is mediated by a protein or protein complex (51). R. amylophilus possesses enzymes of the Embden — Meyerhof-Pamas pathway (52). The presence of fumarate reductase enzyme suggests that succinate is produced by the reduction of fumarate (16). The production of succinate involves fixation of carbon dioxide which is incorporated as the carboxyl group (16). The pathway for succinate production is thought to be identical as the pathway for F. succinogenes shown in Figure 1.

Succinimonas amylolytica. Succinimonas amylolytica was isolated from bovine rumen (53). The organism is a gram negative, motile, anaerobic, non-sporeforming, short, rounded to coccoid bacterium 1.0 to 1.5 pm by 1.2 to 3 pm. S. amylolytica grows in a temperature range of 30 to 37°C and utilize glucose, maltose, starch or dextrin as a substrate. Fermentation products include succinate, acetate and trace propionate. The organisms grow well in media containing trypticase and yeast extract, and do not grow in the absence of either bicarbonate or carbon dioxide (53). The concentration of carbon dioxide required for the initiation of growth and optimal growth is at least 0.1% (15). S. amylolytica also requires acetate and other volatile fatty acids or casein hydrolysate (48). Even though S. amylolytica is normally associated with starch digestion in the rumen, the species possesses a wide range of glycoside hydrolases which aid in the utilization of plant cell wall degradation products (54).

Succinivibrio dextrinosolvens. Bryant and Small (55) first isolated Succinivibrio dextrinosolvens from a bovine rumen and described the species as an anaerobic, non­sporeforming, gram negative, mobile, helicoidal rod-shaped bacterium, 0.3 to 0.5 pm by 1 to 5 pm. The organisms can metabolize glucose, fructose, L-arabinose, D — xylose, galactose, maltose, sucrose, dextrins or pectins (55). S. dextrinosolvens cannot degrade cellulose, hemicellulose or starch, although the organism possesses a wide range of monosaccharide-generating glycoside hydrolases (54,55). Glucose fermentation yields principally acetate and succinate and is accompanied by significant carbon dioxide uptake (55). Formate is a minor product, while some strains also produce trace lactate (55). Similar organisms isolated from an ovine rumen by Wilson (56) were later also termed S. dextrinosolvens strains by Bryant (36).

S. dextrinosolvens has an absolute requirement for naphthoquinone, menadione or vitamin K5, with naphthoquinone resulting in best growth (57). S. dextrinosolvens possesses several nitrogen-assimilating enzymes such as urease, glutamate dehydrogenase and glutamine synthetase. Under ammonia limiting conditions the organism uses the ATP-driven glutamine synthetase system, while under excess ammonia the glutamate dehydrogenase enzyme is utilized (58). In addition to ammonia, S. dextrinosolvens requires an exogenous supply of amino acids to satisfy nitrogen requirements (48). In the absence of carbon dioxide, limited growth occurs after an extended lag phase (15). For optimal growth, a carbon dioxide concentration above 0.1% is required (55). S. dextrinosolvens requires volatile fatty acids and Na+ ion (which cannot be replaced by K+, Li+, Cs+ or Rb+). The Na+ concentration affects both the growth rate and the growth yield of S. dextrinosolvens (59).

The catabolic end products of glucose fermentation by S. dextrinosolvens are affected by the growth rate. Increased growth rate results in decreased succinate and acetate production and increased lactate formation (60). The pathway of succinate production appears to be identical to other rumen organisms already described (see Figure 1) with fixed carbon dioxide incorporated into the carboxyl group of succinate. S. dextrinosolvens has been shown to produce formate from free carbon dioxide present in the media (60).

Prevotella ruminocola. Bryant et al. (53) first isolated Prevotella ruminicola (previously named Bacteroides ruminicola) from bovine rumen and characterized the species as gram negative, non-motile, rod-shaped 0.8-1 pm by 0.8-3 pm, with slightly tapered, rounded ends (53). The organism recently was renamed Prevotella based on its genetic material (61). Several subspecies of P. ruminicola may be distinguished on the bases of morphology, substrates fermented and nutrient requirement. Most strains belonging to P. ruminicola subsp. ruminicola can utilize xylose, glucose and maltose. Some strains have the ability to hydrolyze starch and to utilize arabinose, sucrose and dextrins. Xylans and pectins are rapidly fermented by this particular subspecies (53). P. ruminicola subsp. ruminicola lacks the enzyme superoxide dismutase which is present in the subspecies brevis (62). Cells of P. ruminicola subsp. brevis are coccoid-to-oval shaped and do not require hemin (48,53). Most strains of this subspeices cannot utilize xylose and xylans, but can use pentoses as carbon sources. They also have the ability to hydrolyze starch and utilize maltose and sucrose (53).

P. ruminicola requires volatile fatty acids and acetate for growth. Use of casein hydrolysate can stimulate the growth of these organisms (48). As all other succinate producing rumen bacteria, P. ruminicola requires carbon dioxide for growth (63). The initiation of growth is achieved at 0.02-0.05% carbon dioxide, while above 0.1% optimal growth occurs (15). P. ruminicola exhibits proteolytic activity and possesses at least three different active proteinases (64).

P. ruminicola produces acetate, succinate and formate as products of sugar fermentation (53). In the presence of vitamin B12, strain 23 of P. ruminicola can also produce propionate (65). In this case the formation of propionate occurs via the direct reductive pathway (acrylate pathway) (66). P. ruminicola has low affinity for maltose, sucrose and cellobiose in comparison to glucose (67). The maintenance coefficient for P. ruminicola when grown on glucose is 0.135 g glucose /g cells • h (68). Glucose toxicity is observed with P. ruminicola strain B,4 (69). A pH below 5.7 halts the growth of P. ruminicola; however, growth is not significantly affected by pH changes above 5.7 (70).

P. ruminicola B{4 grows rapidly in a batch culture with a doubling time of 1.65 h (63). The conversion of glucose to phosphoenolpyruvate occurs via the Embden-Meyerhof pathway. The routes of synthesis for succinate and acetate are identical to previously described organisms (see Figure 1), with carbon dioxide again incorporated into the carboxyl group of succinate (63, 71).

Wolinella succinogenes. Wolinella succinogenes (previous named as Vibrio

succinogenes) was isolated by Wolin et al. (72) from bovine rumen and characterized as a curved rod-shaped, motile, anaerobic bacterium approximately 0.6 by 0.3 pm in size (72). A more recent study of the ribosomal ribonucleic acid content and evidence of the organism’s inability to ferment sugars resulted in their renaming to Wolinella succinogenes (73). W. succinogenes contains cytochromes which impart pink color to the cells. These organisms conserve energy by oxidation-reduction reaction in which hydrogen or formate acts as an electron donor and fumarate, malate, asparagine, nitrate, elemental sulfur or nitrous oxide acts as an electron acceptor (72,74,75). These oxidation-reduction reactions can be represented as (72, 75):

formate + H+ + fumarate —► C02 + succinate H2S + fumarate -► succinate + S formate + H+ + S -► C02 + H2S

Major fermentation products are carbon dioxide and succinate when these organism grow on fumarate and formate (72). Nitrous oxide can be reduced to nitrogen, nitrate to nitrite or ammonia, and elemental sulfur to hydrogen sulfide (74, 75).

W. succinogenes can use oxygen as an electron acceptor, but only at a low partial pressure of oxygen (72). At such low oxygen concentrations, hydrogen peroxide generated is degraded by peroxidase, but at high oxygen concentrations, hydrogen peroxide is hypothesized to accumulate and inhibit growth (76). W. succinogenes has a requirement for some succinate when grown on formate and nitrate. However, the presence of succinate does not appear to be necessary when the organisms are grown on fumarate, which is itself reduced to succinate (77).

The reduction of fumarate to succinate is associated with generation of ATP via electron transport phosphorylation (78). W. succinogenes has a doubling time of 3.2 h when grown on formate and fumarate, with growth yield of 4.8 g dry cell/mol formate (79). When grown on hydrogen sulfide and fumarate, the doubling time is 3.8 h and 6.0 g dry cell/mol fumarate is achieved. Use of formate and elemental sulfur results in a 1.2 h doubling time and a growth yield of 3.5 g dry cell/mol formate (75).

W. succinogenes can synthesize its cellular components from fumarate. Pyruvate acts as an intermediate for carbohydrates, nucleotides, phospholipids and for most of the amino acids. Glutamate is derived from a-ketoglutarate and further acts as a intermediate for the synthesis of amino acids belonging to its family (79).