Patent Application
20160002645
Petroleum-based fuels and commodities are commonplace, and their widespread use is growing despite evidence that the earth's petroleum resources are dwindling (Kerr (2008) Science 322: 1178-1179). It is therefore desirable to find renewable sources of carbon that can be used as an alternative to petroleum. Lignocellulosic biomass is an obvious choice since it constitutes more than half of the organic carbon in the biosphere (Boerjan et al. (2003) Ann. Rev. Plant Biol., 54: 519-546; Reddy and Yang (2005) Trends in Bbiotechnology 23: 22-27; Werpy et al. (2004) Top Value Added Chemicals From Biomass. Volume 1-Results of Screening for Potential Candidates From Sugars and Synthesis Gas. DTIC Institution.). A major obstacle to its cost effective commercialization, however, is its recalcitrance to hydrolysis into fermentable sugars (primarily glucose and xylose) (Lynd et al. (2005) Curr. Opin. Biotechnol. 16: 577-583; Mielenz (2001) Curr. Opin. Microbiol. 4: 324-329). Many currently used industrial methods degrade lignocellulose using a two-step process in which it is thermochemically pretreated and then hydrolyzed using enzymes produced by Trichoderma reesei. While high yields can be obtained using this approach, it can be costly and inefficient (Wilson (2009) Curr. Opin. Biotechnol. 20: 295-299; Himmel et al. (2007) Science 315: 804-807; Hendriks and Zeeman (2009) Bioresource Technol., 100: 10-18; Zhao et al. (2009) Appl. Microbiol. Biotechnol. 82: 815-827; Yeoman et al. (2010) Adv. Appl. Microbiol. 70: 1-55; Miller and Blum (2010) Environmental Technol., 31: 1005-1015). The creation of recombinant microbes that can degrade biomass efficiently is an attractive alternative to currently used methods. It is also an essential step towards the creation of a consolidated bioprocessor (CBP), a single microbe that has the capacity to convert lignocellulose into valuable end products such as ethanol (Olson et al. (2012) Curr. Opin. Biotechnol., 23: 396-405; Zhang and Zhang (2010) Engineering in Life Sci., 10: 398-406; Liao et al. (2011) Biotechnol. J. 6: 1409-1418; Lynd et al. (2008) Nature Biotechnol., 26: 169-172; la Grange et al. (2010) Appl. Microbiol. Biotechnol., 87: 1195-1208).
Lignocellulose consists of cellulose and hemicellulose polymers that are surrounded by lignin (Harris and DeBolt (2010) Plant Biotechnol. J. 8: 244-262; Carroll and Somerville (2009) Ann. Rev. Plant Biol., 60: 165-182). Cellulose is a homopolymer of beta-1,4 linked glucose monomers, which hydrogen bond with similar polymers to form both crystalline and amorphous regions. The crystalline regions are in part degraded by exoglucanases, which act on either the reducing or non-reducing ends of the cellulose polymer (Ghose (1977) Adv. Biochem. Engineer., Vol. 6. Springer Berlin/Heidelberg). The amorphous regions within cellulose are less ordered and are accessible to endoglucanases that cleave internal beta-1,4-glucosidic bonds. Endoglucanases also cleave chains within the crystalline region, but at a much slower rate (Id.). Hemicellulose, on the other hand, is a heteropolymer with relatively high xylan content (Pauly and Keegstra (2010) Curr. Opin. Plant Biol., 13: 305-312). It has an amorphous structure that can be easily hydrolyzed by acid or base, but enzymatic degradation requires several hemicellulase enzymes, including exoxylanases and endoxylanases (Banerjee et al. (2010) Biotechnol. Bioengineer., 106: 707-720; McCann and Carpita (2008) Curr. Opin. Plant Biol., 11: 314-320). Lignin also contributes substantially to the hydrolytic recalcitrance of lignocellulose as this extremely complex polymer consists of many types of monomers connected by a diverse array of covalent linkages (Boerjan et al. (2003) Ann. Rev. Plant Biol., 54: 519-546; Wardrop (1969) Australian J. Botany 17: 229-240).
Despite its complexity, several naturally occurring microorganisms have evolved the capacity to efficiently break down lignocellulose and use it as a nutrient (Ransom-Jones et al. (2012) Microbial Ecol., 63: 267-281; Wilson (2011) Curr. Opin. Microbiol., 14: 259-263). Significantly, anaerobic and aerobic microorganisms use different strategies to degrade lignocellulose. Aerobic fungi secrete enzymes with different cellulolytic activities, whereas anaerobic bacteria incorporate cellulases into a cell-surface displayed super-structure known as a cellulosome (Miller and Blum (2010) Environmental Technol., 31: 1005-1015; Doi and Kosugi (2004) Microbiology 2: 541-551; Doi (2008) Ann. New York Acad. Sci., 1125: 267-279; Bayer et al. (2004) Ann. Rev. Microbiol., 58: 521-554; Ding et al. (2008) Curr. Opin. Biotechnol., 19: 218-227). Although their architectures vary, cellulosomes from different microbes consist of a backbone scaffoldin protein that contains several cohesin modules capable of non-covalently binding in a 1:1 ratio to the dockerin modules of the cellulase enzymes. By clustering the cellulases into a cellulosome, the microbe is able to increase the effective enzyme concentration near its cell surface and to combine many enzymes with different activities into a single complex, enabling them to function synergistically (Bayer et al. (1998) Curr. Opin. Structural Biol., 8: 548-557). Although these organisms have potent cellulolytic activity, most are unattractive candidates for use as a CBP as they are difficult to genetically manipulate or cultivate.
Towards the goal of creating a robust CBP microbe, two model microorganisms (Bacillus subtilis and Saccharomyces cerevisiae) have been engineered to display small artificial cellulosomes (i.e., minicellulosomes) (Lilly et al. (2009) FEMS Yeast Res., 9: 1236-1249; Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858; Steen et al. (2010) Nature 463: 559-562; You et al. (2012) Appl. Environ. Microbiol., 78: 1437-1444; Tsai et al. (2009) Appl. Environ. Microbiol., 75: 6087-6093; Fan et al. (2012) Proc. Natl. Acad. Sci. USA, 109: 13260-13265). In most of these systems, a miniscaffoldin containing one or more cohesin modules is covalently or non-covalently attached to the cell surface. The minicellulosome is then often assembled ex vivo by adding purified cellulase enzymes that are fused to dockerin modules. While these recombinant microorganisms are able to degrade amorphous purified cellulose (e.g., regenerated amorphous cellulose (RAC), phosphoric acid swollen cellulose) or soluble cellulose (e.g., carboxymethyl cellulose (CMC)), their ability to degrade industrially relevant forms of biomass such as corn stover, switchgrass, and straw has not been demonstrated. Moreover, the requirement for ex vivo assembly of their cellulosomes can make some of these microbes impractical for use as an industrial CBP.
In certain embodiments recombinant modified microorganisms (e.g., Gram-positive bacteria, etc.) are provided that display on their surface a minicellulosome comprising two or more cellulolytic enzymes where the minicellulosome is self-assembled by the microorganism and resulting microorganism is capable of growing on untreated biomass (e.g. biomass that is not acid treated and/or enzymatically pre-digested). In certain embodiments the microorganism grows on lignocellulose as the sole carbon source.
In one illustrative embodiment a recombinant modified Gram-positive bacterium is provided that displays on its surface a minicellulosome comprising two or more cellulolytic enzymes, where the bacterium comprises: a protein encoding two or more cohesin domains wherein said protein is covalently linked to the surface of said microorganism, and wherein each of said cohesin domains is docked to a dockerin attached to a cellulolytic enzyme; and the one or more constructs that encode dockerin(s) attached to said cellulolytic enzyme(s); and the minicellulosome is self-assembled by said bacterium.
In various aspects, the invention(s) contemplated herein may include, but need not be limited to, any one or more of the following embodiments:
A recombinant modified Gram-positive bacterium that displays on its surface a minicellulosome including two or more cellulolytic enzymes, wherein said bacterium includes: a protein encoding two or more cohesin domains wherein said protein is covalently linked to the surface of said microorganism, and wherein each of said cohesin domains is docked to a dockerin attached to a cellulolytic enzyme; and said bacterium includes one or more constructs that encode said dockerin(s) attached to said cellulolytic enzyme(s); and wherein said minicellulosome is self-assembled by said bacterium.
The bacterium of embodiment 1, wherein said bacterium grows on untreated biomass.
The bacterium according to any one of embodiments 1-2, wherein said bacterium grows on lignocellulose as the sole carbon source.
The bacterium according to any one of embodiments 1-3, wherein said minicellulosome includes at least three cellulolytic enzymes and all of said enzymes are encoded by said bacterium.
The bacterium according to any one of embodiments 1-4, wherein said protein encoding two or more cohesin domains includes a secretory signal sequence at the N-terminus and a cell wall sorting signal (CWSS) at the carboxyl terminus.
The bacterium of embodiment 5, wherein said cell wall sorting signal includes an LPXTG motif.
The bacterium of embodiment 5, wherein said cell wall sorting signal includes a sequence shown in Table 1.
The bacterium of embodiment 5, wherein said cell wall sorting signal includes a CWSS from Staphylococcus aureus fibronectin binding protein B.
The bacterium according to any one of embodiments 5-8, wherein said secretory signal sequence includes a B. subtilis phrC secretory signal or homologues thereof.
The bacterium of embodiment 9, wherein said secretory signal sequence includes a secretion signal derived from B. subtilis phrC.
The bacterium according to any one of embodiments 1-10, wherein said protein encoding two or more cohesin domains encodes a carbohydrate binding module (CBM).
The bacterium of embodiment 11, wherein said carbohydrate binding module is a family 3 carbohydrate binding module.
The bacterium according to any one of embodiments 1-12, wherein said two or more cohesin domains are type I cohesin modules.
The bacterium according to any one of embodiments 1-13, wherein said two or more cohesin domains are cohesin domains from different microorganisms.
The bacterium according to any one of embodiments 1-14, wherein said two or more cohesin domains includes a cohesin domain from an organism selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus.
The bacterium according to any one of embodiments 1-15, wherein said two or more cohesin domains comprise cohesin domains from two organisms selected from the group consisting of C. thermocellum (t), C. cellulolyticum (c) and R. flavefaciens (f).
The bacterium according to any one of embodiments 1-16, wherein said two or more cohesin domains comprise cohesin domains from C. thermocellum (t), C. cellulolyticum (c) and R. flavefaciens (f).
The bacterium according to any one of embodiments 1-15, wherein said dockerins comprise one or more dockerin domains from organism(s) selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus.
The bacterium according to any one of embodiments 1-18, wherein said dockerins dockerin domains from two organisms selected from the group consisting of C. thermocellum (t), C. cellulolyticum (c) and R. flavefaciens (f).
The bacterium according to any one of embodiments 1-19, wherein said dockerins comprise dockerins from C. thermocellum (t), C. cellulolyticum (c) and R. flavefaciens (f).
The bacterium according to any one of embodiments 1-20, wherein said cellulolytic enzyme(s) on dormant bacteria are stable for at least 1 day, more preferably for at least 2 days, and most preferably at least 3 days.
The bacterium according to any one of embodiments 1-21, wherein said cellulolytic enzyme(s) comprise one or more enzymes selected from the group consisting of an endocellulase, an exocellulase, a beta-glucosidase (cellobiase), an oxidative cellulase, a xylanase, a hemicellulase, a lichenase, a chitenase, and a cellulose phosphorylase.
The bacterium according to any one of embodiments 1-22, wherein said minicellulosome includes at least two different cellulolytic enzymes.
The bacterium according to any one of embodiments 1-23, wherein said minicellulosome includes at least three different cellulolytic enzymes.
The bacterium according to any one of embodiments 1-24, wherein said minicellulosome includes at least one endoglucanase.
The bacterium according to any one of embodiments 1-25, wherein said minicellulsome includes at least one exoglucanase.
The bacterium according to any one of embodiments 1-26, wherein said minicellulsome includes at least two endoglucanases and at least one exoglucanase.
The bacterium according to any one of embodiments 1-27, wherein said minicellulosome includes Clostridium cellulolyticum endoglucanase Cel5A.
The bacterium according to any one of embodiments 1-28, wherein said minicellulosome includes C. cellulolyticum endoglucanase Cel48F.
The bacterium according to any one of embodiments 1-29, wherein said minicellulosome includes C. cellulolyticum exoglucanase Cel9E 31:
The bacterium according to any one of embodiments 1-30, wherein said Gram-positive bacterium includes a Gram-positive bacterium that encodes a sortase.
The bacterium of embodiment 31, wherein said Gram-positive bacterium includes a Gram-positive bacillus.
The bacterium of embodiment 32, wherein said Gram-positive bacterium includes a genus selected from the group consisting of Corynebacterium, Clostridium, Listeria, and Bacillus.
The bacterium of embodiment 33, wherein said bacterium is a Clostridium acetobutylicum.
The bacterium of embodiment 33, wherein said Gram-positive bacterium comprise is B. subtilis.
The bacterium of embodiment 31, wherein said Gram-positive bacterium includes a thermophilic Geobacillus spp.
The bacterium of embodiment 31, wherein said Gram-positive bacterium includes a Gram-positive coccus.
The bacterium of embodiment 37, wherein said bacterium is selected from the group consisting of S. aureus, S. epidermis, and S. saprophyticus.
A recombinant modified Gram-positive bacterium that displays on its surface a minicellulosome including two or more cellulolytic enzymes, wherein said bacterium includes (e.g., the minicellulosome includes): a first protein encoding one or more cohesin domains wherein said protein is covalently linked to the surface of said microorganism, and wherein each of one or more cohesin domains is docked to a dockerin attached to a cellulolytic enzyme and said second protein additionally encodes a linking dockerin or a linking cohesin; a second protein encoding one or more cohesin domains wherein each of one or more cohesin domains is docked to a dockerin attached to a cellulolytic enzyme and said second protein additionally encodes a linking dockerin or a linking cohesin; wherein said second protein is docked to said first protein by a dockerin/cohesin interaction between said linking dockerin or linking cohesin encoded by said second protein and said linking dockerin or linking cohesin encoded by said first protein, where when said linking dockerin or linking cohesin on said first protein is a linking cohesin, said linking dockerin or linking cohesin on said second protein is a linking dockerin, and when said linking dockerin or linking cohesin on said first protein is a linking dockerin, said linking dockerin or linking cohesin on said second protein is a linking cohesin.
The bacterium of embodiment 39, wherein said first protein encodes a linking cohesin and said second protein encodes a linking dockerin and said second protein is attached to said first protein by a dockerin/cohesin interaction between said linking cohesin on said first protein and said linking dockerin on said second protein.
The bacterium of embodiment 39, wherein said first protein encodes a linking dockerin and said second protein encodes a linking cohesin and said second protein is attached to said first protein by a dockerin/cohesin interaction between said linking dockerin on said first protein and said linking cohesin on said second protein.
The bacterium according to any one of embodiments 39-41, wherein said one or more cohesin domains in said first protein comprise at least two cohesin domains each docked to a cellulolytic enzyme attached to a dockerin.
The bacterium according to any one of embodiments 39-42, wherein said one or more cohesin domains in said first protein comprise at least three cohesin domains each docked to a cellulolytic enzyme attached to a dockerin.
The bacterium according to any one of embodiments 39-43, wherein said one or more cohesin domains in said second protein comprise at least two cohesin domains each docked to a cellulolytic enzyme attached to a dockerin.
The bacterium of embodiment 44, wherein said one or more cohesin domains in said second protein comprise at least three cohesin domains each docked to a cellulolytic enzyme attached to a dockerin.
The bacterium according to any one of embodiments 39-45, wherein: said bacterium includes a third protein encoding one or more cohesin domains wherein each of one or more cohesin domains is docked to a dockerin attached to a cellulolytic enzyme and said third protein additionally encodes a linking dockerin or a linking cohesin; said second protein includes a second linking dockerin or a second linking cohesin; wherein said third protein is docked to said second protein by a dockerin/cohesin interaction between said second linking dockerin or second linking cohesin encoded by said second protein and said linking dockerin or linking cohesin encoded by said third protein, where when said second linking dockerin or linking cohesin on said second protein is a linking cohesin, said linking dockerin or linking cohesin on said third protein is a linking dockerin, and when said second linking dockerin or linking cohesin on said second protein is a linking dockerin, said linking dockerin or linking cohesin on said second protein is a linking cohesin.
The bacterium of embodiments 46, wherein said second linking dockerin or linking cohesin on said second protein is a second linking cohesin, said linking dockerin or linking cohesin on said third protein is a linking dockerin and said third protein is attached to said second protein by a dockerin/cohesin interaction between said second linking cohesin on said second protein and said linking dockerin on said third protein.
The bacterium of embodiment 46, wherein said second linking dockerin or linking cohesin on said second protein is a linking dockerin and said linking dockerin or linking cohesin on said third protein is a linking cohesin and said third protein is attached to said second protein by a dockerin/cohesin interaction between said second linking dockerin on said second protein and said linking cohesin on said third protein.
The bacterium according to any one of embodiments 46-48, wherein said one or more cohesin domains in said third protein comprise at least two cohesin domains each docked to a cellulolytic enzyme attached to a dockerin.
The bacterium of embodiment 49, wherein said one or more cohesin domains in said third protein comprise at least three cohesin domains each docked to a cellulolytic enzyme attached to a dockerin.
The bacterium according to any one of embodiments 39-50, wherein said cellulases form a cellulosome that self assembles on said bacterium.
The bacterium according to any one of embodiments 39-51, wherein said bacterium grows on untreated plant biomass.
The bacterium according to any one of embodiments 39-52, wherein said bacterium grows on lignocellulose as the sole carbon source.
The bacterium according to any one of embodiments 39-53, wherein said minicellulosome includes at least three cellulolytic enzymes and all of said enzymes are encoded by said bacterium.
The bacterium according to any one of embodiments 39-54, wherein one or more of the cohesin domains in said first protein that are docked to dockerin-bearing enzymes are Type-I cohesins.
The bacterium according to any one of embodiments 39-55, wherein one or more of the cohesin domains in said second protein that are docked to dockerin-bearing enzymes are Type-I cohesins.
The bacterium according to any one of embodiments 46-64, wherein one or more of the cohesin domains in said third protein that are docked to dockerin-bearing enzymes are Type-I cohesins.
The bacterium according to any one of embodiments 39-57 wherein the linking dockerin/cohesins joinining said first protein to said second protein are Type II dockerins and cohesins.
The bacterium according to any one of embodiments 39-57 wherein the linking dockerin/cohesins joining said second protein to said third protein, when said third protein is present, are Type II dockerins and cohesins.
The bacterium according to any one of embodiments 39-59, wherein said first protein encoding two or more cohesin domains includes a secretory signal.
The bacterium of embodiment 60, wherein said secretory signal sequence includes a B. subtilis phrC secretory signal or homologues thereof.
The bacterium of embodiment 61, wherein said secretory signal sequence includes a secretion signal derived from B. subtilis phrC.
The bacterium according to any one of embodiments 39-62, wherein said first protein encoding two or more cohesin domains includes a cell wall sorting signal (CWSS).
The bacterium of embodiment 63, wherein said protein encoding two or more cohesin domains includes a secretory signal sequence at the N-terminus and a cell wall sorting signal (CWSS) at the carboxyl terminus.
The bacterium according to any one of embodiments 63-64, wherein said cell wall sorting signal includes an LPXTG motif.
The bacterium according to any one of embodiments 63-64, wherein said cell wall sorting signal includes a sequence shown in Table 1.
The bacterium according to any one of embodiments 63-64, wherein said cell wall sorting signal includes a CWSS from Staphylococcus aureus fibronectin binding protein B.
The bacterium according to any one of embodiments 39-67, wherein said first protein encoding two or more cohesin domains and/or said second protein encoding two or more cohesin domains encodes a carbohydrate binding module (CBM).
The bacterium of embodiment 68, wherein said carbohydrate binding module is a family 3 carbohydrate binding module.
The bacterium of embodiment 68, wherein said carbohydrate binding module is a carbohydrate binding module derived from C. thermocelllum CipA.
The bacterium according to any one of embodiments 39-69, wherein said two or more cohesin domains including said first protein, and/or said two or more cohesin domains including said second protein and/or said two or more cohesin domains including said third protein, when present, are cohesin domains from different microorganisms.
The bacterium according to any one of embodiments 39-71, wherein said two or more cohesin including said first protein, and/or said two or more cohesin domains including said second protein and/or said two or more cohesin domains including said third protein, when present, comprise a cohesin domain from an organism selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus.
The bacterium according to any one of embodiments 39-72, wherein said two or more cohesin including said first protein, and/or said two or more cohesin domains including said second protein and/or said two or more cohesin domains including said third protein, when present, comprise cohesin domains from two organisms selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus.
The bacterium according to any one of embodiments 39-73, wherein said two or more cohesin including said first protein, and/or said two or more cohesin domains including said second protein and/or said two or more cohesin domains including said third protein, when present, comprise cohesin domains from three organisms selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus
The bacterium according to any one of embodiments 39-73, wherein said two or more cohesin including said first protein, and/or said two or more cohesin domains including said second protein and/or said two or more cohesin domains including said third protein, when present, comprise cohesin domains from C. thermocellum (t), C. cellulolyticum (c) and R. flavefaciens (f).
The bacterium according to any one of embodiments 39-75, wherein the dockerins coupling said cellulolytic enzymes to the cohesins including said first protein and/or said second protein, and/or said third protein when present comprise one or more dockerin domains from organism(s) selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus.
The bacterium according to any one of embodiments 39-75, wherein the dockerins coupling said cellulolytic enzymes to the cohesins including said first protein and/or said second protein, and/or said third protein when present comprise two or more dockerin domains from organism(s) selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus.
The bacterium according to any one of embodiments 39-75, wherein the dockerins coupling said cellulolytic enzymes to the cohesins including said first protein and/or said second protein, and/or said third protein when present comprise three or more dockerin domains from organism(s) selected from the group consisting of Clostridium thermocellum, Clostridium cellulolyticum, Ruminococcus flavefaciens, C. cellulovorans, C. acetobutylicum, C. josui, C. papyrosolvens, A. cellulolyticus, and R. albus.
The bacterium according to any one of embodiments 39-75, wherein the dockerins coupling said cellulolytic enzymes to the cohesins including said first protein and/or said second protein, and/or said third protein when present comprise dockerin domains from C. thermocellum (t), C. cellulolyticum (c) and R. flavefaciens (f).
The bacterium according to any one of embodiments 39-79, wherein said cellulolytic enzyme(s) on dormant bacteria are stable for at least 1 day, more preferably for at least 2 days, and most preferably at least 3 days.
The bacterium according to any one of embodiments 39-80, wherein said cellulolytic enzyme(s) comprise one or more enzymes selected from the group consisting of an endocellulase, an exocellulase, a beta-glucosidase (cellobiase), an oxidative cellulase, a xylanase, a hemicellulase, a lichenase, a chitenase, a mannase, an exogluconase, an endoxylanase, exogluconase, and a cellulose phosphorylase.
The bacterium according to any one of embodiments 39-81, wherein said minicellulosome includes at least two different cellulolytic enzymes, or at least 3 different cellulolytic enzymes, or at least 4 different cellulolytic enzymes, or at least 5 different cellulolytic enzymes, or at least 6 different cellulolytic enzymes.
The bacterium according to any one of embodiments 39-82, wherein said minicellulosome includes at least 6 different cellulolytic enzymes.
The bacterium according to any one of embodiments 39-83, wherein said minicellulosome includes at least one endoglucanase.
The bacterium according to any one of embodiments 39-84, wherein said minicellulsome includes at least one exoglucanase.
The bacterium according to any one of embodiments 39-85, wherein said minicellulsome includes at least two endoglucanases and at least one exoglucanase.
The bacterium of embodiment 83, wherein said minicellulosome includes an endoglucanase, a xylanase, an exoglucanase, an endoxylanase, and a mannase.
The bacterium according to any one of embodiments 82-83, wherein said minicellulosome includes Cel5A.
The bacterium according to any one of embodiments 82-83 and 88, wherein said minincellulosome includes XynA.
The bacterium according to any one of embodiments 82-83 and 88-89, wherein said minincellulosome includes Cel48F.
The bacterium according to any one of embodiments 82-83 and 88-90, wherein said minincellulosome includes CelS.
The bacterium according to any one of embodiments 82-83 and 88-91, wherein said minincellulosome includes Cel9E.
The bacterium according to any one of embodiments 82-83 and 88-92, wherein said minincellulosome includes Man5A.
The bacterium according to any one of embodiments 39-93, wherein said Gram-positive bacterium includes a Gram-positive bacterium that encodes a sortase.
The bacterium of embodiment 94, wherein said Gram-positive bacterium includes a Gram-positive bacillus.
The bacterium of embodiment 95, wherein said Gram-positive bacterium includes a genus selected from the group consisting of Corynebacterium, Clostridium, Listeria, and Bacillus.
The bacterium of embodiment 96, wherein said bacterium is a Clostridium acetobutylicum.
The bacterium of embodiment 96, wherein said Gram-positive bacterium comprise is B. subtilis.
The bacterium of embodiment 94, wherein said Gram-positive bacterium includes a thermophilic Geobacillus spp.
The bacterium of embodiment 94, wherein said Gram-positive bacterium includes a Gram-positive coccus.
The bacterium of embodiment 100, wherein said bacterium is selected from the group consisting of S. aureus, S. epidermis, and S. saprophyticus.
A method of degrading cellulosic biomass into fermentable sugars, said method including: contacting said cellulosic biomass with a bacterium according to any one of embodiments 1-101, under conditions in which said bacteria partially or fully degrade cellulose in said cellulosic biomass to form one or more fermentable sugars.
The method of embodiment 102, wherein said contacting includes contacting dormant bacteria to said cellulosic biomass.
The method of embodiment 102, wherein said contacting includes culturing said bacteria with said cellulosic biomass.
The method according to any one of embodiments 102-104, wherein said cellulosic biomass includes lignocellulosic biomass.
The method according to any one of embodiments 102-105, wherein said cellulosic biomass comprise one or more materials selected from the group consisting of an agricultural plant waste (e.g., corn stover, cereal straw, sugarcane bagasse), a plant waste from an industrial process (e.g., sawdust, paper pulp), and a non-food energy crop (e.g., switchgrass).
The method of embodiment 106, wherein said cellulosic biomass includes one or more materials selected from the group consisting of grasses, rice hulls, bagasse, jute, hemp, flax, bamboo, sisal, abaca, straw, corn cobs, corn stover, alfalfa, hay, coconut hair, seaweed, and algae.
The method according to any one of embodiments 102-106, wherein said cellulosic biomass is not acidified and/or enzymatically pre-digested.
A consolidated bioreactor for the conversion of a lignocellulosic biomass into bioethanol said bioreactor including: a culture system that cultures bacteria according to any one of embodiments 1-101 under conditions in which said bacteria partially or fully degrade cellulose in said lignocellulosic biomass to form one or more fermentable sugars; and a culture system that ferments said sugars to form a biofuel.
A method of identifying cellulolytic enzyme combinations that enhance degradation of a particular substrate said method including: providing a plurality of recombinant bacteria according to any one of embodiments 1-101, wherein said bacteria each display at least two cellulolytic enzymes and different bacteria display different enzymes; contacting said substrate with said bacteria; and selecting bacteria that show enhanced degradation of said substrate and/or improved growth on said substrate.
A method of identifying cellulolytic enzyme variants that enhance degradation of a particular substrate said method including: providing a plurality of recombinant bacteria according to any one of embodiments 1-101, wherein said bacteria each display at least one cellulolytic enzyme variant and different bacteria display different cellulolytic enzyme variants; contacting said substrate with said bacteria; and selecting bacteria that show enhanced degradation of said substrate and/or improved growth on said substrate.
The method according to any one of embodiments 110 to 111, wherein said cellulolytic enzyme(s) and/or said cellulolytic enzyme variants comprise a mutant cellulolytic enzyme.
The method of embodiment 112, wherein said mutant cellulolytic enzyme includes a mutant cellulase.
The method according to any one of embodiments 110 to 113, wherein said selecting includes selecting bacteria that show improved growth on said substrate.
The term “nucleic acid” refers to a nucleotide polymer, and unless otherwise limited, includes known analogs of natural nucleotides that can function in a similar manner (e.g., hybridize) to naturally occurring nucleotides. The term nucleic acid includes any form of DNA or RNA, including, for example, genomic DNA; complementary DNA (cDNA), which is a DNA representation of mRNA, usually obtained by reverse transcription of messenger RNA (mRNA) or by amplification; DNA molecules produced synthetically or by amplification; and RNA. The term nucleic acid encompasses double- or triple-stranded nucleic acid, as well as single-stranded molecules. In double- or triple-stranded nucleic acids, the nucleic acid strands need not be coextensive (i.e., a double-stranded nucleic acid need not be double-stranded along the entire length of both strands). The term nucleic acid also encompasses any chemical modification thereof, such as by methylation and/or by capping. Nucleic acid modifications can include addition of chemical groups that incorporate additional charge, polarizability, hydrogen bonding, electrostatic interaction, and functionality to the individual nucleic acid bases or to the nucleic acid as a whole. Such modifications may include base modifications such as 2′-position sugar modifications, 5-position pyrimidine modifications, 8-position purine modifications, modifications at cytosine exocyclic amines, substitutions of 5-bromo-uracil, backbone modifications, unusual base pairing combinations such as the isobases isocytidine and isoguanidine, and the like.
The terms “isolated”, when referring to an isolated nucleic acid or nucleic acid construct refers to a nucleic acid that either does not exist normally in nature, and/or that is constructed using for example, recombinant DNA techniques, and/or that is removed from nucleic acid sequences that would normally flank it in vivo, and/or that is removed from a cellular milieu. Isolated nucleic acids also include nucleic acids derived from the foregoing isolated nucleic acids, e.g., by propagation of a construct/vector/organism/virus/or microorganism containing such nucleic acid sequences.
“Operably linked” means that a gene (or other sequence to be expressed) and transcriptional regulatory sequence(s) are connected in such a way as to permit expression of the gene under control of the regulatory sequence(s).
“Exogenous” means a nucleic acid sequence that has been inserted into a host cell or a nucleic acid sequence or amino acid sequence derived from a nucleic acid sequence that has been inserted into a host cell. This includes introduced (inserted) nucleic acids that remain into the cytoplasm and introduced nucleic acids that integrate into the host cell genome (e.g., plasmids inserted into the host genome) as well as nucleic acid sequences and/or amino acids sequences derived from such. In certain embodiments an exogenous sequence can result from the cloning of a native gene from a host cell and the reinsertion of that sequence back into the host cell. In most instances, exogenous sequences are sequences that are derived synthetically, or from cells that are distinct from the host cell.
The terms “host cells” and “recombinant host cells” are used interchangeably herein. It is understood that such terms refer not only to the particular subject cell but to the progeny or potential progeny of such a cell. Because certain modifications may occur in succeeding generations due to either mutation or environmental influences, such progeny may not, in fact, be identical to the parent cell, but are still included within the scope of the term as used herein.
The term “cellulolytic enzyme” refers to an enzyme that can participate in the degradation of cellulose or a cellulosic biomass.
The term “cellulosic biomass” refers to plant, algal, or other biomass that contains cellulose.
Lignocellulosic biomass refers to plant biomass that typically contains cellulose, hemicellulose, and lignin. The carbohydrate polymers (cellulose and hemicelluloses) are often tightly bound to the lignin. Lignocellulosic biomass can be grouped into four main categories: (1) agricultural residues (including corn stover and sugarcane bagasse), (2) dedicated energy crops, (3) wood residues (including sawmill and paper mill discards), and (4) municipal paper waste. Illustrative lignocellulosic biomass sources include, but are not limited to grasses, rice hulls, bagasse, jute, hemp, flax, bamboo, sisal, abaca, straw, corn cobs, corn stover, alfalfa, hay, coconut hair, seaweed, algae, and the like.
A cellulase is an enzyme that breaks down cellulose, especially in the wall structures, and a “cellulosome” is an array, cluster, or sequence of enzymes or cellulases that degrades cellulose. In various embodiments cellulosomes comprise catalytic subunits such as glycoside hydrolases, polysaccharide lyases and carboxyl esterases bound together by scaffoldins consisting of cohesins (cohesin domains) connected to other functional units such as the enzymes and carbohydrate binding modules via dockerins.
A “cohesin” or “cohesin domain” refers to a protein domain that interacts with a complementary domain, termed a “dockerin” or “dockerin domain”. Cohesin-dockerin interactions mediate the formation of cellulosome, or minicellulosomes.
The terms “linking dockerin” and “linking cohesin” refers to cohesins (cohesin domains) and dockerins (dockerin domains) that joint two backbone proteins (e.g., scaffoldins) to each other through a cohesin/dockerin interaction. in certain embodiments the linking dockerin is a type II dockerin and the linking cohesin is a type II cohesin.
A “protein encoding one or more cellulolytic enzymes” or a “protein comprising one or more cellulolytic” refers to a protein at least a portion of which displays cellulolytic activity. In certain embodiments the protein comprises a single cellulolytic enzyme and substantially the entire protein (absent processing and/or signaling sequences) comprises a single enzyme (e.g., a cellulase). In certain embodiments the protein comprises multiple (e.g., 2, 3, 4, 5, 6, or more) cellulolytic enzymes and in such instances each enzyme comprises a different “domain” in said protein. Similarly a protein comprising or encoding multiple cohesins refers to a protein comprising one or more domains each of which has the amino acid sequence of a cohesin, and in certain embodiments, is capable of binding to a corresponding dockerin.
When a Markush Group is described in the specification and/or claims it is intended that in various additional or alternative any subset of that Markush group is contemplated. Thus, for example, a Markush group consisting of elements A, B, and C also comprises a disclosure of a Markush Group consisting of A, and B, a Markush Group consisting of B, and C, and a Markush Group consisting of A and C as well as elements A, B, and C individually.
Where a range of values is provided, it is understood that each intervening value between the upper and lower limit of that range and any other stated or intervening value in that stated range, is contemplated. The upper and lower limits of these smaller ranges may independently be included in the smaller ranges, and are also contemplated, subject to any specifically excluded limit in the stated range. Where the stated range includes one or both of the limits, ranges excluding either or both of those included limits are also contemplated.
It is desirable to produce biofuels and other bio-based chemicals and materials from renewable plant biomass (lignocellulose). One promising strategy is to create microbes that are consolidated bioprocessors (CBP). These microorganisms will breakdown biomass into its component sugars and then convert the degradation products into desired chemicals. At present, only a few CBP microbes have been developed and to the best of our knowledge none of them is widely used in industry. Bacillus subtilis is a promising CBP as it is already used in industry to produce a range of compounds (proteins, antibiotics and insecticides). Moreover, it has a robust genetic system, making it well suited for metabolic engineering which could enable it to produce other useful compounds. However, native strains of B. subtilis cannot efficiently degrade lignocellulose and use it as a nutrient to grow.
In various embodiments a protein display system is provided that enables multi-enzyme complexes to be self-assembled on the surface of B. subtilis. It is demonstrated that this new system can be used to create recombinant B. subtilis strains that can efficiently degrade lignocellulosic biomass. Furthermore, these cells can use biomass as a nutrient to grow. Additional modifications of the protein display system enable the number and types of enzymes displayed to be significantly increased to make even more potent cellulolytic organisms. The protein system can also be readily ported to other Gram-positive microbes which will enable them be used as a CBP.
This work has several potentially useful applications. Using the protein display system described herein: (1) the cellulolytic B. subtilis cells can be further engineered to develop a CBP that produces biocommodities such as ethanol from biomass, (2) It can be used to create highly cellulolytic B. subtilis cells that can replace more costly enzyme cocktails that are currently being used in industry to degrade biomass, and (3) The system we have developed can readily be ported to other Gram-positive bacterial species so as to enable them to use lignocellulose biomass as a nutrient.
Unlike most other recombinant cellulosome or minicellulosome systems, in the systems/organisms described herein all of the components of the minicellulosome are expressed in the microbe which effectively self-assembles the cellulosome. In addition, the recombinant organisms described herein display cellulase enzymes that are better suited for degrading lignocellulosic biomass (e.g., switchgrass, straw, corn stover, and the like).
It is believed the constructs described herein represent the first example of a self-assembling type-1 minicellulosome on the surface of B. subtilis. We have demonstrated that B. subtilis cells displaying the minicellulosome can efficiently degrade untreated biomass (corn stover, switchgrass, and straw). This is beneficial in that it potentially avoids costly pretreatment of the biomass (e.g. acid treatment) that is currently being used in industry. To the best of our knowledge, no recombinant microbe has ever been shown to be capable of growing on untreated biomass. Other microbes have been engineered to have cellulolytic activity, but they have only been shown to degrade purified cellulose substrates, such as phosphoric acid swollen cellulose and regenerated amorphous cellulose.
An illustrative schematic of one embodiment of the minicellulosome is shown in
In fact, it has recently been noted that only twelve biomass-derived building blocks are needed to produce a range of commercial products. We have shown that our protein display system can be used to create B. subtilis strains that can grow on plant biomass. It is believed that cells or variant thereof can be used to produce these biocommodities or building-blocks from cheap plant biomass. An immediate application is to introduce the cellulolytic system described herein into existing organisms that are already used industrially to produce commercial products. This would enable the products to be produced from biomass and could significantly reduce costs.
Another application of the system described herein is to use the biomass-degrading cells as a replacement for enzyme cocktails that are currently used in industry to degrade biomass. The cells can be produced more cheaply than the enzymes and thereby reduce the costs associated with degrading biomass into its component sugars. An immediate application is lignocellulose degradation needed to produce lignocellulosic ethanol. Cells displaying the minicellulosomes would degrade the biomass into component sugars, which are then used in downstream ethanol producing fermentation reactions performed by S cerevisiae.
The system we have developed could also be used to engineer other species of Gram-positive bacteria to convert them into CPB or microbes that are dedicated to degrading biomass into sugars. The working prototype described herein has been demonstrated to successfully degrade lignocellulosic biomass into monosaccharides and oligosaccharides. It has also been demonstrated to grow on lignocellulose as the sole carbon source, indicating that these strains are well suited for use in a consolidated bioprocessor.
As explained herein, in various embodiments, a sortase transpeptidase (e.g., Sortase A or analogues, homologues, or orthologues thereof) is exploited to couple a protein (e.g., a “scaffoldin” protein comprising one or a plurality of cohesin domains) to the surface (e.g., cell wall) of a Gram-positive microorganism. To facilitate this, the peptide can be provided with a cell wall sorting signal sequence that is recognized by the sortase transpeptidase.
The examples provided herein use a C-terminal portion of the Staphylococcus aureus Fibronectin Binding Protein B, which contains a 123 amino acid spacer segment and the cell wall sorting signal (CWS). This S. aureus CWS sequence is identical to CWS found in many B. anthracis surface proteins that are anchored to the cell wall of B. anthracis by SrtA
Typically cell wall sorting signals comprise an LPXTG (SEQ ID NO: 1) motif (where X is any amino acid), a C-terminal hydrophobic domain and a charged tail. Homologous sequences are found in many surface proteins of Gram-positive bacteria (see, e.g., Schneewind et al. (1993) EMBO J., 12(12): 4803-4811, which describes a number of cell wall sorting signals, illustrated below in Table 1).
LPETGEENPFIGTTVFGGLSLALGAALLAGRRREL
S. aureus
LPETGGEESTNKGMLFGGLFSILGLALLRRNKKNHKA
S. aureus
LPATGDSSNAYLPLLGLVSLTAGFSLLGLRRKQD
S. sobrinus
LPKTGEKQNVLLTVVGSLAAMLGLAGLGFKRRKETK
E. faecalis
LPSTGSIGTYLFKAIGSAAMIGAIGIYIVKRRKA
S. pyogenes
LPTTGDSDNALYLLLGLLAVGTAMALTKKARASK
L. monocytogenes
LPLTGANGVIFLTIAGALLVAGGAVVAYANKRRHVAKH
A. viscosus
LPYTGVASNLVLEIMGLLGLIGTSFIAMKRRKS
S. agalactiae
LPKTGMKIITSWITWVFIGILGLYLILRKRFNS
S. aureus
LPSTGEQAGLLLTTVGLVIVAVAGVYFYRTRR
S. mutans
LPSTGETANPFFTAAALTVMATAGVAAVVKRKEEN
S. pyogenes
These cell wall sorting signals are intended to be illustrative and not limiting. Using the teachings provided herein, numerous other cell wall sorting signals can be incorporated in the expression/display systems described herein.
While in certain embodiments, cell wall sorting signals comprising the LPXTG motif are preferred, they need not be limited to this motif. Based on homology sortases thus far identified are typically grouped into four or five subgroups or classes (see, Table 2). Each subgroup, in addition to distinctions in sequence, can be distinguished from one another based on membrane topology, genome position, and preference for substrates with specific amino acids within the cell wall sorting signal pentapeptide motif (Comfort and Clubb (2004) Infect. Immun., 72: 2710-2722; Dramsi et al. (2005) Res. Microbiol. 156: 289-297). As indicated above, the prototypical sortase is sortase A, first identified in S. aureus. Sortase A appears to anchor a large number and broad range of surface proteins. The sortase A subgroup of enzymes also seems to share a preference for the LPXTG (SEQ ID NO: 13) cell wall sorting signal motif. The second subgroup of enzymes, sortase B, along with its substrate (IsdC in S. aureus), is encoded in an iron transport operon involved in heme-iron uptake. Enzymes belonging to the sortase B subgroup contain three amino acid segments not found in sortase A and recognize substrates containing an NPQTN (SEQ ID NO:14) motif rather than the canonical LPXTG (SEQ ID NO: 15). The third class, designated sortase C or subfamily 3, contains a C-terminal hydrophobic domain (Id.). Subfamily 3 enzymes also share a preference for substrates containing the LPXTG cell wall sorting signal motif, often followed by a second G residue (i.e., LPXTGG, (SEQ ID NO:16). A fourth subgroup can be defined after alignment of sortase sequences. It has been noted as the sortase D subgroup (Dramsi et al. (2005) Res. Microbiol. 156: 289-297) or subfamilies 4 and 5, as sortases in this subgroup can be distinguished based on the cell wall sorting signals of their associated substrates (Comfort and Clubb (2004) Infect. Immun., 72: 2710-2722). Sortases belonging to subfamily 4 are predicted to anchor proteins bearing the unique LPXTA(ST) (SEQ ID NO:17) motif (Id.). An alanine residue in the last position of the substrate motif suggests that the subfamily 4 enzymes fulfill a nonredundant role within the cell (Id.). Many high-G/C bacteria contain sortases belonging to subfamily 5, and most do not harbor sortase A. This subgroup of sortase enzymes shares substrate specificity for proteins containing an LAXTG (SEQ ID NO:18) motif (Id.).
Bacillus, Listeria,
Staphylococcus, Enterococcus,
Bacillus, Listeria,
Staphylococcus,
Enterococcus,
Bacillus
bCell wall sorting signal pentapeptide motif. Uppercase letters represent amino acids that are absolutely conserved. Asterisks indicate that the cleavage site has been verified experimentally.
Accordingly in various embodiments, display systems that utilize any of these cell wall sorting sequences are contemplated for use in the methods and constructs described herein.
As described herein, in various embodiments, the display system(s) utilize more proteins (e.g., scaffoldins) comprising one or more, preferably two or more, cohesin domains (e.g., cohesin I domains) that interact with dockerin domains to anchor and/or organize one or more enzymes on the surface of the Gram-positive bacterium.
In various embodiments the systems contemplated herein can comprise one or more dockerin domains selected from the group consisting of a dockerin I domain, a dockerin II domain, and a dockerin III domains. Correspondingly, in various embodiments the systems contemplated herein can comprise one or more cohesin domains selected from the group consisting of a cohesin I domain, a cohesin II domain, and a cohesin III domains that binds to its corresponding dockerin sequence. In certain embodiments the dockerin and/or cohesin domains comprise a domain derived from Clostridium thermocellum.
The sequences of cohesins and dockerins are well known to those of skill in the art (see, e.g., Ding et al. (2003) Genet. Eng. (N Y) 25: 209-225 for cellulosome cohesins and dockerins, and Peer et al. (2009) FEMS Microbiol Lett. 291(1): 1-16 for non-cellulosome cohesins and dockerins). In various embodiments specific cohesin-dockerin pairs are chosen so as to enable specific complexes to form on the cell surface even if multiple enzymes are present.
While the display system described herein is exemplified using cohesin-dockerin pairs, it will be recognized that, in certain embodiments, other protein-protein interaction pairs can be used as long as one member of the pair becomes covalently attached to the cell wall and the other is fused to the cellulolytic enzyme(s) so as to enable enzyme complex formation on the cell surface.
In various embodiments Gram-positive bacteria are engineered using the methods described herein to display one or more enzymes. In certain embodiments the enzymes are cellulolytic enzymes and/or other enzymes useful in the synthesis of biofuels from lignocellulosic biomass. In various embodiments it will be recognized that the “cellulases” can include, but are not limited to, the cellobiohydrolases, e.g., cellobiohydrolase I and cellobiohydrolase II, as well as the endoglucanases. In various embodiments “cellulolytic enzymes” include, but are not limited to, cellobiohydrolases, e.g. cellobiohydrolase I and cellobiohydrolase II, as well as endoglucanases and beta-glucosidases.
In various embodiments the digestion of cellulose and hemicellulose is facilitated by the use of several types of enzymes acting cooperatively. In certain embodiments at least three categories of enzymes are utilized to convert cellulose into fermentable sugars: endoglucanases that cut the cellulose chains at random; cellobiohydrolases that cleave cellobiosyl units from the cellulose chain ends and beta-glucosidases that convert cellobiose and soluble cellodextrins into glucose. Among these three categories of enzymes involved in the biodegradation of cellulose, cellobiohydrolases are useful for the degradation of native crystalline cellulose. Cellobiohydrolase I, also referred to as a cellulose 1,4-beta-cellobiosidase or an exoglucanase, exo-cellobiohydrolase or 1,4-beta-cellobiohydrolase catalyzes the hydrolysis of 1,4-beta-D-glucosidic linkages in cellulose and cellotetraose, by the release of cellobiose from the non-reducing ends of the chains. Cellobiohydrolase II activity is identical, except that cellobiohydrolase II attacks from the reducing ends of the chains.
In various embodiments the cellulolytic enzymes are organized into a cellulosome or minicellulosome (see, e.g.,
The displayed cellulosomes can be simple cellulosome systems containing a single scaffoldin or complex cellulosome systems that exhibit multiple types of interacting scaffoldins. In various embodiments each scaffoldin can contain one, two, three, four, five, six, seven, eight, nine, or 10 or more cohesin domains. The arrangement of the modules on the scaffoldin subunit and the specificity of the cohesin(s) and/or dockerin for their modular counterpart determine the overall architecture of the cellulosome. Several different types of scaffoldins have been described and are useful in the construction of minicellulosomes according to the methods described herein. The primary scaffoldins incorporate the various dockerin-bearing subunits directly into the cellulosome complex, adaptor scaffoldins increase the repertoire or number of components into the complex, and the anchoring scaffoldins attach the complex to the bacterial cell surface.
Scaffoldins are well known to those of skill in the art and can readily be identified with a simple GenBank search for the term “scaffoldin”.
In certain embodiments the cellulolytic enzymes comprising the cellulosome or individually displayed on the surface of the bacteria comprise one or more enzymes collected from the group consisting of an exoglucanase, an endoglucanase, a glycosyl hydrolase, a cellulase, a hemicellulase, a xylanase, a cellobiohydrolase, a beta-glucosidase, a mannanse, a xylosidase (e.g., a β-xylosidase), an arabinofuranosidase, and/or a glucose oxidase. Illustrative, but non-limiting, enzymes suitable for display using the systems described herein are shown in Table 3.
Clostridium thermocellum endoglucanse CelG
Clostridium thermocellum endoglucanse CelD
Clostridium thermocellum endoglucanse CelQ
Clostridium thermocellum endoglucanse CelR
Clostridium thermocellum endoglucanse CelN
Clostridium thermocellum exoglucanase CelS
Clostridium thermocellum cellobiohydrolase CbhA
Clostridium thermocellum cellobiohydrolase CelK
Clostridium thermocellum cellobiohydrolase CelO
Clostridium thermocellum xylanase XynD
Clostridium thermocellum xylanase XynC
Clostridium thermocellum xylanase XynA
Clostridium thermocellum lichenase LicB
Clostridium thermocellum chitinase ChiA
Clostridium thermocellum mannase ManA
In addition, a large number of other suitable enzymes are described in U.S. Patent Publication 2010/0189706 which is incorporated herein by reference for any one or more of the cellulolytic enzymes described herein. Cellulosomes are also described by Fontes and Gilbert (2010) Annu. Rev. Biochem., 79: 655-681.
In certain embodiments the cellulosome that is to be displayed can be engineered based upon the cellulosic material to be metabolized. For example, different cellulases and other enzymes may be engineered into a cellulosome pathway depending upon the sources of substrate. Illustrative substrate sources include, but are not limited to, alfalfa, corn stover, crop residues, debarking waste, forage grasses, forest residues, municipal solid waste, paper mill residue, pomace, sawdust, spent grains, spent hops, switchgrass, and wood chips. Some substrate sources can have a larger percentage of cellulose compared to other source, which may have a larger percentage of hemicellulose.
A hemicellulose substrate typically comprises short, branched chains of sugars and can comprise a polymer of five different sugars. Hemicellulose comprises five-carbon sugars (e.g., D-xylose and L-arabinose) and six-carbon sugars (e.g., D-galactose, D-glucose, and D-mannose) and uronic acid. The sugars are typically substituted with acetic acid. Hemicellulose is relatively easy to hydrolyze to its constituent sugars. When hydrolyzed, the hemicellulose produces xylose (a five-carbon sugar) or six-carbon sugars from hardwoods or softwoods, respectively.
Proteins or polypeptides having the ability to convert the hemicellulose components into carbon sources that can be used as a substrate for biofuel production includes, for example, cellobiohydrolases (Accessions: AAC06139, AAR87745, EC 3.2.1.91, 3.2.1.150), cellulases (E.C. 3.2.1.58, 3.2.1.4, Accessions: BAA12070, BAB64431); chitinases (E.C. 3.2.1.14, 3.2.1.17, 3.2.1.-, 3.2.1.91, 3.2.1.8, Accessions: CAA93150, CAD12659), various endoglucanases (E.C. 3.2.1.4, Accessions: BAA92430, AAG45162, P04955, AAD39739), exoglucanases (E.C. 3.2.1.91, Accessions: AAA23226), lichenases (E.C. 3.2.1.73, Accessions: P29716), mannanases (E.C. 3.2.1.4, 3.2.1.-, Accessions: CAB52403), pectate lyases (E.C. 4.2.2.2, Accessions: AAG59609), xylanase (E.C. 3.2.1.136, 3.2.1.156, 3.2.1.8, Accessions: BAA33543, CAA31 109) and silase (E.C. 3.2.2.-, 2.7.7.7, Accessions: CQ80097S).
Cellulases are a class of enzymes produced chiefly by fungi, bacteria, and protozoans that catalyze the hydrolysis of cellulose. However, there are also cellulases produced by other types of organisms such as plants and animals. Several different kinds of cellulases are known, which differ structurally and mechanistically. The EC number for cellulase enzymes is E.C. 3.2.1.4. Assays for testing cellulase activity are known in the art.
Polypeptides having xylanase activity are also useful in synthetic cellulosomes. Xylanase is the name given to a class of enzymes which degrade the linear polysaccharide beta-1,4-xylan into xylose, thus breaking down hemicellulose. The EC number for xylanase enzymes is E.C. 3.2.1.136, 3.2.1.156, 3.2.1.8. Assays for testing xylanase activity are known in the art.
In certain embodiments the minicellulosome comprises at least two different, or at least three different, or at least four different, or at least five different, or at least six different, or at least seven different, or at least 8 different, or at least 9 different, or at least 10 different, or at least 11 different, or at least 12 different cellulolytic (or other degredative) enzymes. In certain embodiments the two enzymes comprise two different or three different, or four different, or five different, or six different enzymes selected from the group consisting of endocellulase/endocellulase, exocellulase/endocellulase, beta-glucosidase (cellobiase)/endocellulase, oxidative cellulase/endocellulase, xylanase/endocellulase, hemicellulase/endocellulase, lichenase/endocellulase, chitenase/endocellulase, xylanase/endocellulase, cellulose phosphorylase/endocellulase, endocellulase/exocellulase, exocellulase/exocellulase, beta-glucosidase (cellobiase)/exocellulase, oxidative cellulase/exocellulase, xylanase/exocellulase, hemicellulase/exocellulase, lichenase/exocellulase, chitenase/exocellulase, xylanase/exocellulase, cellulose phosphorylase/exocellulase, endocellulase/beta-glucosidase, exocellulase/beta-glucosidase, beta-glucosidase (cellobiase)/beta-glucosidase, oxidative cellulase/beta-glucosidase, xylanase/beta-glucosidase, hemicellulase/beta-glucosidase, lichenase/beta-glucosidase, chitenase/beta-glucosidase, xylanase/beta-glucosidase, cellulose phosphorylase/beta-glucosidase, endocellulase/oxidative cellulase, exocellulase/oxidative cellulase, beta-glucosidase (cellobiase)/oxidative cellulase, oxidative cellulase/oxidative cellulase, xylanase/oxidative cellulase, hemicellulase/oxidative cellulase, lichenase/oxidative cellulase, chitenase/oxidative cellulase, xylanase/oxidative cellulase, cellulose phosphorylase/oxidative cellulase, endocellulase/xylanase, exocellulase/xylanase, beta-glucosidase (cellobiase)/xylanase, oxidative cellulase/xylanase, xylanase/xylanase, hemicellulase/xylanase, lichenase/xylanase, chitenase/xylanase, xylanase/xylanase, cellulose phosphorylase/xylanase, endocellulase/hemicellulase, exocellulase/hemicellulase, beta-glucosidase (cellobiase)/hemicellulase, oxidative cellulase/hemicellulase, xylanase/hemicellulase, hemicellulase/hemicellulase, lichenase/hemicellulase, chitenase/hemicellulase, xylanase/hemicellulase, cellulose phosphorylase/hemicellulase, endocellulase/lichenase, exocellulase/lichenase, beta-glucosidase (cellobiase)/lichenase, oxidative cellulase/lichenase, xylanase/lichenase, hemicellulase/lichenase, lichenase/lichenase, chitenase/lichenase, xylanase/lichenase, cellulose phosphorylase/lichenase, endocellulase/chitenase, exocellulase/chitenase, beta-glucosidase (cellobiase)/chitenase, oxidative cellulase/chitenase, xylanase/chitenase, hemicellulase/chitenase, lichenase/chitenase, chitenase/chitenase, xylanase/chitenase, cellulose phosphorylase/chitenase, endocellulase/xylanase, exocellulase/xylanase, beta-glucosidase (cellobiase)/xylanase, oxidative cellulase/xylanase, xylanase/xylanase, hemicellulase/xylanase, lichenase/xylanase, chitenase/xylanase, xylanase/xylanase, cellulose phosphorylase/xylanase, endocellulase/cellulose phosphorylase, exocellulase/cellulose phosphorylase, beta-glucosidase (cellobiase)/cellulose phosphorylase, oxidative cellulase/cellulose phosphorylase, xylanase/cellulose phosphorylase, hemicellulase/cellulose phosphorylase, lichenase/cellulose phosphorylase, chitenase/cellulose phosphorylase, xylanase/cellulose phosphorylase, and cellulose phosphorylase/cellulose phosphorylase.
In certain embodiments the minicellulosome comprises at least three different cellulolytic (or other degredative) enzymes. In certain embodiments the three different enzymes comprise an enzyme pair selected from the group listed above, combined with one enzyme selected from the group consisting of an endocellulase, an exocellulase, a beta-glucosidase (cellobiase), an oxidative cellulase, a xylanase, a hemicellulase, a lichenase, a chitenase, a xylanase, and a cellulose phosphorylase.
It will be recognized that the enzymes, and enzyme combinations, identified above are intended to be illustrative and not limiting. Using the teachings provided herein, the display of numerous other enzymes will be available to one of skill in the art.
In various embodiments, to facilitate interaction of displayed enzyme(s) with their substrate (e.g., cellulose) the displayed protein comprises a substrate binding domain (e.g., a carbohydrate binding domain). Suitable substrate binding domains include, but are not limited to, carbohydrate binding domains, cellulose binding domains, cellulose binding modules, or other binding domains.
The amino acid sequence of cellulose binding peptides and/or binding domains are well known to those of skill in the art. Carbohydrate binding peptides include peptides e.g., proteins and domains (portions) thereof, that are capable of, binding to a plant derived cellulosic (e.g., lignocellulosic) material. Carbohydrate binding peptides include, for example, peptides screened for their cellulose binding activity out of a library, as well as naturally occurring cellulose binding peptides or peptide domains.
The carbohydrate binding domain can include any amino acid sequence expressible in plants which binds to a cellulose polymer. For example, the cellulose binding domain or protein can be derived from a cellulase, a binding domain of a cellulose binding protein or a protein screened for, and isolated from, a peptide library, or a protein designed and engineered to be capable of binding to cellulose or to saccharide units thereof. The cellulose binding domain or protein can be naturally occurring or synthetic. Suitable polysaccharidases from which a carbohydrate binding domain can be obtained includes, but is not limited to a β-1,4-glucanase. In certain embodiments, a cellulose binding domain or protein from a cellulase or scaffoldin is used.
Carbohydrate binding domains/modules are well known to those of skill in the art (see, e.g., Tomme et al. (1995) in Enzymatic Degradation of Insoluble Polysaccharides (Saddler and Penner, eds.), Cellulose-binding domains: classification and properties. pp. 142-163, American Chemical Society, Washington). Cellulose binding domains are also described in U.S. Pat. No. 5,837,814 and in U.S. Patent publication 2011/0005697 which are incorporated herein by reference for the cellulose binding domains described therein. In particular, U.S. Patent Publication No: 2011/0005697 identifies proteins containing putative β-1,3-glucan-binding domains (see, e.g., Table 1 therein, Table 4 below); proteins containing Streptococcal glucan-binding repeats (Cp1 superfamily) (see e.g., Table 2 therein, Table 5 below), and the like.
B. circulans (WL-12)
B. circulans (IAM 1165)
Actinomadura sp. (FC7)
Arthrobacter sp.
O. xanthineolytica
R. faecitabidus
R. communis
S. ividans (1326)
R. tridentatus
S. downei (sobrinus)
S. downei (sobrinus)
S. downei (sobrinus)
S. downei (sobrinus)
S. downei (sobrinus)
S. mutans (Ingbritt)
S. mutans (GS-5)
S. mutans (GS-5)
S. mutans
S. mutans (GS-5)
S. mutans (GS-5)
S. mutans (GS-5)
S. salivarius
S. salivarious
S. salivarious
S. salivarious
S. pneumoniae R6
S. pneumoniae
C. difficile (VPI
C. difficile (BARTS
C. difficile (VPI
C. difficile (1470)
C. novyi
C. novyi
C. acetobutylicum
C. acetobutylicum
C. acetobutylicum
C. acetobutylicum
In various embodiments the Ka. for binding of the carbohydrate binding domains/proteins to cellulose is at least in the range of weak antibody-antigen extractions, i.e., at least 103 M−1, preferably at least 104 M−1, most preferably at least 106 M−1.
In various embodiments the peptide comprising the cell wall sorting signal (CWS) also contains a secretory signal sequence to enhance/facilitate transport through the cell membrane. Typical Gram-positive secretory signal peptides are N-terminal peptides.
Gram-positive secretion signals are well known to those of skill in the art. In certain embodiments the secretory signal sequence comprises a B. subtilis phrC secretory signal or homologues thereof.
In various embodiments, the number and types of enzymes that can be displayed in each cellulosome (minicellulosome) can be increased by using multiple polypeptide fragments to construct a cell wall attached extended scaffoldin that in turn coordinates the binding of cellulases that are displayed on the cell's surface. This approach eliminates the need to express and display a single large scaffoldin polypeptide which can be problematic. The use of multiple polypeptide fragments to construct an “extended” scaffolidin provides a simple and effective way in which to expand the number of enzymes displayed on the surface of a Gram-positive bacterium (e.g., B. subtilis).
In certain illustrative, but non-limiting embodiments, the polypeptide fragments are expressed with “complementary” “linking scaffoldins and linking dockerins” that join the polypeptide fragment into an extended scaffoldin. Thus, for example a first polypeptide can be attached to the bacterial cell wall and bear a terminal “linking” (linker) cohesin or dockerin. This terminal linking cohesin or dockerin interacts with a corresponding cohesin or dockerin on a second polypeptide thereby providing an extended scaffoldin. Optional, the second polypeptide can bear a second linking cohesin or dockerin that interacts with a corresponding linker or dockerin on a third polypeptide fragment thereby facilitating the attachment of the third polypeptide to the second.
As proof of principle, the utility of this method is demonstrated herein in Example 2 by displaying a complex that contains six enzymes (instead of three enzymes). Cells containing the enlarged six enzyme complex have significantly improved cellulolytic activity. The new method can readily be used to construct larger complexes that contain more than six enzymes.
It is noted that the bacterial cells described herein do not require in vitro assembly of the complex. Thus, for example, no purified cellulases must be added to the cells to form the minicellulosome. It is also noted that it is believed the B. subtilis cells described herein (e.g., in Example 2) exhibit the highest cellulolytic activity of any recombinant microorganism yet reported.
The scaffold extension method described herein provides a simple approach to quickly increase the number of enzymes that are housed in multi-cellulase complexes displayed on the surface of a Gram-positive bacterium (e.g., B. subtilis). By using an extended scaffoldin approach, larger cellulase complexes can be assembled using smaller proteins. This is advantageous because it overcomes problems associated with secreting and folding of larger polypeptides. With this new system, Gram-positive bacteria (e.g., B. subtilis) can be engineered to display complexes that contain 4 or more, 5 or more, 6 or more, 7 or more, 8, or more, 9 or more, 10 or more, 11 or more, or 12 or more enzymes.
The new B. subtilis cells displaying the extended, six enzyme complex described herein in Example 2 exhibit improved cellulolytic activity and are therefore of greater potential use. Importantly, the method described herein is general, and can therefore be applied to construct cells that contain complexes that house more than six enzymes.
One illustrative, but non-limiting embodiments is shown in
Briefly, strain TDA21 was generated to co-express nine proteins: (1) the SrtA sortase from B. anthracis, (2) a chimeric scaffoldin (Scaf-I) composed of three type-I cohesin modules and one type-II cohesin module that is covalently attached to the cell wall by SrtA, (3) a second chimeric scaffoldin (Scaf-II) composed of three type-I cohesin modules and a type-II dockerin module, and (4-9) six dockerin-cellulase fusion proteins that bind to the scaffoldin non-covalently via species-specific dockerin-cohesin interactions (see, e.g., Table 7 in Example 2). The six cellulases were derived from C. cellulolyticum and C. thermocellum and have complementary cellulose degrading activities: Cel5A (endoglucanase/xylanase, family 5 glycoside hydrolase (GH)), Cel48F (processive endoglucanase, family 48 GH), Cel9E (exoglucanase, family 9 GH), CelS (exognlucanase, family 48 GH), Man5A (mannanase, family 5 GH), and XynA (xylanase, family 11 GH). Each protein component of the minicellulosome also contained an N-terminal signal sequence enabling them to be exported to the cell surface. The Scaf-I and Scaf-II proteins contain cohesin modules derived from C. cellulolyticum, C. thermocellum, and Ruminococcus flavefaciens that selectively bind to their cognate dockerin modules fused to Cel5A, Cel48F, Cel9E, CelS, Man5A, and XynA (
The construct described in Example 2 is intended to be illustrative and non-limiting. Using the methods described herein numerous other minicellulosomes comprising 4 or more, 5 or more, 6 or more, 7 or more, 8, or more, 9 or more, 10 or more, 11 or more, or 12 or more enzymes can be constructed and expressed on Gram-positive bacteria.
It has recently been noted that only twelve biomass-derived building blocks are needed to produce a range of commercial products. It is demonstrated that the protein display systems described herein can be used to create B. subtilis strains that can grow on plant biomass. Therefore, metabolic engineering of the cells described herein can create a consolidated bioprocessing microbe (CBP) that produces these biocommodities or building-blocks from cheap plant biomass. An immediate application is to introduce the cellulolytic system into existing microorganisms (e.g., bacteria) that are already used industrially to produce commercial products. This would enable the products to be produced from biomass and would significantly reduce costs.
Another illustrative, but non-limiting application of the systems described herein would be to use to use the biomass degrading cells as a replacement for enzyme cocktails that are currently used in industry to degrade biomass. The cells could be produced more cheaply than the enzymes and thereby reduce the costs associated with degrading biomass into its component sugars. An immediate application is lignocellulose degradation needed to produce lignocellulosic ethanol. Cells displaying the minicellulosomes would degrade the biomass into component sugars, which would then be used in downstream ethanol producing fermentation reactions performed by S cerevisiase.
The system we have developed can also be used to engineer other species of Gram-positive bacteria to convert them into CPB or microbes that are dedicated to degrading biomass into sugars.
In various embodiments it is contemplated that the display methods described herein can be used with virtually any microorganism capable of exploiting a sortase A transpeptidase reaction to anchor a protein to the cell surface. In various embodiments the microorganism is a Gram-positive microorganism (e.g., a Gram-positive bacterium).
The term “Gram-positive bacteria” generally refers to bacteria that are stained dark blue or violet by Gram staining Gram-positive microorganisms are well known to those of skill in the art. Gram-positive bacteria are generally divided into the Actinobacteria and the Firmicutes. The Actinobacteria or actinomycetes are a group of Gram-positive bacteria with high G+C ratio. They include some of the most common soil bacteria. Other Actinobacteria inhabit plants and animals and including some pathogens, such as Mycobacterium, Corynebacterium, Nocardia, Rhodococcus and a few species of Streptomyces. The majority of Firmicutes have Gram-positive cell wall structure. Illustrative Gram-positive bacteria include, but are not limited to Acetobacterium, Actinomyces (e.g., A. israelii), Arthrobacter, Bacillus (e.g., B. subtilis), Bifidobacterium, Clostridium, Clostridium spp. (e.g., C. perfringens, C. septicum, C. tetanomorphum), Corynebacterium, Enterococcus, Eubacterium, Frankia, Heliobacterium, Heliospirillum, Lactobacillus, Lactococcus, Leuconostoc, Listeria, Listeria spp., Megasphaera, Micrococcus spp., Micromonospora, Mycobacterium, Nocardia, Pectinatus, Pediococcus, Propionibacterium, Selenomonas, Sporomusa, Staphylococcus spp. (e.g., S. aureus), Streptococcus spp., (e.g., S. pneumoniae, B group streptococci), Streptomyces, and Zymophilus. Similarly, sortases, secretion signals cell wall sorting signals can include, but are not limited to, those derived from any Gram-positive microorganism.
In certain embodiments the bacterial host is selected from the group of non-pathogenic and/or non-invasive, Gram-positive bacteria consisting of Lactobacillus, Lactococcus, Pediococcus, Carnobacterium, Bifidobacterium, Oenococcus, Bacillus subtilis, Streptococcus thermophilus, and other non-pathogenic and/or non-invasive Gram-positive bacteria known in the art. In certain embodiments the bacterial host cell preferably is a Gram-positive bacterium, more preferably a Gram-positive bacterium that belongs to a genus selected from the group consisting of Lactobacillus, Lactococcus, Leuconostoc, Carnobacterium, Bifidobacterium, Bacillus, Streptococcus, Propionibacterium, Oenococcus, Pediococcus, Enterococcus. In certain embodiments the bacterial host cell is a bacterium that belongs to a species selected from the group consisting of L. acidophilus, L. amylovorus, L. bavaricus, L. brevis, L, caseii, L. crispatus, L. curvatus, L. delbrueckii, L. delbrueckii subsp. bulgaricus, L. fermentum, L. gallinarum, L. gasseri, L. helveticus, L. jensenii, L. johnsonii, L. minutis, L. murinus L. paracasei, L. plantarum, L. pontis, L. reuteri, L. sacei, L. salivarius, L. sanfrancisco, Lactobacillus ssp., C. piscicola, B. subtilis, Leuconostoc mesenteroides, Leuconoctoc lactis, Leuconostoc ssp, L. lactis subsp. lactis, L. lactis subsp. cremoris, Streptococcus thermophilus, B. bifidum, B. longum, B. infantis, B. breve, B. adolescente, B. animalis, B. gallinarum, B. magnum, and B. thermophilus.
As indicated above, in certain embodiments, microorganisms are engineered to contain a nucleic acid construct that exploits a sortase pathway to covalently anchor a protein to the surface of the cell. In certain embodiments the nucleic acid construct encodes a protein comprising one or more, preferably two or more cohesin domains attached to a secretory signal sequence (e.g., at the N-terminus of the protein) and a cell wall sorting signal (e.g., at the carboxyl terminus of the protein). In certain embodiments the same or additional constructs encode dockerin domains attached to a cellulolytic enzyme. The dockerin domains are selected to mate/bind with the cohesin domains on the “scaffoldin” protein. As described herein and illustrated in the examples, the entire system is designed to create a self-assembling minicellulosome.
In one illustrative, but non-limiting embodiment, a microorganism is transfected with the construct(s) and as encoded protein is transcribed it is displayed on the surface of the microorganism, e.g., through the transpeptidase reaction mediated by a sortase. The sortase can be an endogenous sortase expressed by the microorganism. In certain embodiments the sortase can be a sortase that is encoded by the same or another nucleic acid construct transfected into the microorganism. In certain embodiments the sortase is a sortase found in the subject microorganism, and in certain embodiments, the sortase is a sortase characteristic of a different microorganism.
In certain embodiments, particularly where a minicellulosome is to be expressed, the same construct or a different nucleic acid construct is provided that encodes one or more dockerins each attached to a different enzyme (e.g., cellulolytic enzyme) as described above.
Methods of making the nucleic acid constructs described herein are well known to those of skill in the art, and specific methods are illustrated in the examples. Cloning and bacterial transformation methods, DNA vectors and the use of regulatory sequences are well known to the skilled artisan and may for instance be found in Current Protocols in Molecular Biology, F. M. Ausubel et al, Wiley Interscience, 2004, incorporated herein by reference.
Many embodiments, of the methods and constructs described herein utilize an expression vector containing a nucleotide sequence that encodes the protein(s) of interest, a cell wall sorting signal and a secretion signal. Suitable expression vectors include, but are not limited to baculovirus vectors, bacteriophage vectors, plasmids, phagemids, cosmids, fosmids, bacterial artificial chromosomes, viral vectors (e.g. viral vectors based on vaccinia virus, poliovirus, adenovirus, adeno-associated virus, SV40, herpes simplex virus, and the like), P1-based artificial chromosomes, and any other vectors specific for specific hosts of interest. Such vectors include chromosomal, nonchromosomal and synthetic DNA sequences, and may comprise a full or mini transposon for the integration of a desired DNA sequence into the host chromosome. Examples of tranposons include but are not limited to TN5, TN7, and TN10, as well as the engineered transposomes from Epicentre (www.epicentre.com).
Numerous suitable expression vectors are known to those of skill in the art, and many are commercially available. The following vectors are provided by way of example; for bacterial host cells: pQE vectors (Qiagen), pBluescript plasmids, pNH vectors, lambda-ZAP vectors (Stratagene); pTrc99a, pKK223-3, pDR540, and pRIT2T (Pharmacia); for eukaryotic host cells: pXTI, pSGS (Stratagene), pSVK3, pBPV, pMSG, and pSVLSV40 (Pharmacia). However, any other plasmid or other vector, with or without various improvements for expression, may be used so long as it is compatible with the host cell.
In certain embodiments the subject vectors will contain a selectable marker gene. In certain embodiments this gene encodes a protein necessary for the survival or growth of transformed host cells grown in a selective culture medium. Host cells not transformed with the vector containing the selection gene will not survive in the culture medium. Typical selection genes encode proteins that (a) confer resistance to antibiotics or other toxins, e.g., ampicillin, neomycin, methotrexate, or tetracycline, (b) complement auxotrophic deficiencies, or (c) supply critical nutrients not available from complex media, e.g., the gene encoding D-alanine racemase for Bacilli, and the like.
The vector(s) of interest can be transfected into and propagated in the appropriate host. Methods for transfecting the host cells with the genomic DNA vector can be readily adapted from those procedures which are known in the art. For example, the vector can be introduced into the host cell by such techniques as the use of electroporation, precipitation with DEAE-Dextran or calcium phosphate, or lipofection.
Suitable promoters for use in prokaryotic host cells include, but are not limited to, a bacteriophage T7 RNA polymerase promoter; a trp promoter; a lac operon promoter; a hybrid promoter, e.g., a lac/tac hybrid promoter, a tac/trc hybrid promoter, a trp/lac promoter, a T7/lac promoter; a trc promoter; a tac promoter, and the like; an araBAD promoter; in vivo regulated promoters, such as an ssaG promoter or a related promoter (see, e.g., U.S. Patent Publication No. 2004/0131637), apagC promoter (Pulkkinen and Miller (1991) J. Bacteriol., 173 (1): 86-93; Alpuche-Aranda et al. (1992) Proc. Natl. Acad. Sci. U.S.A. 89(21): 10079-83), a nirB promoter (Harborne et al. (1992) Mol. Micro. 6: 2805-2813), and the like (see, e.g., Dunstan et al. (1999) Infect. Immun. 67: 5133-5141; McKelvie et al. (2004) Vaccine 22: 3243-3255; Chatfeld et al. (1992) Biotechnol. 10: 888-892, and the like); a sigma70 promoter, e.g., a consensus sigma70 promoter (see, e.g., GenBank Accession Nos. AX798980, AX798961, and AX798183); a stationary phase promoter, e.g., a dps promoter, a spy promoter, and the like; a promoter derived from the pathogenicity island SPI-2 (see, e.g., WO96/17951); an actA promoter (see, e.g., Shetron-Rama et al. (2002) Infect. Immun. 70:1087-1096); an rpsM promoter (see, e.g., Valdivia and Falkow (1996). Mol. Microbiol. 22:367-378); a tet promoter (see, e.g., Hillen, W. and Wissmann, A. (1989) In Saenger, W. and Heinemann, U. (eds), Topics in Molecular and Structural Biology, Protein-Nucleic Acid Interaction. Macmillan, London, UK, Vol. 10, pp. 143-162); an SP6 promoter (see, e.g., Melton et al. (1984) Nucl. Acids Res. 12:7035-7056); and the like.
In certain embodiments the nucleic acid constructs of interest are operably linked to an inducible promoter or to a constitutive promoter. Inducible and constitutive promoters are well known to those of skill in the art.
Where the host cell is genetically modified to express two or more gene products (e.g., a sortase and a protein comprising a sorting signal), nucleotide sequences encoding the two or more gene products will in some embodiments each be contained on separate expression vectors and in some embodiments contained in the same vector.
Where nucleotide sequences encoding the two or more gene products are contained in a single expression vector, in some embodiments, the nucleotide sequences will be operably linked to a common control element (e.g., a promoter), e.g., the common control element controls expression of all gene product-encoding nucleotide sequences on the single expression vector. In some embodiments, the nucleotide sequences encoding different gene products are operably linked to different control element(s) (e.g., promoter(s)). In some embodiments, one of the nucleotide sequences will be operably linked to an inducible promoter, and one or more of the other nucleotide sequences will be operably linked to a constitutive promoter.
As described above, the nucleic acid constructs may be introduced into the host cell as extra-chromosomal genetic materials that can replicate themselves (e.g., plasmids), or as genetic material integrated into the host genome. Regardless of whether the heterologous genes are integrated into the host genome, or exist as extra-chromosomal genetic materials, the optimal expression of the constructs heterologous genes belonging to a new metabolic pathway can on occasion benefit from coordinated expression of such genes, tight control over gene expression, and consistent expression in all kinds of host cells.
Methods and systems are provided that fine-tune the expression of heterologous genes, which in turn allow reproducible manipulation of metabolism in model microbes, such as E. coli, Bacillus subtillis, and Aspergillus nidulans. These methods allow balanced expression of the heterologous genes (e.g., those encoding the cellulosome) by techniques such as fine-tuning mRNA stability, the use of inducible promoters of various strengths, etc. See, for example, Keasling et al., New tools for metabolic engineering of E. coli. In Metabolic Engineering, S.-Y. Lee and E. T. Papoutsakis, eds. Marcel Dekker, New York, N.Y. (1999); Keasling, Gene-expression tools for the metabolic engineering of bacteria. Trends in Biotechnology 17:452-460, 1999; Martin et al., Redesigning cells for production of complex organic molecules. ASM News 68: 336-343, 2002 (all incorporated herein by reference).
While the foregoing discussion and examples below focus on Gram-positive bacteria and Sortase A, it will be appreciated that the methods described herein are amenable for use in any microorganism in which a sortase is found or can be expressed and is functional. Thus, for example, sortase enzymes have also been identified in the gram-negative organisms Bradyrhizobium japonicum, Colwellia psychroerythraea, Microbulbifer degradans, Shewanella oneidensis, and Shewanella putrefasciens, as well as in Methanobacterium thermoautotrophicum, a thermophilic archaeon (Pallen et al. (2003) Curr. Opin. Microbiol. 6: 519-527.). The use of the methods described herein with any of these organisms is also contemplated.
The foregoing methods and constructs are intended to be illustrative and not limiting. Using the teachings provided herein, numerous proteins, enzymes, minicellulosomes and the like can be stably displayed on the surface of a microorganism.
The following examples are offered to illustrate, but not to limit the claimed invention.
This example describes the engineering of B. subtilis to display a cell wall attached minicellulosome that assembles spontaneously. We show that these recombinant cells degrade both pretreated and untreated forms of lignocellulosic biomass, enabling them to grow robustly when these substances are provided as a primary nutrient source. This is an important step in the development of a CBP that can cost-efficiently convert biomass into valuable commodities.
Construction of B. Subtilis Strains.
A description of the strains and plasmids created in this study can be found in Table 6. The genes encoding srtA and scaf were integrated into the thrC locus by homologous recombination using the pSrtA/Scaf plasmid derived from vector pBL112 (Lanigan-Gerdes et al. (2007) Molecular Microbiol., 65: 1321-1333). Both genes are IPTG (Isopropyl-β-D-1-thiogalactopyranoside) inducible under the Pspac promoter. srtA encodes the B. anthracis sortase A and has been described previously (Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858). The scaf gene encodes a fusion protein that contains three type I cohesin modules derived from three different bacterial species: C. cellulolyticum (CipC), C. thermocellum (CipA), and R. flavefaciens (ScaB) (Fierobe et al. (2005) J. Biol. Chem., 280: 16325-16334). It also contains a family 3 carbohydrate binding module (CBM) from C. thermocellum CipA and the cell wall sorting signal (CWSS) from Staphylococcus aureus fibronectin binding protein B (Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858). The genes encoding the cellulase enzymes used in this study have been described previously, and were cloned into pHCMC05 (Bacillus Genetic Stock Center) to create plasmid pCellulase (Fierobe et al. (2005) J. Biol. Chem., 280: 16325-16334). Plasmid pCellulase contains genes encoding the three cellulase enzymes. cel9E encodes a fusion protein that contains a N-terminal VSV-g epitope tag, a CBM, immunoglobulin-like domain, a family 9 glycoside hydrolase (GH) domain, and the R. flavefaciens type I dockerin module. cel48F encodes an N-terminal Myc epitope tag, a family 48 GH, and a type I dockerin module from C. thermocellum. cel5A contains a family 5 GH with its native type I dockerin module and a C-terminal hexahistidine (His6) tag. In addition, a nucleotide sequence encoding a ribosome binding site and secretion signal derived from B. subtilis phrC was appended to scaf cel9E, cel48F, and cel5A. Similar methods were used to create plasmids pCel5A, pCel9E, pCel48F, pCel5A/9E, pCel5A/48F, and pCel9E/48F that contain one or two cellulase genes. Strains containing pSrtA/Scaf and/or plasmids encoding the cellulase genes were generated by transforming the plasmids into B. subtilis BAL2238 using standard methods, and involve plating on Luria-Bertani (LB) agar plates containing 1 μg/ml erythromycin or 5 μg/ml chloramphenicol (Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858).
Bacillus subtilis strains and plasmids used in this Example.
B. subtilis-E. coli shuttle plasmid that integrates into thrC
B. subtilis expression plasmid with IPTG inducible promoter,
B. anthracis srtA and scaf in pBL112
C. cellulolyticum cel5A in pHCMC05
C. cellulolyticum cel9E in pHCMC05
C. cellulolyticum cel48F in pHCMC05
C. cellulolyticum cel5A, C. cellulolyticum cel48F in
C. cellulolyticum cel5A, C. cellulolyticum cel9E in
C. cellulolyticum cel5A, C. cellulolyticum cel48F in
C. cellulolyticum cel5A, C. cellulolyticum cel9E,
C. cellulolyticum cel5A in pHCMC05
aProtein(s) expressed by the strains.
bFlag-tagged full length sortaseA from B. anthracis strain Ames
cScaf contains a three cohesin containing polypeptide (type I cohesins from C. thermocellum CipA, C. cellulolyticum CipC, and R flavefaciens ScaB) and a family 3 CMB that is anchored to the cell by SrtA vi the S. aureus fibronectin binding protein cell wall sorting signal.
dCel % A contains C. cellulolyticum Cel5A endoglucanase/xylanase fused to its native dockerin, an N-terminal secretory peptide, and a C-terminal His6-tag.
eCel9E contains the C. cellulolyticum Cel9E exoglucanase fused to an R. flavefaciens type-I dockerin, an N-terminal secretory peptide, and VSV-G epitope tag.
fCel48F contains C. cellulolyticum Cel48F processive endoglucanase fused to a C. thermocellum type I dockerin, and N-terminal secretory peptide, and Myc epitope tag.
gAmpr, ampicillin resistance; Eryr, erythromycin resistance; Cmr, chloramphenicol resistance.
h
Bacillus Genetic Stock Center.
Cell Fractionation and Immunoblot Analysis.
Cultures were grown overnight at 37° C. to saturation in 5 ml LB supplemented with 5 μg/ml chloramphenicol. A total of 500 μl of the overnight culture was then used to inoculate 50 ml of similar media and grown at 37° C. until the cells reached an optical density at 600 nm (OD600) of 0.1. At this point, IPTG was added to 1 mM to induce expression of the srtA, scaf, and cellulase genes. After the cells reached saturation, they were collected by centrifuging at 3,000×g for 10 min, washed with 1 ml STM buffer (50 mM Tris-HCl, pH 8.0, 25% sucrose, 5 mM MgCl2), centrifuged at 3,000×g for 5 min, and then re-suspended in STM such that the cell densities between samples were identical (OD600 ˜10). The cells were then fractionated by incubating with lysozyme (500 μg/ml) for 30 min at 37° C. to solubilize the cell walls. The suspensions were then centrifuged at 20,000×g to pellet the protoplasts, and the supernatant, which contains solubilized cell wall components, was collected. Secreted proteins were also collected from the spent growth medium, which was filtered through a 0.2 μm filter to remove cells. The proteins in the medium were precipitated with 10% trichloroacetic acid, centrifuged, and the pellet re-dissolved in water for immunoblot analysis. Samples of the solubilized cell walls (equivalent to 2.5×104 cells) and precipitated secreted protein (equivalent to protein secreted by 2.5×104 cells) were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to polyvinylidene fluoride (PVDF) membranes. Standard procedures for immunoblot analysis were used as described previously (Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858). Anti-His6 (0.25 μg/ml dilution for 1 h, Abgent) and anti-Myc (1 μg/ml dilution for 1 h, Syd Labs) primary antibodies were used to probe for Cel5A and Cel48F, respectively, and visualized using horseradish peroxidase conjugated to a rabbit anti-mouse immunoglobulin G secondary antibody (1:50,000 dilution for 1 h, Sigma). Cel9E was visualized using an anti-VSV-g primary antibody (0.5 μg/ml dilution for 1 h, Acris Antibodies) and horseradish peroxidase conjugated to a goat anti-biotin secondary antibody (1:20,000 dilution for 1 h, Cell Signaling).
Biomass Growth Studies.
Untreated corn stover, switchgrass, and hatched wheat straw were ground, washed with deionized water, and dried in an oven at 100° C. For some assays, the corn stover was first pretreated using dilute sulfuric acid as described previously (Jensen et al. (2010) Bioresource Technol., 101: 2317-2325). Briefly, 90 ml of 0.8% sulfuric acid was incubated with 3 g ground corn stover. A laboratory autoclave was then utilized to heat the corn stover-sulfuric acid suspension at 120° C. for 15 min. Following heating, the suspension was neutralized by washing with deionized water and dried in a laboratory oven at 100° C. Strains were tested for their ability to grow on untreated and pretreated biomass. Colonies from agar plates were used to inoculate a 5 mL LB culture supplemented with 1 μg/ml erythromycin and/or 5 μg/ml chloramphenicol, in order to select for srtA/scaf integrants and cells containing plasmids encoding cellulase enzymes, respectively. After 8 hours of growth at 37° C., 100 μl of each culture was transferred into 5 ml of M9 medium that contained 0.5% w/v glucose (Zhang et al. (2011) Metabolic Engineering 13: 364-372). The media also contained 0.004% tryptophan, 0.004% phenylalanine, and 0.004% threonine as the parent strain is auxotrophic for these amino acids. After 16 hours of growth, 100 μl of each culture was used to inoculate a 5 ml culture that contained biomass as the sole carbon source. This media consists of M9 minimal media and 0.5% w/v treated/untreated biomass. In control experiments the biomass was replaced with 0.5% w/v glucose. To induce protein expression 1 mM IPTG was added immediately after inoculating the biomass-containing culture. The OD600 of the cultures were measured over a 72 hour period. In addition to monitoring the cell density (OD600), colony forming units (CFUs) of strains TDA17 and TDA18 cultured in the presence of glucose or biomass were determined by plating 100 μl of the 102-106 dilutions onto LB plates supplemented with 5 μg/ml chloramphenicol and the resultant colonies that grew counted. Cells assayed for growth include those capable of displaying three (strain TDA17), two (strains TDA14 (Cel9E+Cel48F), TDA15 (Cel5A+Cel9E) and TDA16 (Cel5A+Cel48F)), or one cellulase enzyme (strains TDA11 (Cel5A), TDA12 (Cel9E) and TDA13 (Cel48F)), or cells only capable of secreting the enzymes (strain TDA18). Growth assays were performed in triplicate and the standard deviation was used as an estimate of the error.
Cells induced for protein expression were grown to saturation in LB media as described above. They were then centrifuged at 3,000×g for 10 min, re-suspended in Assay buffer (20 mM Tris-acetate, pH 6.0, 1 mM CaCl2, 0.1% sodium azide), re-centrifuged, and the final cell pellet was re-suspended in Assay buffer. Lignocellulosic biomass was then added to the cell suspension such that there was a total of ˜15 mg of cell displayed cellulase enzymes per gram of biomass; 10 ml suspensions containing cells at an OD600 of 2.5 were incubated with 60 mg of biomass at 37° C. with shaking (Banerjee et al. (2010) Biotechnol. Bioengineer., 106: 707-720; Banerjee et al. (2010) Bioresource Technol., 101: 9097-9105). In some instances, exogenous β-glucosidase (Sigma) was added to the cell-biomass mix (1 mg/g biomass), and the amount of cell suspension used was correspondingly adjusted to maintain a ratio of ˜15 mg enzyme per g biomass. For the assays performed using commercially available cocktails, a mixture containing 13.5 mg of Ctec2 and 1.5 mg of Htec2 enzyme cocktails (Novozymes Inc.) per gram of biomass was shaken in 10 ml of assay buffer at 37° C. To measure the amount of total biomass degraded, the cell-biomass mixture was removed at various times from the shaker and the insoluble biomass was allowed to settle. After decanting the cell, the residual biomass was washed with deionized water, decanted again, and the insoluble fraction dried (100° C. for 1 h). Measurement of reducing sugars released into the medium was accomplished as described previously and made use of dinitrosalicylic acid (glucose was used as the standard) (Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858; Tsai et al. (2009) Appl. Environ. Microbiol., 75: 6087-6093). Glucose was assessed using a glucose assay kit (Eton Biosciences) that makes use of the glucose oxidase enzyme and followed procedures outlined by the manufacturer (Banerjee et al. (2010) Biotechnol. Bioengineer., 106: 707-720; Banerjee et al. (2010) Bioresource Technol., 101: 9097-9105). Xylose release was analyzed using phloroglucinol (Fisher) as described previously (Eberts et al. (1979) Clinical Chem., 25: 1440-1443; Akin and Rigsby (2008) Appl. Biochem. Biotechnol., 144: 59-68). Control experiments made use of strain TDA18 which lacks scaf and srtA, but contains the cellulase expressing plasmid resulting in cellulase secretion. Assays were performed in triplicate and the standard deviation was used as an estimate of the error.
Determination of Surface-Displayed Scaf Levels and the Degree of Saturation.
For these studies full-length Cel5A, Cel9E, and Cel48F were overexpressed in E. coli and purified. Plasmids expressing each protein were created by sub-cloning their genes into plasmid pET28a using standard methods. After transformation into E. coli strain BL21(DE3), the proteins were over-expressed using standard procedures described by Novagen. For protein purification, cell pellets were re-suspended in lysis buffer (25 mM Tris-Cl, pH 7.0, 250 mM NaCl, 25 mM CaCl2), lysed by sonication, and the supernatant collected by centrifugation at 20,000×g for 30 min. The supernatant was then passed through Co-NTA resin, washed with 10 column volumes of lysis buffer supplemented with 10 mM imidazole, and eluted in lysis buffer supplemented with 100 mM imidazole. The proteins were then buffer exchanged into binding buffer (20 mM Tris-Cl pH 6.0, 2 mM CaCl2), concentrated, and the amount of protein quantified using a BCA assay.
To estimate the amount of Scaf displayed per cell, purified Cel5A was added to a known number of cells that display Scaf attached to the cell wall (strain TDA10). The amount of Cel5A adhered to Scaf was then estimated by measuring the cellulolytic activity of the cells. Briefly, a 50 ml LB culture of strain TDA10 supplemented with 1 μg/ml erythromycin was grown to an OD600 of 0.1, and IPTG was added to a final concentration of 1 mM to induce SrtA and Scaf expression. After 4 hrs, the cells were collected by centrifugation (3,000×g for 5 minutes), and re-suspended in binding buffer. To ensure that non-covalently bound Scaf was removed from the cells, this wash step was repeated. Increasing amounts of purified Cel5A was added to 2.4×1010 cells and incubated on ice for 1 hr. They were then centrifuged at 3,000×g for 5 minutes, the supernatant was removed, and the cells were re-suspended in 1 ml of binding buffer. This washing step was performed twice. The cells were then pelleted by centrifugation, the supernatant was removed, and the cells were re-suspended in 1 ml 0.5% CMC (dissolved in assay buffer). The amount of reducing sugar produced was then determined using DNS as described above. Control experiments were performed using strain TDA19, which does not produce SrtA needed to attach Scaf to the cell wall.
To determine if the cohesin domains within Scaf are saturated with enzymes TDA17 cells displaying minicellulosomes were exposed to purified enzymes and an immunoblot was performed to determine if they could bind additional protein. A 50 ml culture of cells was induced to express the minicellulosome as described above. The cells were collected by centrifugation and the pellet re-suspended in binding buffer. This procedure was repeated to wash the cells. A total of 5×1010 cells were then incubated with an excess of Cel5A, Cel9E, and Cel48F; to the cells, 2 mg of each purified enzyme was added. After incubation on ice for 1 hr the cells were then fractionated and subjected to immunoblot analysis as described above. Additional control experiments were performed using strains TDA10, TDA18 and TDA19 instead of TDA17.
B. subtilis Cells Display a Self-Assembled Minicellulosome.
Cells that display ex vivo assembled minicellulosomes are of questionable practicality for industrial applications; thus, we engineered B. subtilis to display a minicellulosome that self-assembles (
Cell fractionation and immunoblotting experiments were performed to confirm that the enzyme components self-assemble to form a minicellulosome on the cell surface (
Experiments were performed to quantify the number of displayed complexes and to determine if the Scaf proteins were saturated with enzymes. Data in
Degradation and Growth on Dilute Acid Pretreated Corn Stover.
Recombinant B. subtilis displaying the full-complement of enzymes in its minicellulosome (strain TDA17) grew when cultured aerobically in minimal media containing 0.5% w/v dilute acid pretreated corn stover as the sole carbon source (
To quantitatively determine the amount of dilute acid pretreated corn stover degraded by the cells, we exposed the biomass to TDA17 cells that were defective in sugar import. TDA17 cells induced to express the minicellulosome were grown to saturation in rich media, killed by adding azide (0.1%), washed, and incubated with pretreated biomass.
Measurement of Sugars Released from Dilute Acid Pretreated Corn Stover.
The amount of soluble reducing sugars (as well as glucose and xylose) liberated from dilute acid pretreated corn stover by azide-treated TDA17 cells was measured to further characterize their capacity to degrade biomass. As shown in
Corn stover is comprised of ˜36% glucose and ˜21% xylose, which reside within its cellulose and hemicellulose components, respectively (46). An analysis of the sugar content of the biomass before and after exposure to dilute acid revealed that pretreatment solubilized only a small fraction of the available sugars; 2% and 12% of the glucose and xylose were solubilized by dilute acid pretreatment, respectively (data not shown). After 48 hours the cells liberated 5% and 33% of the total available glucose and xylose in the biomass, respectively. Interestingly, the cells released ˜4 times more xylose than glucose, even though the pretreated biomass is primarily glucan (compare
Growth on Untreated Corn Stover, Switchgrass, or Hatched Straw.
We cultured the minicellulosome displaying cells with various types of untreated biomass to determine whether thermochemical pretreatment was required for degradation. Cells grew to saturation in ˜60 hours when cultured in minimal media containing untreated corn stover, switchgrass, or hatched straw (
The production of ethanol and other commodities from sustainable biomass promises to reduce the world's dependency on petroleum. A major obstacle to its commercialization is the high cost of degrading biomass into fermentable sugars, which is typically achieved industrially through a two-step process in which the biomass is first thermochemically pretreated before it is degraded by adding cellulase enzymes (Himmel et al. (2007) Science 315: 804-807; Hendriks and Zeeman (2009) Bioresource Technol., 100: 10-18; Yeoman et al. (2010) Adv. Appl. Microbiol. 70: 1-55). In principle, major reductions in cost and gains in efficiency could be achieved by using bacteria to degrade the biomass instead of enzyme cocktails. One promising approach to achieve this objective is to create cellulolytic microbes that display multi-cellulase containing minicellulosomes. This has now been accomplished in S. cerevisiae and B. subtilis (Wilson (2011) Curr. Opin. Microbiol., 14: 259-263; Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858; You et al. (2012) Appl. Environ. Microbiol., 78: 1437-1444; Tsai et al. (2009) Appl. Environ. Microbiol., 75: 6087-6093). While these microbes are capable of degrading amorphous purified cellulose and soluble cellulose (e.g., CMC), in some ex vivo assembly is required making them impractical for industrial purposes and it remains unknown whether they can grow using bona fide biomass as a primary nutrient source. Moreover, their cellulolytic activity has not been rigorously investigated. Here we report the construction of B. subtilis cells that display a minicellulosome that assembles without experimenter intervention. We demonstrate that the cells can degrade untreated biomass and use it as a nutrient source to grow.
To construct cells that display a self-assembled minicellulosome that could degrade biomass, we substantially redesigned our sortase-mediated protein display system, which we have shown is capable of displaying ex vivo assembled minicellulosomes on the surface B. subtilis (Anderson et al. (2011) Appl. Environ. Microbiol., 77: 4849-4858). Two major changes were made. First, we reengineered the cells to co-express all four components of the minicellulosome (previously, only the scaffoldin was expressed). This was achieved by expressing five genes: the sortase from B. anthracis (SrtA), a chimeric scaffoldin (Scaf) composed of three cohesin modules, and three dockerin-cellulase fusion proteins (Cel5A, Cel9E, and Cel48F) (
The self-assembled minicellulosomes enabled B. subtilis to grow robustly when dilute acid pretreated biomass was provided as a nutrient. Cells cultured with dilute acid pretreated corn stover approached similar CFU/mL values as those grown in soluble glucose (
Significantly, the minicellulosome displaying cells grew on three industrially relevant forms of biomass that did not require pretreatment with dilute acid: corn stover, hatched straw, and switchgrass. In all cases, after a significant lag, the cells achieved densities similar to those grown on glucose, and growth required that the cells display the enzymes on their surface (
Cellulase mixtures used in industry to degrade biomass contain as many as sixty distinct enzymes and can completely hydrolyze the cellulosic and hemicellulosic components of pretreated lignocellulose within 24 to 48 hours (Banerjee et al. (2010) Biotechnol. Bioengineer., 106: 707-720). In order to benchmark our recombinant cells against these enzyme mixtures, we quantified the amount of sugar released from both untreated and dilute acid pretreated corn stover following exposure to azide-killed cells. In these studies, the conditions were chosen such that 15 mg of total cellulase enzymes were exposed to 1 g of biomass. This cellulase:biomass ratio is identical to that used by Walton and colleagues to study biomass degradation using enzyme mixtures and assumes that ˜150,000 minicellulosomes are displayed on each cell. This number was calculated by measuring the cellulolytic activity of Cel5A that has been bound to cells displaying Scaf. Consistent with the growth data, after washing, only cells that displayed a minicellulosome released significant amounts of oligosaccharides from both untreated and dilute acid pretreated corn stover. Moreover, the cellulolytic activity of the azide-killed cells was stable for at least 48 hours (
An analysis of the sugar content of dilute acid pretreated corn stover before and after cell exposure indicates that 21% of total glucan and 33% of the xylan is digested into its component monosaccharides by the cells (
Recently, several model organisms have been engineered to grow on pretreated biomass. However, to the best of our knowledge, the B. subtilis strain reported herein is the first recombinant bacterium that has been demonstrated to have the ability to grow on untreated biomass. While native strains of B. subtilis can potentially subsist on untreated plant biomass, the laboratory strains created in this study could only grow on untreated biomass when functional minicellulosomes were displayed. The robustness of our recombinant B. subtilis cells was likely due to the sortase-mediated attachment system that allowed high copy-number display of the minicellulosome without the need for ex vivo assembly. In addition, unlike other previously described systems, the minicellulosomes are covalently anchored to the peptidoglycan, and thus presumably more stable and better suited for industrial applications (Lilly et al. (2009) FEMS Yeast Res., 9: 1236-1249; You et al. (2012) Appl. Environ. Microbiol., 78: 1437-1444; Tsai et al. (2009) Appl. Environ. Microbiol., 75: 6087-6093). Biofuels and many other high-value bio-based chemicals and materials can be produced from only twelve biomass-derived building blocks (Reddy and Yang (2005) Trends in Bbiotechnology 23: 22-27; Werpy et al. (2004) Top Value Added Chemicals From Biomass. Volume 1-Results of Screening for Potential Candidates From Sugars and Synthesis Gas. DTIC Institution). B. subtilis shows great promise for producing several of these compounds, since unlike many other currently used industrial microbes, it naturally imports and metabolizes cellobiose and C5 sugars (Tobisch et al. (1997) J. Bacteriol., 179: 496-506; Stulke and Hillen (2000) Ann. Rev. Microbiol., 54: 849-880). Moreover, using its robust genetic system, several investigators have already introduced into B. subtilis the relevant metabolic pathways needed to produce some of these compounds (Tobisch et al. (1997) J. Bacteriol., 179: 496-506; Schallmey et al. (2004) Canadian J. Microbiol., 50: 1-17; Li et al. (2011) Appl. Microbiol. Biotechnol., 91: 577-589; Xue and Ahring (2011) Appl. Environ. Microbiol., 77: 2399-2405; Romero et al. (2007) Environ. Microbiol., 73: 5190-5198), which, when paired with the cellulose degrading system we have created, enables the direct production of many valuable biocommodities from biomass.
Dwindling supplies of petroleum have encouraged the production of renewable biofuels from lignocellulosic biomass (Kerr (2008) Science, 322: 1178-1179). However, a significant problem deterring the enhanced use of fuels derived from lignocellulose is the cost of the cellulase enzyme cocktails that are used to degrade the plant biomass into fermentable sugars (Banerjee et al. (2010) Biotechnol. Bioeng. 106: 707-720; Banerjee et al. (2010) Bioresour. Technol. 101: 9097-9105). An alternative to the use of purified cocktails to hydrolyze lignocellulose includes the construction of recombinant cellulolytic microbes that display cellulase enzymes (la Grange et al. (2010) Appl. Microbiol. Biotechnol. 87: 1195-1208). Microbes that display multi-cellulase complexes generally experience high degrees of synergy between the displayed enzymes and can effectively enable the microorganism to adhere to the biomass which make them an attractive replacement of the cellulase cocktails (Fontes and Gilbert (2010) Ann. Rev. Biochem., 79: 655-681). Due to the promising applications of cellulosomes in industrial lignocellulose degradation, several research groups have displayed designer cellulosomes on the surface of Saccharomyces cerevisiae and Bacillus subtilis (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876; Anderson et al. (2011) Appl. Environ. Microbiol. 77: 4849-4858; Fan et al. (2012) Proc. Natl. Acad. Sci. USA, 109: 13260-13265; Goyal et al. (2011) Microb. Cell Fact., 10: 89; Kim et al. (2013) Microb. Cell. Fact. 12: 14; Tsai et al. (2009) Appl. Environ. Microbiol. 75: 6087 ˜6093; Tsai et al. (2010) Appl. Environ. Microbiol. 76: 7514-7520; Wen et al. (2010) Appl. Environ. Microbiol. 76: 1251-1260; You et al. (2012) Appl. Environ. Microbiol. 78: 1437-1444).
Though displaying the Clostridium cellulolyticum cellulases Cel5A, Cel9E and Cel48F in a designer cellulosome on the B. subtilis cell surface enabled these cells to efficiently degrade lignocellulose, the total amounts of reducing sugars released were three to six fold lower than commercially available cocktails, indicating that certain enzyme activities could be missing or that the absolute number of enzymes was too low (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876). This hypothesis can be supported by the notion that essential activities such as xylanases were missing in the surface displayed cellulosome and that the commercial cellulase cocktails contain up to eighty different enzymes, while the cell-displayed designer cellulosomes consisted of only three (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876; Banerjee et al. (2010) Biotechnol. Bioeng. 106: 707-720; Banerjee et al. (2010) Bioresour. Technol. 101: 9097-9105). Therefore, it became apparent that perhaps both of these issues could be resolved by increasing the number of enzymes displayed whose activities are different, yet complementary, to what is already present.
The choice of cellulase and hemicellulase enzymes to incorporate into the designer cellulosome is critical for obtaining optimal lignocellulose degradation. Work has been published using proteomics approaches to determine what cellulases are most prominently displayed in naturally assembled cellulosomes (Blouzard et al. (2010) Proteomics, 10: 541-554; Fendri et al. (2009) FEBS J., 276: 3076-3086; Raman et al. (2009) PloS One 4: e5271). Some of these studies were performed by culturing the bacteria on model substrates such as cellobiose, regenerated amorphous cellulose (RAC), and phosphoric acid swollen cellulose (PASC) (Blouzard et al. (2010) Proteomics, 10: 541-554). After culturing, the bacteria were collected and the purified cellulosomes subjected to mass spectrometry to identify and characterize the abundance of each protein found within the cellulosome. At least two dozen enzymes were found to be prominent in the isolated cellulosomes, but because the bacteria were not cultured on lignocellulosic substrates, these proteomic profiles may not necessarily reflect the composition of the complex when they are cultured on plant-based biomass. Therefore, in order to determine how the cellulosomal composition of the C. thermocellum and C. cellulolyticum cellulosomes changes when exposed to industrially relevant forms of biomass, these bacteria were cultured on switchgrass and straw, respectively (Fendri et al. (2009) FEBSJ., 276: 3076-3086; Raman et al. (2009) PloS One 4: e5271). Interestingly, it was observed that glycoside hydrolase (GH) family 9 and 48 enzymes appear to be particularly enriched in cultures that have been grown on pretreated lignocellulose (Fendri et al. (2009) FEBSJ., 276: 3076-3086). This may be due to a possible enhanced synergy between enzymes of these families. In addition, it has been observed that purified designer cellulosomes containing combinations of the C. cellulolyticum enzymes Cel9G paired with Cel9E or Cel48F experience at least a seven fold enhancement on crystalline cellulose (Fierobe et al. (2001) J. Biol. Chem. 276: 21257-21261; Fierobe et al. (2005) J. Biol. Chem. 280: 16325-16334; Mingardon et al. (2007) Appl. Environ. Microbiol. 73: 7138-7149). Family5 GH enzymes were also highly abundant and included the Man5A mannanase that could be crucial for efficient degradation of the hemicellulose component (Fendri et al. (2009) FEES J., 276: 3076-3086; Raman et al. (2009) PloS One 4: e5271). The interesting conundrum with Man5A is that it has also been found to be abundant when the bacteria were cultured on crystalline cellulose as opposed to lignocellulose, though it is not clear why this may be happening (Raman et al. (2009) PloS One 4: e5271). This phenomenon has also been observed with other hemicellulase enzymes and demonstrates that a global view of cellulosome composition does not necessarily indicate why some enzymes are present (Fendri et al. (2009) FEES J., 276: 3076-3086; Raman et al. (2009) PloS One 4: e5271).
Based on inter alia results described above, lignocellulose degradation can potentially be improved by the expansion of the surface-displayed cellulosome to include more than three enzymes. One goal of this work is to expand the cellulosome to as many as nine enzymes in order to resemble the C. thermocellum cellulosome. To determine the feasibility of efficiently displaying a cellulosome containing nine different enzymes, a smaller, more manageable complex containing six different cellulases was constructed. In addition to Cel5A, Cel9E and Cel48F which have previously been displayed, the C. thermocellum enzymes XynA (endoxylanase), Man5A (mannanase) and CelS (exoglucanase) were chosen to be displayed based on proteomics data showing their high abundance in cellulosomes isolated from C. thermocellum cultured on switchgrass (Raman et al. (2009) PloS One 4: e5271). Strains displaying six unique enzymes proved to be more effective at degrading both dilute-acid pretreated and untreated lignocellulosic substrates. This increased efficiency enabled the cells to reach similar degradation capacity as purified cellulase cocktails and permitted faster growth on lignocellulosic substrates. This is an important demonstration in the feasibility of enlarging the cellulosome to display more than three enzymes, and can be used as a model for the creation of strains displaying even larger cellulosome complexes.
Construction of B. subtilis Strains.
Strains of B. subtilis were created that could assemble a six-enzyme containing designer cellulosome (
Bacillus subtilis strains and plasmids used in this study.
B. subtilis-E. coli shuttle plasmid that integrates into thrC
B. subtilis expression plasmid with IPTG inducible promoter,
B. subtilis expression plasmid with IPTG inducible promoter,
B. anthracis srtA and scaf in pBL112
B. anthracis srtA and scaf in pBL112
B. anthracis srtA, scaf-I, and scaf-II in pBL112
C. cellulolyticum cel5A, cel9E, cel48F in pHCMC05
C. thermocellum celS, man5A, xynA in pDG148
aProtein(s) expressed by the strains.
bFlag-tagged full length sortaseA from B. anthracis strain Ames
cScaf contains three type I cohesin domains from C. thermocellum, C. cellulolyticum, and R. flavefaciens, a CBM, and the sorting signal from Staphylococcus aureus fibronectin binding protein B and a HA epitope tag.
d
C. cellulolyticum Cel5A appended with a His6 tag, C. cellulolyticum Cel9E appended with a VSV-G epitope tag and C. cellulolyticum Cle48F appended with a Myc epitope tag.
eScaf was appended with a type-ll cohesin from C. thermocellum immediately before the sorting signal domain, creating Scaf-I. Scaf-ll was generated by removing the sorting signal domain and replacing it with a type-ll dockerin from C. thermocellum
f
C. thermocellum CelS, C. thermocellum Man5A, C. thermocellum XynA. All proteins were appended with a His6 tag.
gAmpr, ampicillin resistance; Eryr, erythromycin resistance; Cmr, chloramphenicol resistance; Neor, neomycin resistance.
h
Bacillus Genetic Stock Center
Cell Fractionation and Immunoblot Analysis of Self-Assembled Cellulosome.
B. subtilis cells were cultured overnight in LB media supplemented with 5 μg/ml chloramphenicol and 5 μg/ml neomycin. A 500 μl aliquot of the overnight culture was then used to inoculate 50 ml of fresh LB media supplemented with the same antibiotics. After the cultures reached an optical density at 600 nm (OD600) of 0.1, 1M IPTG was added to a final concentration of 1 mM to induce protein expression. After the cells have reached saturation, generally after 4 h, the cells were collected by centrifuging at 3,000×g for 5 min, and the cell pellet resuspended in 1 ml STM buffer (25% sucrose, 50 mM Tris-HCl pH 8.0, 5 mM MgCl2). Following resuspension, the cells were centrifuged at 3,000×g for 5 min, and the pellet again resuspended in STM buffer such that the cell densities between cultures was the same (OD600 ˜10). The cells were then subjected to lysozyme treatment by the addition of 500 μg/ml lysozyme, and the cell walls solubilized by incubation at 37° C. for 30 min. The suspension was then centrifuged at 20,000×g for 10 min and the solubilized cell walls (found in the supernatant) were collected and stored at −20° C. The solubilized proteins found within the cell wall fraction were then analyzed by immunoblot, and has been described elsewhere (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876).
Determination of the Levels of Scaf-I/Scaf-II Displayed on the Cell Surface.
For these studies, full-length Cel5A was expressed and purified from E. coli. Cel5A was subcloned into a pET28a plasmid. After transformation into E. coli BL21 (DE3) cells, the protein was expressed using standard procedures at 18° C. overnight. Following protein expression, the cell pellet was resuspended in lysis buffer (25 mM Tris-HCl, pH 7.0, 250 mM NaCl, 25 mM CaCl2) and sonicated; the supernatant was collected by centrifugation at 20,000×g for 30 min. The supernatant was then passed through HisPur (Pierce, Inc.) cobalt-nitrilotriacetic acid (Co-NTA) resin, washed with 10 column volumes of lysis buffer supplemented with 10 mM imidazole, and eluted in lysis buffer supplemented with 150 mM imidazole. The proteins were then buffer exchanged into binding buffer (20 mM Tris-HCl, pH 6.0, 2 mM CaCl2) and concentrated, and the amount of protein was quantified using a bicinchoninic acid (BCA) assay.
In order to determine the amount of Scaf-I and Scaf-II displayed on the cell surface, increasing amounts of purified Cel5A was incubated with cells expressing and displaying Scaf or Scaf-I and Scaf-II. To 50 ml of fresh LB media, an inoculant containing 500 μl of an overnight culture was added and the culture induced to express Scaf or Scaf-I/Scaf-II for 4 hr. After the cells reached saturation, they were collected by centrifugation at 3,000×g for 10 min. To prevent non-bound Scaf or Scaf-I/Scaf-II from interfering in the assay, the cells were resuspended in 1 ml binding buffer and centrifuged at 3,000×g for 5 min, and repeated for a total of three times. To a total of 2.4×1010 cells used for each assay, an increasing amount of Cel5A was added, and allowed to incubate with the cells. Following incubation on ice for 1 hr, the cells were collected by centrifugation at 3,000×g for 5 min, resuspended in 1 ml binding buffer and centrifuged again at 3,000×g for a total of three times. Following the third wash, the cells were resuspended in 1 ml 0.5% carboxymethyl cellulose (CMC) dissolved in assay buffer (20 mM Tris-Acetate pH 6.0, 2 mM CaCl2) and incubated at 37° C. for 1 hr. Following incubation, the suspension was centrifuged and the reducing sugars found in the supernatant analyzed with the dinitrosalicylic acid assay solution (1% dinitrosalicylic acid, 1% NaOH, 0.2 phenol, 0.1 NaSO3). The solution was boiled for 10 min and the amount of reducing sugars present determined by reading the absorbance at 575 nm. Glucose was used as a standard. Control strains in which SrtA was not present, preventing covalent attachment of the scaffoldin proteins, were used. The assays were performed in triplicate and the error represented is the standard deviation.
Studies of Bacterial Growth on Lignocellulosic Biomass.
Untreated corn stover, switchgrass, and hatched wheat straw were washed with deionized water and frozen at −20° C. For some assays, the corn stover was first pretreated using dilute sulfuric acid as described previously (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876; Jensen et al. (2010) Bioresource Technol., 101: 2317-2325). Following autoclaving, the suspension was neutralized by washing with deionized water and stored at −20° C. Strains displaying three and six cellulases (TDA17 and TDA22, respectively) were tested for their ability to grow on untreated and pretreated biomass. Colonies from agar plates were used to inoculate a 5 ml LB culture supplemented with 5 μg/ml chloramphenicol and/or 5 μg/ml neomycin in order to select for transformants that contain plasmid pCellulase and/or pSXM. After 8 h of growth at 37° C., 100 μl of each culture was transferred into 5 ml of M9 medium that contained 0.5% (wt/vol) glucose. The medium also contained 0.004% tryptophan, 0.004% phenylalanine, and 0.004% threonine, as the parent strain is auxotrophic for these amino acids. After 16 h of growth, 100 μl of each culture was used to inoculate a 5 ml culture that contained biomass as the sole carbon source. This medium consists of M9 minimal medium and 0.5% (wt/vol) pretreated/untreated biomass. In control experiments, the biomass was replaced with 0.5% (wt/vol) glucose. To induce protein expression, 1 mM IPTG was added immediately after inoculating the biomass-containing culture. The OD600s of the cultures were measured over a 72 to 96 h period. Control strains in which no cellulases were displayed served as negative controls. Growth assays were performed in triplicate, and the errors represented are the standard deviation.
Whole-Cell and Cellulase Cocktail Sugar Release Assays.
Cells were induced for protein expression and were grown to saturation in LB medium as described above. They were then centrifuged at 3,000×g for 10 min, resuspended in assay buffer (20 mM Tris-acetate, pH 6.0, 1 mM CaCl2, 0.1% sodium azide), recentrifuged, and the final cell pellet was resuspended in assay buffer. Lignocellulosic biomass was then added to the cell suspension such that there was a total of ˜15 mg of cell-displayed cellulase enzymes per gram of biomass; 10 ml suspensions containing cells at an OD600 of 2.5 were incubated with 50 mg of biomass at 37° C. with shaking. For the assays performed using commercially available cocktails, a mixture containing 13.5 mg of CTec2 and 1.5 mg of HTec2 enzyme cocktails (Novozymes Inc.) per gram of biomass was shaken in 10 ml of assay buffer at 37° C. To measure the amount of total biomass degraded, the cell-biomass mixture was removed at various times from the shaker and the insoluble biomass was allowed to settle. Measurement of reducing sugars released into the medium was accomplished as described previously and made use of dinitrosalicylic acid (glucose was used as the standard). Glucose was assessed using a glucose assay kit (Eton Biosciences) that makes use of the glucose oxidase enzyme, and the assay followed procedures outlined by the manufacturer. Xylose release was analyzed using phloroglucinol (Fisher) as described previously (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876). Control experiments made use of strain TDA19, which lacks scaf and srtA but contains the cellulase-expressing plasmids, resulting in cellulase secretion. Assays were performed in triplicate, and the standard deviation was used as an estimate of the error.
Dionex Analysis of Sugars Released by Cells Displaying Three or Six Enzymes and Cellulase Cocktails.
In order to identify the types of cellodextrins and xylodextrins solubilized by azide-killed cells displaying three or six enzymes and purified cellulase cocktails when degrading untreated corn stover, similar experiments were performed as described above. Samples from cells that had grown on the untreated corn stover were also collected in order to determine the degree of biomass solubilization during growth and the methods used are identical to those described above. After incubation of the untreated corn stover with cells displaying three or six enzymes or the cellulase cocktail, the samples were transferred to a glass culture tube and autoclaved on the liquid cycle for 10 min. Following autoclaving, the samples, including supernatant and insoluble material, was transferred to 15 ml plastic tubes and frozen at −80° C. Before shipping to our collaborator, Dr. Henri-Pierre Fierobe at CNRS in France, the tubes were placed in a styrofoam box and filled with dry ice for shipping.
The insoluble biomass was completely hydrolyzed using concentrated sulfuric acid and high temperature to release all remaining monosaccharides that were not solubilized by the cells or cellulase cocktail. To both the solubilized monosaccharides from the residual lignocellulose and the soluble carbohydrates released into the supernatant by the cell-displayed enzymes and cellulase cocktail, 200 μl samples were taken and analyzed using a Dionex PA-1 anion exchange HPLC column, and this procedure has been described elsewhere (Westereng et al. (2013) J. Chromatog. A 1271: 144-152; Widmer (2010) Biotechnol. Letts., 32: 435-438). The residual sugars and solubilized carbohydrates from three independent experiments were collected.
B. subtilis Cells Display Enlarged Six-Enzyme Containing Designer Cellulosomes.
Recombinant B. subtilis displaying three cellulase enzymes (strain TDA17) enabled effective growth of the cells when lignocellulose was provided as the sole carbon source (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876). However, these strains were more than three-fold less active than commercially available cellulase cocktails on dilute acid-pretreated corn stover, and had a significant lag phase when cultured on plant biomass as a nutrient. This indicated that the numbers of enzymes and/or enzyme activities were lacking, and to enable for more efficient degradation and growth on lignocellulose, the cellulosome was enlarged to include more enzymes with complementary activities. In addition to the enzymes previously displayed, three new C. thermocellum enzymes: CelS (exoglucanase), XynA (endoxylanases), and Man5A (mannanase) were also expressed and displayed (
Western blotting analysis of cells expressing all six cellulase enzymes and the two scaffoldin proteins (strain TDA 22) indicate that the enzymes can correctly incorporate into the cellulosome as cells that lack the Scaf-I and Scaf-II scaffoldins (TDA20) cannot anchor the cellulase proteins to the cell wall (
Cells of B. subtilis that displayed cellulosomes containing three enzymes averaged 150,000 complexes displayed per cell (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876). Similar experiments to determine the number of complexes displayed by strain TDA21 were performed, and nearly twice as much activity on CMC was observed, indicating that about twice as much Cel5A was displayed as compared to the strain displaying only a single scaffoldin (
B. subtilis Cells Displaying Six Enzymes Enable Enhanced Growth on Dilute Acid Pretreated Corn Stover and Untreated Lignocellulosic Biomass Substrates.
B. subtilis that displayed only Cel5A, Cel9E, and Cel48F were able to utilize dilute acid pretreated corn stover as a nutrient for growth. Though after 60 hours the cultures were able to reach similar densities as those cultured with glucose, a significant lag phase was observed (10-12 hours) before transition into exponential growth (Anderson et al. (2013) Appl. Environ. Microbiol. 79: 867-876). It was hypothesized that cells of strain TDA17 were initially unable to quickly degrade the lignocellulose into metabolizable soluble monosaccharides and dextrans, resulting in this lag phase, until enough cellulosome complexes were assembled that enabled more robust lignocellulose degradation. Increasing the number of enzymes would presumably improve the rate of degradation because the display of more enzymes should increase solubilization of the insoluble carbohydrates. Cells of strain TDA22 that displayed the six enzymes did show an increased growth rate, and the lag was reduced to ˜5 hours (
A similar pattern of growth was observed when cells displaying six cellulases were cultured on untreated corn stover, switchgrass and straw (
Determination of Soluble Glycans/Xylans Released by Cells Displaying Six Cellulases on Dilute Acid Pretreated Corn Stover and Untreated Lignocellulose Substrates.
In order to more fully characterize the cellulolytic capabilities of cells displaying six cellulases in their cellulosomes, solubilized reducing sugars, glucose and xylose were analyzed from lignocellulose digestions using azide-treated cells. The standard 15 mg cell-displayed cellulase protein/g biomass was maintained in order to compare against the three cellulase displaying strain (TDA17) and commercial cellulase cocktails. Experiments were performed as described previously, in which cultures of cells displaying three or six enzymes were induced to express the cellulosomes. After reaching saturation, the cultures were collected and incubated with dilute acid pretreated corn stover or untreated lignocellulose substrates and samples taken periodically for 48 hours. Cells displaying six cellulase enzymes were able to release more than two-fold more reducing sugars from pretreated corn stover than cells displaying three enzymes (
Enzymatic degradation of untreated lignocellulosic substrates was also markedly better for the cells that displayed six cellulases. Overall, released soluble reducing sugars increased three to four fold over the amount solubilized by cells displaying only three enzymes (
Though these results indicate that the rationally chosen enzymes to be displayed in the six-cellulase cellulosome enabled a dramatic increase in lignocellulose hydrolysis, the substrates were not completely degraded. This could be due to a buildup of product inhibitors, such as cellobiose (Demain et al. (2005) MMBR 69: 124-154) or indicate the utility of displaying more enzymes to fully degrade the biomass.
Dionex analysis of Carbohydrates Released from Untreated Corn Stover by Three and Six Enzyme Displaying Cells.
In order to rationally determine what additional enzyme activities are required to improve degradation of lignocellulose, samples of strain TDA17 and TDA22 were azide-treated and incubated with untreated corn stover as described above. In addition, these strains were cultured on untreated corn stover to characterize the efficiency of lignocellulose saccharification when the cells are growing on it. Corn stover was chosen due to its use in cellulosic biofuel production and it was left untreated to determine what additional enzyme activities are required to efficiently digest untreated biomass.
To ascertain the types of carbohydrates that are released by cells displaying designer cellulosomes containing three or six cellulase enzymes, these strains were cultured in rich medium, induced to express the cellulosomes, and collected after the cells have reached saturation. Following expression, the cells were collected, washed, and resuspended in buffer containing 5 g/L untreated corn stover and 1% azide to make the cells inert. Following a 48 hour digestion at 37° C., the samples were autoclaved to stop enzymatic degradation, and the residual insoluble material was separated from the supernatant. The remaining biomass not solubilized by the cells was then completely saccharified using concentrated sulfuric acid and analyzed by Dionex high performance liquid chromatography (HPLC). It was determined that the untreated corn stover alone (not digested by the cells) consisted of 59% glucose, 15% xylose, 1% arabinose, and undetectable amounts of galactose and mannose (data not shown). As shown in Table 8 when strain TDA17 was exposed to biomass, 35% of it could be liberated, including 10% of the cellulose, 39% xylose, and 10% arabinose. However, when the lignocellulose was exposed to strain TDA22, more than half could be solubilized, resulting in 35% of the available glucose to be released into the supernatant, nearly half of the xylan and a third of the arabinan. Interestingly, arabinose solubilization was not expected as no arabinase was present in the cellulosome, but could have been released due to the hemicellulase activity of XynA. In addition, cells displaying three or six enzymes that were grown on the lignocellulose were able to degrade significantly higher amounts of the untreated lignocellulose than azide-killed cells. The increased degradation could likely be due to the constant removal of product inhibitors like cellobiose which would not be possible when the cells have been rendered metabolically inert (Demain et al. (2005) MMBR 69: 124-154). Furthermore, cells of strain TDA22 that were azide-killed or growing on untreated corn stover appeared to be almost as effective at solubilizing lignocellulose as the Ctec2/Htec2 cellulase cocktail (Table 8). In particular, more hemicellulose was removed by cells displaying cellulosomes than the Ctec2/Htec2 cocktail and could indicate that the cocktail may be catered toward the hydrolysis of cellulose and not hemicellulose. However, since the cellulase cocktail can solubilize nearly fifty percent more of the cellulose, the surface displayed cellulosomes may need to have additional endo- or exoglucanases incorporated into the complex to more efficiently work on the cellulose.
aPercentages shown in parenthesis indicates prevalence in 5 g/L untreated corn stover sample.
Analysis of the types of carbohydrates solubilized by cells displaying three or six enzymes was also performed using Dionex HPLC in order to determine what additional enzyme activities may be required for better lignocellulose degradation. Overall, more glycans and xylans were released by cells displaying six enzymes (Tables 9 and 10). This is consistent with the amounts of carbohydrates that were remaining in the insoluble biomass pellet after saccharification by the cells. The data suggests that expansion of the cellulosome to include three new enzymes is essential for more efficient saccharification. However, a significant portion of the glycans and xylans solubilized by cells displaying three and six enzymes were long chain carbohydrates (greater than four sugars in length) and revealed that additional endoglucanases may be necessary to further degrade them. Interestingly, the Ctec2/Htec2 cocktail also had a significant amount of the solubilized long chain carbohydrates. This may indicate that product inhibition could indeed be partly responsible in the incomplete conversion of the carbohydrates to glucose when using azide-killed TDA22 cells or the Ctec2/Htec2 cocktail.
aNot detected.
Further improvement of the cellulosome display system could include enlarging the complex to display an additional three enzymes, bringing the total number displayed to nine. To exploit the synergy of the enzymes currently displayed, Cel9G, an endoglucanase from C. cellulolyticum, can be incorporated because it has been demonstrated to work well together with Cel9E and Cel48F; a six-fold increase in digestion on wheat straw was observed when Cel9G is part of a complex containing Cel9E or Cel48F (Fierobe et al. (2001) J. Biol. Chem. 276: 21257-21261; Fierobe et al. (2005) J. Biol. Chem. 280: 16325-16334). Incorporation of another endoglucanase, C. cellulolyticum Cel8C could potentially enhance cellulose degradation as well to further produce more ends within the glycan chains that can be acted upon the exoglucanases present (Belaich et al. (1997) J. Biotechnol., 57: 3-14; Fierobe et al. (1993) Eur. J. Biochem./FEBS, 217: 557-565). In addition, another xylanase incorporated into the cellulosome could be beneficial to complete the removal of the hemicellulose. Since the incorporation of one xylanase enabled the solubilization of nearly 50% of the xylan, two xylanases may be able to function synergistically to completely remove the xylan. Xylanase XynZ from C. thermocellum has been shown to be abundant when cells were grown on cellulose, indicating that it could be essential in efficient xylan degradation (Gold and Martin (2007) J. Bacteriol., 189: 6787-6795; Raman et al. (2009) PloS One 4: e5271). Alternatively, addition of ligninases into the nine-cellulase cellulosome could prove to be critical in enhancing lignocellulose degradation. If the lignin could be removed easily by using these enzymes, there should no longer be a barrier for the cellulases and hemicellulases to function optimally. Incorporation of these additional enzymes should permit enhanced lignocellulose decomposition, and could potentially cause cells displaying cellulosomes to overcome the degradative capacity of the Ctec2/Htec2 cocktail.
The ultimate goal of this work is to develop strains of bacteria that efficiently degrade lignocellulose into its component sugars. These strains could potentially replace purified cellulase cocktails and act as a “bag of enzymes”, with the distinct advantage of being easily recyclable between lignocellulose digestions. The B. subtilis strains that have been constructed demonstrate lignocellulose degradation that is comparable to that of the cellulase cocktails, on both untreated and pretreated biomass. The use of cells to degrade lignocellulose has many advantages over these cocktails. Most importantly, one would only have to maintain bacterial cultures as opposed to the production and purification of multiple cellulase enzymes. In addition, potential cost savings could be realized due to the easy recyclability of cells displaying cellulases; cells displaying cellulases have recently been demonstrated to be functional after five digestion cycles (Matano et al. (2012) Bioresour Technol. 135: 403-409).
Another potential application and long-term goal of this work is to create consolidated bioprocessing (CBP) microorganisms that can directly convert lignocellulose into useful biofuels and other high-value biocommodities. A CBP could also potentially dramatically reduce production costs by combining all steps into a single organism, reducing the necessity to first hydrolyze the lignocellulose into fermentable sugars and then feed to yeast for fermentation (Mazzoli et al. (2012) Trends Biotechnol., 30: 111-119; Olson et al. (2012) Curr. Opin. Biotechnol., 23: 396-405). Work described in this thesis has created strains of bacteria that can effectively utilize lignocellulose as a source for growth, suggesting that this type of system could be useful in the construction of a CBP organism. In addition, B. subtilis has been demonstrated to be able to produce ethanol and other biofuels from glucose (Romero et al. (2007) Appl. Environ. Microbiol. 73: 5190-5198). It therefore seems feasible that engineering metabolic pathways to produce isobutanol, or other biocommodities, could be integrated into the cellulolytic strains of B. subtilis created. This result would be interesting because only a small number of CBPs created have demonstrated the ability to produce biofuels directly from lignocellulosic substrates (Bokinsky et al. (2011) Proc. Natl. Acad. Sci. USA, 108: 19949-19954). For these reasons, the strains developed in this thesis will likely be further optimized to produce chemicals of great social value.
It is understood that the examples and embodiments described herein are for illustrative purposes only and that various modifications or changes in light thereof will be suggested to persons skilled in the art and are to be included within the spirit and purview of this application and scope of the appended claims. All publications, patents, and patent applications cited herein are hereby incorporated by reference in their entirety for all purposes.
This application claims priority to and benefit of U.S. Ser. No. 61/727,601, filed on Nov. 16, 2012, which is incorporated herein by reference in its entirety for all purposes.
This invention was made with Government support of Grant No: DE-FC02-02ER63421, awarded by the U.S. Department of Energy. The Government has certain rights in this invention.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US2013/070393 | 11/15/2013 | WO | 00 |
| Number | Date | Country | |
|---|---|---|---|
| 61727601 | Nov 2012 | US |
