Transcriptional Regulation of Plant Biomass Degradation and Carbohydrate Utilization Genes in the Extreme Thermophile Caldicellulosiruptor bescii

Abstract

ABSTRACT

Extremely thermophilic bacteria from the genus Caldicellulosiruptor can degrade polysaccharide components of plant cell walls and subsequently utilize the constituting mono- and oligosaccharides. Through metabolic engineering, ethanol and other industrially important end products can be produced. Previous experimental studies identified a variety of carbohydrate-active enzymes in model species Caldicellulosiruptor saccharolyticus and Caldicellulosiruptor bescii, while prior transcriptomic experiments identified their putative carbohydrate uptake transporters. We investigated the mechanisms of transcriptional regulation of carbohydrate utilization genes using a comparative genomics approach applied to 14 Caldicellulosiruptor species. The reconstruction of carbohydrate utilization regulatory network includes the predicted binding sites for 34 mostly local regulators and point to the regulatory mechanisms controlling expression of genes involved in degradation of plant biomass. The Rex and CggR regulons control the central glycolytic and primary redox reactions. The identified transcription factor binding sites and regulons were validated with transcriptomic and transcription start site experimental data for C. bescii grown on cellulose, cellobiose, glucose, xylan, and xylose. The XylR and XynR regulons control xylan-induced transcriptional response of genes involved in degradation of xylan and xylose utilization. The reconstructed regulons informed the carbohydrate utilization reconstruction analysis and improved functional annotations of 51 transporters and 11 catabolic enzymes. Using gene deletion, we confirmed that the shared ATPase component MsmK is essential for growth on oligo- and polysaccharides but not for the utilization of monosaccharides. By elucidating the carbohydrate utilization framework in C. bescii, strategies for metabolic engineering can be pursued to optimize yields of bio-based fuels and chemicals from lignocellulose.

IMPORTANCE To develop functional metabolic engineering platforms for nonmodel microorganisms, a comprehensive understanding of the physiological and metabolic characteristics is critical. Caldicellulosiruptor bescii and other species in this genus have untapped potential for conversion of unpretreated plant biomass into industrial fuels and chemicals. The highly interactive and complex machinery used by C. bescii to acquire and process complex carbohydrates contained in lignocellulose was elucidated here to complement related efforts to develop a metabolic engineering platform with this bacterium. Guided by the findings here, a clearer picture of how C. bescii natively drives carbohydrate utilization is provided and strategies to engineer this bacterium for optimal conversion of lignocellulose to commercial products emerge.

INTRODUCTION

Plant biomass contains carbohydrate-rich cell walls whose composition is highly diverse and differs significantly between plants. A limited number of monosaccharides comprise the majority of plant polysaccharides (1). d-Glucose is the major component of glucans, cellulose in particular, while d-xylose is the predominant component of hemicelluloses, such as xylan, glucuronoxylan, arabinoxylan, and glucuronoarabinoxylan. The latter four hemicelluloses also contain residues of l-arabinose, d-glucuronate, and more rarely, d-galactose. Mannans (glucomannan, galactomannan, and galactoglucomannan) are composed of d-mannose, d-glucose, and d-galactose (2). Pectins (homogalacturonans and rhamnogalacturonans), acting as binders between cellulose and hemicelluloses fibers, are composed of d-galacturonate and l-rhamnose residues with various branching side chains (3). Starches (amylose and amylopectin) function as glucose storage polysaccharides in many plants and contain linear or branching d-glucose chains with either α-1,4 or α-1,6 glycosidic linkages, often referred to as α-glucans.

Microbial conversion of plant biomass into biofuels involves extensive and highly diversified carbohydrate utilization (CU) pathways that usually involve three major components. First, degradation of plant cell wall polysaccharides requires an array of extracellular carbohydrate-active enzymes (CAZymes), including glycosyl hydrolases (GHs) with carbohydrate-binding modules (CBMs), and polysaccharide lyases (PLs). Second, the extracellular biomass-derived oligo- and monosaccharides are transported into the cytoplasm via specific uptake transporters. Third, the imported saccharides undergo intracellular processing via sets of enzymes organized into individual catabolic pathways. These ultimately yield central metabolites that are further degraded through major glycolytic pathways (glycolysis or pentose-phosphate pathway) and natively fermented to final products (e.g., acetate, lactate, or ethanol).

Extremely thermophilic, cellulolytic bacteria from the genus Caldicellulosiruptor possess extensive and highly diversified machinery for plant biomass degradation and carbohydrate utilization (4). Caldicellulosiruptor species are common in neutral pH terrestrial hot springs globally where they are the dominant inhabitants of these high-temperature (70 to 80°C) environments and are capable of growth on cellulose, xylan, pectin, and other complex glucan substrates (5). Recent availability of genomic sequences of 14 Caldicellulosiruptor species provides opportunities for comparative analyses of inventories of CAZymes and CU genes, thereby linking them to species with distinct physiological properties. The glucan degradation locus (GDL) in C. bescii encodes six multidomain GHs/CBMs (multifunctional cellulases), three PLs (pectate lyases and rhamnogalacturonan lyase), and several other cellulose-binding proteins. This arsenal of extracellular proteins participates in primary breakdown of a variety of complex polysaccharides, including cellulose, β-glucan, xylan, mannan, and pectin (6, 7). Other cellulolytic Caldicellulosiruptor species possess GDLs with highly variable GH/CBM domain contents, suggesting dynamic evolutionary adaptation to variable plant biomass composition (8). Despite a diversity of CAZymes found in the genus, representing at least 45 GH and 20 CBM protein families, only some of these proteins have been characterized (5). Also, many metabolic gaps remain in describing CU pathways in Caldicellulosiruptor spp., and the identity of many mono- and oligosaccharide transporters is still unknown.

Transcriptomic analysis provides an opportunity to identify missing enzymes and transporters of CU pathways in Caldicellulosiruptor species. Previous transcriptomic studies in C. bescii, C. saccharolyticus, and C. kronotskyensis identified groups of genes encoding GHs/CBMs and ABC transporters that are upregulated on plant biomass substrates cellulose and switchgrass (9). Furthermore, the transcriptional response of C. saccharolyticus to various monosaccharides (10) and polysaccharides, including cellulose, mannan, pectin, and xylan (11), revealed patterns of genes that are coordinately regulated in response to specific carbohydrates. However, the specific transcription factors (TFs) and regulatory mechanisms for the CU gene network in Caldicellulosiruptor species remain unexplored. The global redox-sensing regulator Rex is the only experimentally characterized TF; it was shown to regulate genes encoding a membrane-bound [NiFe]-hydrogenase (MBH), and cytosolic electron-bifurcating [FeFe]-hydrogenase (BF-H₂ase) and pyruvate ferredoxin oxidoreductase (POR) in C. bescii and C. saccharolyticus (12, 13).

Accurate functional assignment of CU genes across diverse bacteria with complete genomes is challenging due to frequent variations of the respective pathways (e.g., nonorthologous gene displacements and functionally divergent paralogs). To enable accurate functional assignment of CU genes and inference of their transcriptional regulatory networks, we previously combined a subsystems-based comparative genomic approach (14, 15) with available experimental data to reconstruct CU metabolic pathways and transcriptional regulons across multiple taxonomic groups of bacteria, including the genera Thermotoga, Shewanella, Bacteroides, and Bifidobacterium (16–20), in addition to microbial communities (21). This approach substantially improved the accuracy of genomic annotations and the predicted functions of many previously uncharacterized CU genes in Thermotoga maritima (22–25), Shewanella spp. (26–29), and other bacterial species (30).

Here, this comparative genomics approach was used to reconstruct CU metabolic and regulatory networks in 14 available genomes from the genus Caldicellulosiruptor, with a specific emphasis on C. bescii. Comparative analysis of upstream regions of CU genes identified candidate DNA-binding motifs and enabled reconstruction of regulons for numerous TFs. Bioinformatics analyses complemented transcriptomics targeted to reveal transcriptional responses of C. bescii to cellulose, cellobiose, glucose, xylan, and xylose, which led to the identification of transcript boundaries for CU genes providing additional validation of the genomic inferences. As result, the reconstructed regulatory network of CU genes refined and improved a metabolic reconstruction describing how C. bescii degrades plant biomass and obtains carbon and energy for growth via cytoplasmic CU pathways. The refined CU pathway reconstructions were further used to construct the first metabolic model of C. bescii (31), thus opening opportunities for metabolic engineering of this species to produce bio-based chemicals from plant biomass.

RESULTS AND DISCUSSION

Genomic reconstruction of carbohydrate utilization pathways and regulons. (i) Glycosyl hydrolases.

To estimate the carbohydrate degradation capabilities of C. bescii, sets of CAZymes (GHs and PLs) that are involved in the breakdown of poly- and oligosaccharides into monosaccharides were identified. Overall, 56 proteins containing at least one GH domain were found in the C. bescii genome. These include six multidomain proteins containing two to five GH domains that also contain N-terminal signal peptides suggesting their extracellular localization. These secreted GHs are encoded within the glucan degradation locus (GDL), which plays an essential role in plant biomass deconstruction (6). Nine additional GHs, four PLs, and one polysaccharide deacetylase have signal peptides and thus are predicted to function as extracellular enzymes (see Data Set S1A in the supplemental material). Most of these secreted CAZymes (16 of 20) also contain one to three non-catalytic carbohydrate-binding modules (CBMs) that promote association of catalytic domains with polysaccharide. Using similarity searches and metabolic reconstructions via comparative genomics techniques, potential functions of 60 CAZymes that are presumably involved in biomass breakdown and carbohydrate utilization (CU) in C. bescii were examined. As a result, 57 CAZymes were assigned putative substrate specificity, Enzyme Commission (EC) number and CU metabolic pathway (see Data Set S1). These include 5 cytoplasmic and 11 secreted enzymes that were previously characterized in C. bescii, C. saccharolyticus, or C. owensensis. Functional roles in CU for the remaining 41 CAZymes were predicted, based on the genomic context analysis combined with metabolic reconstruction (see below) and by taking into account annotation of experimentally characterized homologs in other organisms (as summarized in Data Set S1A). In summary, the secreted CAZymes are involved in degradation of plant-derived polymers, including cellulose, mannans, hemicelluloses, pectin, and starches, while the cytoplasmic GH enzymes have broader functionalities since they are capable of catabolizing a variety of oligo- and disaccharides to release sugars that enter the central carbohydrate metabolism of C. bescii.

(ii) Carbohydrate utilization pathways.

To identify the repertoire of known and putative CU genes in C. bescii, a subsystem-based bioinformatics workflow for the analysis of 14 Caldicellulosiruptor genomes was generated. Functional annotation of the identified genes was conducted using genome context analysis, metabolic reconstruction, and homology searches. The complete list of CU genes with their assigned functional roles, CU pathways, protein families, and cellular localization, along with reference information on functionally characterized homologs, are provided in Data Set S1A. The identified CU machinery consists of 222 C. bescii genes, including 119 genes encoding catabolic enzymes, 68 genes for 21 carbohydrate transport systems, and 35 genes for DNA-binding transcription factors (TFs) and their cognate sensor kinases (Table 1). Functional annotations of 112 CU genes was supported by published literature, since they were homologous to characterized genes from Caldicellulosiruptor or other organisms (see Data Set S1A). Using the subsystem-based workflow, we inferred specific functional assignments for 94 genes whose functions were previously unknown or defined at the level of general class only. Most of these novel functional assignments constitute components of carbohydrate-specific ABC transporters (54 genes) and 29 transcriptional regulators. The ABC transporter systems so identified were classified into two large families, CUT1 and CUT2, that are implicated in uptake of oligo- and monosaccharides, respectively. Eleven novel cytoplasmic enzymes were implicated in the catabolic pathways for galacturonate, arabinose, rhamnose, and fucose (Fig. 1). Seventeen putative genes, including 5 CAZymes, a putative oligosaccharide ABC transporter (Athe_0085-87), a hypothetical CU gene locus (Athe_1993-97), and a putative dehydrogenase in the rhamnose utilization gene locus (Athe_0702-4), have not been assigned a specific function or substrate specificity. Overall, metabolic pathways for the degradation of the major plant polysaccharides, cellulose, xylan, pectin, glucomannan, and α-glucans (or starches) were identified, together with the uptake and intracellular catabolism of derived oligo- and monosaccharides (Table 1 and Fig. 1).

FIG 1

Reconstruction of metabolic pathways and regulons involved in plant polysaccharide degradation and carbohydrate utilization in C. bescii. Polysaccharide substrates are in red bent rectangles. Extracellular enzymes, transporters, and cytoplasmic enzymes are indicated by rectangles with red, blue, and black text, respectively. CAZymes and other classes of enzymes are indicated by large and small arrows, respectively. Novel carbohydrate-specific ABC transporters from the CUT1 and CUT2 families are in boxes with red and green outlines, respectively. The PTS transporter for fructose is in a blue outline box. The genes regulated by the same TF are indicated by matching colored symbols described in the lower inset. The names of differentially expressed (DE) genes on five carbohydrate substrates measured in RNA-Seq experiments are highlighted by light blue (cellulose), green (cellobiose), yellow (glucose), pink (xylan), and pastel blue (xylose). The central carbohydrate metabolism, fermentation, and hydrogen production pathways are indicated by a light yellow background. Detailed information on each gene from the reconstructed CU pathways, including gene IDs, functional roles, etc., is provided in Data Set S1A.

TABLE 1

Distribution of carbohydrate utilization genes in C. bescii^a

Sugar utilization pathway	Regulator(s)b	Transporter(s)c	No. of enzymes (no. secreted)	Total no. of genes	Total no. of genes in regulons
Monosaccharides	AxgR, GxgRS, AraR*	AxgFGHG1, GxgABCA1X		12	12
Xylan, xylose	XylR, XynR, XylQ, AxuRS, BxgRS	XynUVW, XloEFG, AxoFGH, AxuABC, BxgLFGL1	16 (6)	39	37
Arabinose, arabinose OS	AraR, ArbA, AxeRS	AxeUVW	5	12	9
Cellulose, cellobiose	AviRS	AviABCD	10 (7)	16	12
Pectin, galacturonate	UxaR, PecR, HemR, PecR2, KdgR*	PecXYZX, HemXYZ	15 (3)	26	20
Glucuronate, glucuronides	KdgR, XynR*		10	11	9
Fucose, fucose OS	FucR		3	4	4
Rhamnose, rhamnose OS, rhamnogalacturonides	RhaR, RhaR2	RhaXYZ, RglFGH	10 (1)	18	8
Galactose, galactose OS			7	7	-
Fructose	FruR	FruAB	3	6	6
Maltose, maltose OS	MalR, MalR1, MalR2	MalEFG, MalEFG2	9 (3)	18	13
Kojibiose, trehalose	KojR	KojEFG	3	7	6
Mannose, mannose OS	MosR, MosSQ	MosABC	9	15	7
Deoxynucleosides	DeoR	DeoUVWZ	6	11	11
Various oligosaccharides	XylR*	MsmKc		1	1
Other CU genes	IolR	Athe_0085-87	10	14	2
Hypothetical CU pathway oligosaccharides	Athe_1993	Athe_1997	3	5	5
Total CU genes	29 (35 genes)	21 (68 genes)	119	222	162
Glycolysis	CggR, Rex*		12	13	6
Central carbon metabolism	Rex		10	30	26

a

The details of all functional assignments summarized here are provided in Data Set S1.

b

An asterisk indicates a regulator that controls multiple metabolic pathways.

c

CUT1 family ABC transporters are underlined. MsmK is a common ATPase component shared by multiple of ABC transporters from the CUT1 family.

(iii) Carbohydrate utilization regulons.

Bacterial CU-specific regulators tend to cluster on the chromosome with their target genes (16, 18, 21, 32). The identified CU gene loci in C. bescii contain 28 genes encoding DNA-binding TFs and 6 genes encoding sensory histidine kinases that are essential components of two-component regulatory systems (TCRS), along with their cognate response regulators (see Data Set S1A). Most of the predicted TFs belong to the protein families whose members were previously known to regulate CU pathways, such as AraC, DeoR, GntR, IclR, LacI, ROK, and RpiR (Fig. 1). In contrast, ArsR constitutes a large family of repressors that collectively provide resistance to a wide range of both biologically required and toxic heavy-metal ions (33). Here, the first example of an ArsR-family TF (named ArbA) involved in CU was found that potentially controls the arabinoside catabolic gene abfA in Caldicellulosiruptor spp. The majority of identified in C. bescii CU-linked TFs lack previously characterized orthologs in any bacterial species and have only homologs with low sequence similarity (below 35 to 40% identity). This suggests that the biological function of these novel TFs (including specificities to effectors and their recognized DNA motifs) cannot be assigned based on homology.

To predict candidate TF binding site (TFBS) motifs for novel regulators, candidate coregulated genes were selected by genome context analysis in all Caldicellulosiruptor genomes (see Data Set S2) to which a phylogenetic footprinting approach to upstream regions of these genes was applied (see Fig. S1). After construction of a positional weight matrix for each identified DNA motif, additional TFBSs were sought in each analyzed genome possessing an orthologous TF gene, for which cross-species comparisons of the predicted regulons were done. As a result, DNA-binding motifs were predicted (Fig. 2), and regulons were reconstructed for 26 of 29 CU-related regulators in C. bescii that control a large transcriptional network of 162 genes involved in the cellulose, glucomannan, glucan, xylan, and pectin degradation and downstream utilization of the released oligo- and monosaccharides (Fig. 1). Below, the reconstructed regulons and catabolic pathways are described in more detail.

FIG 2

DNA-binding motifs for reconstructed carbohydrate utilization regulons. TFBS sequence logos were built by WebLogo using all candidate TFBSs identified by comparative genomics techniques in C. bescii and related Caldicellulosiruptor genomes. The identified TFBSs have either inverted-repeat (palindromes, represented as a single DNA element) or direct-repeat (tandems, represented by two identical DNA elements with a nonconserved linker of specified length) structures. Predicted repression or activation mechanism of regulation for each TF is specified by circled “R” and “A” letters, respectively.

(iv) Cellulose and mannan degradation.

The glucan degradation locus (GDL) is essential for plant biomass degradation and encodes six multidomain GHs/CBMs that catalyze the initial breakdown of β-glucans (such as cellulose) and mannans to generate an extracellular pool of cellulose oligosaccharides (COS) and mannose oligosaccharides (MOS), respectively (Fig. 1). These oligosaccharides are presumably delivered to the cytoplasm by CUT1-family ABC transporters, AviABCD and MosABC, predicted to be activated by local TCRS regulators AviRS and MosSQ, respectively (Fig. 3). The avi gene cluster contains an additional AviRS-regulated gene, celD, encoding a secreted bifunctional endoglucanase/endoxylanase. Cytoplasmic β-glucosidase BglA and two cellobiose phosphorylases (Cbp) required for COS utilization are encoded in a gene locus preceded by a new direct repeat DNA motif for an unknown TF (see Fig. S1). The mannan-derived MOS residues are degraded by cytoplasmic mannosidases (MnnA and MnnB) and phosphorylases (MosP and ManP). The novel LacI-family regulator MosR coregulates the MosABC transporter with the MOS phosphorylase MosP and cellobiose 2-epimerase MosE (Fig. 3), to convert 4-O-β-d-glucopyranosyl-d-mannose (epicellobiose) to cellobiose (34). As such, the mos gene cluster is involved in the utilization of oligosaccharides derived from degradation of mannan and glucomannan.

FIG 3

Genomic context of reconstructed regulons for catabolic and transporter genes involved in glucan, cellulose, mannan and pectin degradation in C. bescii. Carbohydrate utilization genes are indicated by arrows with colors according to the functional gene classification and numbers corresponding to their locus tag with Athe_ prefix. Candidate TF binding sites are indicated by colored symbols and connected to a cognate TF. Putative DNA-binding sites of as-yet-unknown TFs are indicated by circled numbers. Predicted effectors of TFs are given in square brackets. Transcription start sites (TSSs) identified by in RNA-Seq experiments are indicated by standing arrows. Genes upregulated on a particular substrate according to RNA-Seq and their corresponding log₂-fold change are shown in Fig. S3.

(v) Pectin degradation.

The last three genes in the GDL encode three extracellular pectate lyases—PecA, PecB, and PecC—that are involved in pectin degradation and are presumably coregulated by a novel regulator from the AraC family, which is encoded by the upstream gene pecR (Fig. 3). Degradation of polygalacturonan and rhamnogalacturonan, two major pectin polymers, results in the release of pectin oligosaccharides (POS) and rhamnogalacturonides, respectively. The derived oligosaccharides are taken up via three predicted CUT1-family ABC transporters and further metabolized using an array of cytoplasmic CAZymes and catabolic enzymes (Fig. 1) that are encoded by several gene clusters and presumably controlled by at least six local TFs (Fig. 3).

Unsaturated galacturonate is released from rhamnogalacturonate by RhiN and utilized via a 2-keto-3-deoxy-d-gluconate (KDG) intermediate via a dedicated pathway controlled by the KDG repressor KdgR from the IclR family. Saturated galacturonate is converted to tagaturonate via the isomerase UxaC; however, other enzymatic steps from conventional galacturonate catabolic pathways are missing in Caldicellulosiruptor spp. (24). The predicted regulator UxaR (DeoR family) coregulates two uncharacterized sugar catabolic gene clusters, hem and uxa (Fig. 3), encoding putative tagaturonate utilization enzymes (Fig. 1). Two predicted POS transporter operons, pecXYZ and hemXYZ, are likely controlled by local regulators from the AraC family, PecR2 and HemR, respectively. The predicted rhamnogalacturonide transporter RglFGH is coexpressed with secreted rhamnogalacturonan exolyase RglX. However, their genomic locus lacks a dedicated TF. Expression of the rhamnose catabolic genes rhaA, rhaB, and rhaD1 is possibly controlled by two DeoR-family paralogs, RhaR and RhaR2. l-Lactaldehyde, a toxic intermediate of rhamnose catabolism, is further metabolized to l-lactate by aldehyde dehydrogenase or to 1,2-propanediol by lactaldehyde reductase (30), although both enzymes are missing in Caldicellulosiruptor genomes. This final step of rhamnose catabolism is likely catalyzed by a novel three-subunit dehydrogenase, encoded in Athe_0702-4 from the RhaR2-regulated gene cluster, which is homologous to acetoin dehydrogenase.

(vi) Xylan degradation.

Xylose is a major component of β-xylans, while arabinoxylan (AX) and glucuronoxylan (GX) contain the side chains of arabinose and glucuronic acid, respectively. In the C. bescii genome, 10 multigene loci and four single gene loci encode CAZymes, transporters and regulators involved in xylan degradation, uptake and utilization of resulting oligosaccharides, and catabolism of xylose, arabinose, and glucuronate (Fig. 4). The ROK-family regulator XylR from C. bescii is 33% identical (amino acid level) to the previously characterized xylose-responsive repressor from T. maritima, which coregulates xylan degradation, XOS and xylose utilization genes (22). In contrast to T. maritima, the reconstructed XylR regulon in C. bescii contains only two target loci – the xylose isomerase xylA and the XOS utilization gene cluster xynUVW-xylR-xynA. The xylan degradation locus (XDL) (Athe-0174-Athe_0185), which encodes two CUT1-family ABC transporters (AxoFGE and XloEFG) and seven CAZymes (three secreted and four cytoplasmic), is presumably controlled by two TFs, a novel LacI-family regulator, named XynR, and the arabinose repressor, AraR. The remaining xylan and XOS catabolic gene loci, the xylose catabolism operon xynR-xylB, the glucuronidase aguX, and glucuronate catabolic genes uxuAB, are coregulated via the reconstructed XynR regulon (Fig. 4). The latter glucuronate utilization genes belong to a larger GX utilization gene cluster, which also encodes a CUT1-family ABC transporter (BxgLFG) and a TCRS regulator (BxgRS), that are presumably involved in utilization of glucuronoxylan oligosaccharides (GXOS) and locally controlled by BxgRS characterized by two conserved TFBSs with tandem repeat structure (see Fig. S1).

FIG 4

Genomic context of reconstructed regulons for catabolic and transporter genes involved in xylan degradation in C. bescii. For abbreviations, see the Fig. 3 legend.

The α-xyloside utilization gene locus, containing a CUT1-family ABC transporter (AxuABC) and cytoplasmic α-xylosidase XylS, encodes two novel TFs from the AraC-family (named AxuRS and XylQ), that are predicted to control axuABC and xylSQ genes, respectively (Fig. 4). Another gene cluster, containing a cytoplasmic endo-α-1,5-l-arabinanase (AbnA), a CUT1-family ABC transporter (AxeUVW), and a TCRS regulator (AxeRS), is likely involved in utilization of arabinoxylo-oligosaccharides (AXOS).

The AraR regulator from the GntR family is 42% identical (amino acid level) to arabinose-responsive repressors from B. subtilis and Clostridium acetobutylicum (35, 36). The reconstructed AraR regulon in C. bescii includes the arabinose catabolic operon araBRD and the novel predicted l-arabinose isomerase araI, the above-mentioned xylan degradation gene locus, the α-l-arabinofuranosidase abfA, and the axg operon encoding a CUT2-family ABC transporter for monosaccharides. The abfA gene in C. bescii and other Caldicellulosiruptor spp. is divergently transcribed with a hypothetical gene encoding novel ArsR-family regulator (ArbA) (Fig. 4). A common intergenic region of abfA and arbA contains a candidate AraR-binding site and three predicted ArbA-binding sites that are conserved across the genus (see Fig. S1). The AraR-controlled axg locus also encodes a candidate local regulator of the axgGHF operon (AxgR) that belongs to the DeoR family of TFs. The axgR-axgGHF intergenic region contains a putative AxgR regulatory site with tandem repeat structure (Fig. 2), which is conserved across all eight Caldicellulosiruptor genomes possessing the axg loci (see Fig. S1). Thus, both AraR and XynR TFs function as master regulators controlling five and six gene loci in C. bescii, respectively, while seven other identified TFs associated with the xylan degradation and utilization loci are presumably local.

(vii) α-Glucan degradation.

Linear and branching alpha-glucans, containing d-glucose monomer linked together through either α-1,4 or α-1,6 glycosidic linkages (i.e., amylose and amylopectin), are two major constituents of starch. Caldicellulosiruptor genomes contain a conserved set of extracellular hydrolases involved in the primary degradation of α-glucans (AmyA2, PulA, and AmyX), the CUT1-family ABC transporter MalEFG for uptake of maltooligosaccharides (MOS), and cytoplasmic CAZymes for MOS degradation (NplT, Cga, AmyA, MalL, and PulA2), resulting in maltose and glucose as the final products (Fig. 1). Maltose is further utilized via the GlgP phosphorylase. The LacI-family regulator MalR controls most of the genes and operons involved in α-glucan degradation (pulA-amyX), MOS uptake and catabolism (malFG-glgP-malR, malE, amyA, nplT, and cga). All of these genes are coregulated by a common palindromic DNA motif (Fig. 2; see also Fig. S1), which is similar to binding motifs of orthologous MalR repressors from Bacillus (37), Streptococcus and Lactobacillus (38), Bacteroides (17), and Bifidobacteria (16). An additional LacI-family regulator (KojR) presumably controls the Athe_0397-0403 gene cluster, containing cytoplasmic phosphorylases (pgmB-kojP and treP) and the CUT1-family ABC transporter (kojEFG) involved in the utilization of α-glucoside disaccharides trehalose and kojibiose.

Characterization of ATPase component for oligosaccharide transporters.

As shown above, Caldicellulosiruptor spp. possess a large number of candidate oligosaccharide transporters from the CUT1 family of ABC transporters (Fig. 1). These include 12 systems implicated in the uptake of oligosaccharides derived from the degradation of cellulose, glucomannan, pectin, and xylans, as well as transporters for maltose/MOS, the disaccharides trehalose and kojibiose, and deoxynucleosides (Table 1). All these CUT1-family transporters lack a dedicated ATP-binding ATPase component and presumably use a shared ATPase encoded by the msmK gene (Athe_1803). To test this hypothesis, we constructed the msmK deletion mutant strain of C. bescii (see Fig. S2) and characterized its growth on mono- and polysaccharides.

Under standard growth conditions on the monosaccharide substrates xylose and glucose, the ΔmsmK strain grew to the same final density as the parent strain (Fig. 5). On both substrates, while the same final cell density was reached, a minor growth phenotype was observed whereby the growth rate was lower during log phase than the parent strain. During growth of the parent strain on both glucose and xylose, maximum cell density was reached at 16 h, while the deletion strain did not reach maximum density until 20 h. Growth of the ΔmsmK strain was severely impaired relative to the parent strain on all tested di- and oligosaccharide substrates (Fig. 5). Substrates that supported diminished growth capacity were cellobiose, crystalline cellulose, xylan, and pectin. The growth phenotype was the least severe on cellobiose, where the final cell density was roughly 8-fold lower than the parent strain. During growth on cellulose and pectin, the final cell densities were over an order of magnitude lower than the parent strain. These growth phenotypes of the ΔmsmK strain of C. bescii suggest that MsmK is a functionally promiscuous ATPase, which is essential for utilization of di- and oligosaccharides derived from degradation of cellulose, pectin and xylan; however, MsmK is not required for monosaccharide uptake.

FIG 5

Comparison of growth for the ΔmsmK strain and the parent strain on various carbon substrates. The ΔmsmK strain and the parent strain were grown in biological triplicate (n = 3) at 75°C on a variety of soluble and insoluble carbohydrate substrates. Growth was compared on glucose (A), xylose (B), xylan (C), cellobiose (D), cellulose (E), and pectin (F). Insoluble xylan particles with similar scale to C. bescii cells interfered with direct quantification of growth (counting cells). Therefore, for growth on xylan, the growth was reported as the endpoint (28 h) protein concentration. For all other substrates, growth was reported as the cell density (cells/ml) over the course of growth (24 to 28 h).

Reconstruction of central carbohydrate metabolism and regulons.

The reconstructed carbohydrate catabolic pathways produce phosphorylated hexoses, pentoses, or trioses that are further catabolized through the central carbohydrate metabolism pathways (Fig. 1). A total of 18 genes encoding 16 enzymes from glycolytic and the pentose phosphate pathways are conserved in all analyzed Caldicellulosiruptor spp. (see Data Set S2). The central glycolytic gene repressor CggR controls the cggR-gap-(pgk-tpi)-pgm-eno operon encoding enzymes from the central part of glycolysis. In Bacillus subtilis, the CggR repressor binding to its DNA operator is modulated by fructose-1,6-bisphosphate (39). By analyzing the upstream gene regions of cggR genes, the putative CggR-binding motif, an inverted repeat with sequence GGGAC-(27)-GTCCC (see Fig. S1), can be identified that is similar to DNA motifs of CggR regulators from other Firmicutes, e.g., Bacillus spp. (37). In addition to the conventional glyceraldehyde-3-phosphate (GAP) dehydrogenase (encoded by the CggR-controlled gap gene), all known Caldicellulosiruptor species have an alternative ferredoxin-dependent GAP oxidoreductase (GOR) (40). GOR in C. bescii is encoded by the two-gene gorSL operon and controlled by a global redox-sensing regulator Rex that responds to the intracellular ratio of NADH:NAD⁺ (13). Among other experimentally determined Rex regulon members in C. bescii are pyruvate ferredoxin oxidoreductase (POR), ferredoxin-dependent membrane-bound [NiFe]-hydrogenase (MBH), and the bifurcating [FeFe]-hydrogenase (BF-H₂ase).

Comparative genomics was used to reconstruct the Rex regulon in the genomes of Caldicellulosiruptor species. The identified 20-bp palindromic motif was similar to previously identified TFBS motifs of Rex in other clostridial lineages (41). A positional-weight matrix was constructed to scan the 14 Caldicellulosiruptor genomes for additional candidate Rex-binding sites. After the removal of false-positive sites by a consistency check approach, the predicted members of Rex regulon in C. bescii and related species were identified. The most conserved members of the reconstructed Rex regulon were involved in central carbohydrate metabolism (GOR, POR, BF-H₂ase, and MBH), acetate fermentation (Pta and AckA) and serine metabolism (SerC and SerA) (Fig. 1; see also Data Set S4). Candidate Rex-binding sites upstream of these operons were conserved in all analyzed Caldicellulosiruptor genomes (see Fig. S1). Among additional Rex regulon members in C. bescii that were also conserved in most other species, a second copy of putative POR-encoding operon was found, an hypothetical operon encoding sulfur carrier protein TusA, CoA-disulfide reductase, an Fe-S oxidoreductase and phosphoglucosamine mutase.

Carbohydrate-specific transcriptomic responses of C. bescii.

To assess the transcriptional responses of C. bescii to glucose, cellobiose, cellulose, xylose, and xylan, transcriptome sequencing (RNA-Seq) analyses were done for mid to late exponentially growing cultures grown in triplicates on each of five carbon sources. Reads per samples ranged from 3.6 to 5.0 million with an average 4.2 million reads across the 15 samples (see Data Set S3A and B). Averaged normalized expression values on log₂ scale (see Data Set S3C) were compared between each pair of conditions to estimate number of differentially transcribed genes in these experiments (Table 2). Overall, 277 C. bescii genes were upregulated at least 4-fold (log₂-fold > 2) for at least one pair of conditions. Growth on polysaccharides cellulose and xylan promoted larger transcriptional responses (64 to 86 upregulated genes) compared to glucose, xylose, and cellobiose (10 to 25 upregulated genes). These upregulated genes were further classified into three groups (see Data Set S3D): (i) 98 genes (36 operons) that belong to the reconstructed CU pathways and TF regulons; (ii) 93 genes involved in other metabolic pathways, including amino acid and nucleotide metabolism (25 genes), peptidases (4 genes), and ABC efflux transporters (9 genes); and (iii) 87 hypothetical genes, including pseudogenes and transposases. The transcriptional responses of CU genes involved xylan, glucan, cellulose, mannan, and pectin degradation genes were placed into the context of the reconstructed TF regulons (Fig. 1 and Table 3; see also Fig. S3).

TABLE 2

Numbers of differentially transcribed genes for C. bescii growth on various carbohydrate substrates

Substrate	No. of differentially transcribed genesa
	Cellulose	Cellobiose	Glucose	Xylan	Xylose
Cellulose (Avi)		86 (49)	68 (35)	60 (17)	81 (31)
Cellobiose (CB)	10 (0)		18 (12)	23 (5)	31 (7)
Glucose (Glc)	25 (4)	23 (21)		30 (13)	43 (8)
Xylan (XN)	37 (15)	70 (50)	64 (42)		65 (39)
Xylose (Xyl)	14 (2)	23 (11)	22 (9)	10 (5)

a

The numbers represent differentially transcribed genes with a log₂-fold change of >2.0 for comparisons between column and corresponding line. The numbers in parentheses correspond to subsets of differentially expressed genes that are involved in the reconstructed carbohydrate utilization pathways/regulons. For example, 86 genes were upregulated on cellulose compared to cellobiose; of these, 49 genes belong to the reconstructed pathways/regulons. The functional classifications of the remaining non-CU genes are provided in Data Set S3D.

TABLE 3

Differential expression of carbohydrate utilization genes in C. bescii grown on carbohydrate substrates

Target operon(s)a	DE substrate(s)b	CU pathway(s)c	TF regulon(s)d
xynUVW, xylA	XN, Avi, Xyl	Xylan	XylR
xyl3A, xynM-xynN-xynC, xynA, X-xynA2	XN, Avi	Xylan	XynR
xynR-xylB	XN	Xylan	XynR
xloEFG-X-GH129-axoFGE-xynF-xynE-xynD-xyn10A-xynX-xynB	XN, Avi	Xylan (XDL)	XynR, AraR
aguX-uxuB-uxuA-bgaL	XN, Avi	Xylan, glucuronate	XynR
xylR	XN	Xylan
msmK	XN, Avi, CB	Oligosaccharides	XylR
bxgL-bxgFG	XN, Avi	Xylan	BxgRS
bxgL1-X	XN, Avi, Xyl, Glc	Xylan	BxgRS
axuABC	XN, Avi	Xylan, α-xylosides	AxuRS
celA1-celB1-X-celA2-celB2-celC; celF	Avi, XN	Cellulose (GDL)	New motifs 1/2
celD; aviABC	Avi, CB, XN, Xyl	Cellulose	AviRS
cbp-cbp2	Avi, CB, XN	Cellobiose	New motif 3
mosABC	Avi, CB, XN	Mannose OS	MosR, MosSQ
axgGHF	XN, Avi, Xyl, Glc	Monosaccharides	AxgR, AraR
gxgABC	Glc, XN, Xyl	Monosaccharides	GxgRS
pulA-amyX	Avi, CB, Glc	Starch	MalR
malE; malFG-glgP-malR; amyA; cga	Avi, CB, Glc	Maltodextrin OS	MalR
hemXYZ	XN	Pectin OS	HemR

a

CU operons with similar transcriptional response, assigned catabolic pathways, and TF regulons are grouped together and separated by semicolon. Detailed information on CU operon-containing genes including their locus tags and functional annotations is provided in Data Set S1. Additional differentially expressed genes involved in the utilization of mannose OS, pectin, fructose, inositol, and deoxynucleosides are listed in Data Set S3D and visualized in Fig. 1.

b

Five analyzed substrates: XN, xylan; Avi, crystalline cellulose; CB, cellobiose; Xyl, d-xylose; Glc, d-glucose. Substrates that demonstrated differential expression (DE) of genes from target operons with log₂-fold change of >2.0 are shown. Substrates and genes with moderate differential expression that showed log₂-fold change between 1.5 and 2 are underlined. Detailed information on differential gene expression values is provided in Data Set S3.

c

Metabolic scheme of the reconstructed CU pathways is given in Fig. 1.

d

TF regulons with predicted TFBSs located in the promoter regions of target operons are indicated. Putative TFBS motifs for yet unknown TFs are numbered as 1, 2, and 3. Consensus sequences for identified TFBS motifs are given in Fig. 2.

(i) Cellulose and mannan degradation.

Extracellular GHs encoded by the GDL locus are required for degradation of both β-glucans (e.g., cellulose) and mannans. The GDL genes were upregulated (4- to 7-fold) on cellulose and xylan (see Fig. S3). Despite the fact that this locus lacks a dedicated TF, the identified common DNA motifs in the celA1 and celF promoter regions (Fig. 2) suggest potential involvement of as-yet-unknown TFs to the observed transcriptional responses to both polysaccharides. The aviABC and celD genes from the COS utilization locus were upregulated on cellulose, cellobiose, and xylan (10- to 15-fold) and presumably controlled by their cognate TCRS regulator AviRS. The cbp-cbp2 operon encoding two cellobiose phosphorylase paralogs was ∼6-fold upregulated on cellulose, cellobiose, and xylan compared to glucose-grown cells and was associated with a novel direct repeat DNA motif for an as-yet-unknown TF (see Fig. S1). The mosABC operon was highly upregulated on cellulose and cellobiose (50- to 90-fold) and also moderately responded to xylan (9-fold), and the cognate TCRS MosSQ is presumably involved in the induction of this mannose OS transporter.

(ii) Xylan degradation.

Transcription levels of all 25 genes from the six operons that constitute the reconstructed XynR regulon were significantly elevated in C. bescii grown on xylan compared to cells grown on cellobiose. The highest fold changes of gene expression (∼10- to 100-fold) were observed for the xylan degradation genes xynA2, xyl3A, and xynMNC and the proximal XDL genes, while the distal XDL genes, as well as xynR-xynB and aguX-uxuBA-bgaL, were moderately upregulated on xylan (4- to 9-fold). In addition, most of the XynR-regulated genes demonstrated modest upregulation on cellulose. The xylUVW and xylA genes from the XylR regulon were highly upregulated on xylan, xylose, and cellulose. A low-scoring XylR-binding site was identified upstream of msmK encoding an ATPase component of CUT1-family transporters (see Fig. S1). msmK is highly expressed in all tested conditions, with only 3- to 4-fold higher expression in C. bescii grown on xylan and cellulose compared to glucose-grown cells (see Fig. S3), suggesting the XylR repressor has a weak effect on its expression. Also highly upregulated on xylan were bxg and axu encoding putative GX oligosaccharides and α-xyloside transporters that are presumably regulated by their local TCRS, BxgRS and AxuRS, respectively.

(iii) Monosaccharide transporters.

C. bescii grows on various monosaccharides, including fructose, glucose, galactose, mannose, xylose, arabinose, fucose, and rhamnose (42, 43). Fructose uptake and utilization are mediated via the dedicated phosphotransferase system (PTS) FruAB-PtsI-Hpr and phosphofructokinase FruK that are controlled by the DeoR-family repressor FruR. Two candidate ABC transporters from the CUT2 family, AxgFGH and GxgABC, are potentially involved in uptake of other monosaccharides in C. bescii and are presumably controlled by their local transcription factors, the AraC-family regulator AxgR and the TCRS regulator GxgRS, respectively. Each of these systems includes a dedicated ATPase (GxgA and AxgG), which agrees with experiments showing that the MsmK ATPase is not essential for monosaccharide uptake. The proposed sugar specificities of these two monosaccharides transporters can be predicted using transcriptomics data. In C. bescii, both axgFGH and gxgABC genes are upregulated on glucose (10- to 25-fold) and xylan (5- to 10-fold), although their basal levels of expression and responses to other carbohydrates were different (see Data Set S3). While the axgFGH genes were highly upregulated on xylose (30- to 100-fold) and cellulose (15- to 30-fold), the gxgABC operon was highly expressed on all carbon sources, with only 3-fold induction on xylose compared to cellobiose-grown cells (minimal expression level). In C. saccharolyticus (10), an ortholog of the axgFGH transporter was induced on arabinose, xylose, and galactose, while the gxgABC ortholog was upregulated on galactose, xylose, and glucose (see Data Set S1B). These results suggest that the cognate AxgR and GxgRS regulators for these monosaccharide transporter operons respond to different carbohydrates.

(iv) Starch and maltodextrin degradation.

The starch degradation operon pulA-amyX, the maltodextrin OS transporter malEFG and cytoplasmic enzymes amyA and cga were moderately upregulated on cellulose, cellobiose and glucose compared to xylan and xylose grown cells (3- to 10-fold). MalR is a predicted global regulator of all these starch/maltodextrin utilization genes that can be partially responsible for their similar transcriptional responses.

5′-Transcription start site mapping for CU genes.

We used 5′-end RNA sequencing for all five C. bescii cultures from RNA-Seq experiments to globally determine the 5′ transcription start sites (5′TSSs) at nucleotide resolution. As result, 5′TSSs were mapped within the upstream regions of 83 of 100 putative operons containing CU genes, as well as for 18 putative operons encoding enzymes from the central carbohydrate metabolism, including all predicted Rex- and CggR-regulated operons (see Data Set S2). We further performed multiple alignments of the 5′TSS-containing gene upstream regions from C. bescii and related Caldicellulosiruptor species and searched for associated candidate −35 and −10 promoter elements (see Fig. S1). The majority of identified 5′TSSs were located within 180 bp upstream of the translation start site (median distance 50 bp). However, the fucI-alfA operon has the largest 5′-untranslated region (5′ UTR) between the transcription and translation start sites (329 bp), containing tandem FucR-binding sites, suggesting FucR acts as a repressor.

To determine whether TFs in CU from the reconstructed regulons were activators or repressors, the positions of their predicted TFBSs were compared to the experimentally mapped 5′TSSs in C. bescii (see Fig. S1). All 12 TFs from the LacI and DeoR families, as well as XylR (ROK family), ArbR (ArsR family), AraR (GntR family), KdgR (IclR family), and CggR (SorC family), have their cognate TFBSs located within 5′ UTRs or overlapping candidate promoter elements, suggesting that they are repressors (Fig. 2). This is in agreement with the previously reported repression mechanism for other TFs from the protein families discussed above (16, 18, 22, 30, 32, 33, 44). In contrast, all four single-component regulators, as well as three of five TCRS containing DNA-binding domains from the AraC family, have their cognate DNA sites (tandem repeats with large spacers) located upstream of candidate 5′TSS-associated promoter elements, suggesting that they are activators. In fact, most characterized regulators in this family (44), including a novel TCRS for xyloglucan degradation in Ruminiclostridium cellulolyticum (45), function as transcriptional activators.

Many global regulators of carbohydrate metabolism in bacteria, including Rex, are capable of acting as both repressors or activators, depending on the location of their binding sites with respect to the promoter elements of their target genes (27, 41, 46). For the majority of core conserved Rex regulon members, Rex-binding sites either overlap with—or are located downstream of—the promoter elements associated with 5′TSSs, suggesting that Rex functions as a transcriptional repressor of these operons (see Fig. S1). However, the MBH-encoding operon has a Rex operator located 45 bp upstream of its 5′TSSs, suggesting a potential activation mechanism. The hypothesis that Rex is an activator of MBH is supported by decreased MBH transcript expression, as previously determined in an RNA-Seq analysis of a rex deletion strain of C. bescii (13). Thus, these results suggest that the NADH-responsive Rex regulator controls expression of two hydrogenases in an opposite manner: namely, Rex represses the NADH-dependent cytoplasmic BF-H₂ase and activates the membrane-bound and NADH-independent MBH, suggesting an adaptation mechanism to fine-tune the NADH concentration in the cell.

Conclusions.

Extremely thermophilic cellulolytic bacteria from the genus Caldicellulosiruptor possess extensive and highly diversified CU machinery. C. bescii is a promising nonmodel species for development of a microbial metabolic engineering platform for conversion of plant biomass to next-generation bioproducts. Accurate functional assignment of CU genes across diverse bacteria with complete genomes is challenging due to frequent variations of the respective pathways. Here, a subsystems-based comparative genomic approach was used in conjunction with available experimental data on genome-wide gene expression to reconstruct CU pathways and transcriptional regulons in C. bescii. The global CU model so developed includes >150 protein families (metabolic enzymes, transporters and transcriptional regulators) spanning >20 distinct pathways. Using a comparative analysis of 14 Caldicellulosiruptor genomes, DNA-binding motifs and reconstructed CU regulatory networks were identified, including both previously known regulons that have orthologous TFs characterized in other Firmicutes (e.g., AraR, KdgR, and XylR), as well as ab initio predicted novel TF regulons from LacI, DeoR, AraC, and other families. Reconstructed regulons provided an additional layer of genome context, helping to significantly improve the accuracy of functional annotations and metabolic reconstruction. For instance, the inferred AraR regulon includes a novel functional variant of an arabinose isomerase gene, which is nonorthologous to previously characterized enzymes. The reconstructed regulatory network was validated by gene expression profiling via RNA-Seq analysis of C. bescii grown on glucose, cellobiose, cellulose, xylose, and xylan. In addition, the TSS data for C. bescii were used to delineate the transcriptional architecture of CU genes and to propose the activation or repression mechanism of regulation for predicted TF regulons. Results of previous gene expression experiments in C. bescii and the related C. saccharolyticus species grown on various mono- and polysaccharides were also considered. The observed upregulation of genes involved in catabolism of specific carbohydrates showed excellent correlation with the reconstructed content of transcriptional regulons.

Among other insights, it was determined that, while the CUT2 family ABC transporters (putatively responsible for monosaccharide transport) are each chromosomally associated with a dedicated ATPase subunit, the CUT1 family transporters each lack an associated ATPase. A similar scenario has been previously observed in Streptococcus pneumoniae, where msmK was characterized as a shared ATPase for six different CUT1 family ATP transporters that each lacked their own ATPase (47). Likewise, in B. subtilis, a homolog (msmX) was demonstrated to be a multipurpose ATPase involved in the uptake of pectin oligosaccharides (48). Deletion of msmK in C. bescii resulted in the predicted phenotypes on all tested carbohydrate substrates, indicating that this ATPase is indeed functionally promiscuous.

This study elucidated the transcriptional regulatory network and mechanisms controlling expression of CU genes in C. bescii, thereby improving the functional annotations of carbohydrate transporters and catabolic enzymes to inform a metabolic model of C. bescii (31). The developed C. bescii model containing the targeted reconstruction of CU pathways supports metabolic engineering strategies for this biotechnology-important thermophilic bacterium capable of unpretreated lignocellulose conversion to bioproducts.

MATERIALS AND METHODS

Bioinformatics analysis of Caldicellulosiruptor genomes. (i) Genomes and bioinformatics resources for functional annotation.

The following 14 Caldicellulosiruptor genomes were downloaded from NCBI Reference Sequence (RefSeq) database: C. bescii DSM 6725 (NC_012034), C. kronotskyensis 2002 (NC_014720), C. hydrothermalis 108 (NC_014652), C. owensensis OL (NC_014657), C. obsidiansis OB473, C. danielii Wai35.B1 (NZ_LACM00000000), C. acetigenus DSM 7040 (NZ_ATXO00000000), C. kristjanssonii 177R1B (NC_014721), C. lactoaceticus 6A (NC_015949), C. saccharolyticus DSM 8903 (NC_009437), Caldicellulosiruptor sp. F32 (NZ_APGP01000000), C. changbaiensis CBS-Z (NZ_CP034791), C. morganii Rt8.B8 (NZ_LACO00000000), and Thermoanaerobacter cellulolyticum NA10 (renamed C. naganoensis NA10, NZ_LACN00000000). All Caldicellulosiruptor genomes were functionally annotated using the Rapid Annotation of microbial genomes using Subsystem Technology (RAST) server (49) and were imported to the SEED genomic database to enable comparative genomic analysis and metabolic reconstruction using metabolic subsystems (collections of functionally related proteins) (15). The complete RAST-annotated proteomes with SEED IDs for protein-encoding genes were used for identification orthologs between 14 Caldicellulosiruptor strains using a pangenome analysis pipeline based on the identification of bidirectional best hits (50). Genome-wide CAZyme family annotations were assigned using dbCAN server (51), cellular localization of CAZymes was assigned using SignalP and PSORTb (52, 53).

(ii) Reconstruction of CU metabolic pathways.

For identification of candidate CU genes in the C. bescii genome and their functional annotation, we used the previously described bioinformatics workflow (19) and several reference databases including KEGG for reference metabolic pathways and biochemical reactions (54), Pfam for Protein Families and Gene Ontology terms (55), TCDB for Transporter Classification (56), TransportDB for genome-wide transporter prediction (57), CAZy for Carbohydrate Active enZYmes (58). For the majority of candidate CU genes, we also performed protein similarity searches using NCBI BLAST versus UniProt Knowledgebase (UniProtKB), a high-quality curated resource on functional information of experimentally characterized proteins (59), and PaperBLAST, a unique and regularly updated database of proteins described or mentioned in published scientific articles obtained by automated text-mining (60). We further used the subsystem-based comparative genomic approach implemented in the SEED genomic platform and database that capture the taxonomically diverse functional variants of CU enzymes and transporters (15) to analyze the genomic and functional context of CU gene candidates in C. bescii. The obtained functional gene annotations are captured in Data Set S1 and summarized in Table 1.

(iii) Ab initio reconstruction of TF regulons.

For ab initio prediction and reconstruction of CU regulons, we utilized the established comparative genomics approach (initially described elsewhere [61] and also reviewed in reference 14) that was previously utilized for de novo inference of TF binding site motifs and regulon analysis for numerous TFs implicated in the transcriptional regulation of various CU pathways in diverse taxonomic groups of bacteria (16–19, 21, 32, 37). First, we identified genes encoding potential CU-related TFs by analyzing all representatives of five TF protein families, namely, AraC, DeoR, GntR, LacI, ROK, and RpiR, that are most commonly implicated in CU transcriptional networks in bacteria according to the RegPrecise database (44). For each candidate TF and its orthologs, we analyzed their genomic context across the analyzed Caldicellulosiruptor species and, thus excluded putative TFs that do not show the conserved chromosomal clustering with CU genes in the genomes. After the initial assignment of TF candidates to their candidate target genes, we also searched for their previously characterized TF orthologs in the literature (using PaperBLAST [60]) and also in the RegPrecise database, a reference collection of manually curated inferences of TF regulons in bacterial genomes (44) (see Data Set S1). For identification of candidate TFBSs, we collected initial training sets of potentially coregulated CU genes that are located in conserved genomic neighborhoods with a TF gene in the analyzed 14 Caldicellulosiruptor genomes. For each training set of genes, we extracted intergenic regions of >50 bp and submitted them to de novo DNA motif identification tool, SignalX (62), that uses the expectation-maximization method to find prioritized sets of potential regulatory sites with one of two type of intrinsic symmetry, inverted repeats (palindromes), or tandem repeats. For palindromes, the DNA site size was between 20 and 24 bp, while for inverted repeats we searched for two repeated units, 7 to 11 bp each, that are mutually located with 21- or 32-bp periodicity, corresponding to two or three DNA turns, respectively. For each candidate TFBS motif, we constructed a positional weight matrix (PWM), as previously described (14). For analysis of the Rex regulon, we collected the upstream regions of known Rex-regulated genes in C. bescii and their orthologs from other Caldicellulosiruptor genomes and applied the motif recognition tool with default parameters to identify a 20-bp palindromic motif. For CggR regulon, its DNA-binding motif was identified as a long-inverted repeat containing two 7-bp half-sites separated by a 23-bp spacer, which is according to the previously reported DNA-binding motifs of CggR from the Bacillus and Lactobacillus species (37, 38).

The obtained PWM motifs were further used to scan each Caldicellulosiruptor genome encoding a TF ortholog via the Genome Explorer software enabling the comparative analysis of identified candidate TF regulon members (62). During the PWM scan, we analyzed the whole-genome upstream regions (up to −300 bp relative to the translation start site) and used thresholds for site scores defined as the lowest score observed in the training set. All identified candidate TFBSs were further validated by phylogenetic footprinting using multiple sequence alignment of upstream gene regions for each group of orthologs, as previously described (16, 17). Multiple alignments of DNA upstream regions (see Fig. S1) were obtained by ClustalW2 integrated in the SEED platform (49). WebLogo (version 2.8.2) was used to draw consensus sequences for derived TFBS motifs (63).

Construction of msmK knockout strain.

Generation of C. bescii MACB1080 (ΔpyrE ΔmsmK) was accomplished with allelic exchange in the MACB1018 background (ΔpyrE), as described previously (64). Plasmid pGR027, Athe_1803 (msmK) deletion construct (see Fig. S2), was generated from PCR products with NEBuilder HiFi DNA Assembly (New England Biolabs, Ipswich, MA). PCR products for construct assembly were generated with PrimeSTAR HS DNA polymerase (TaKaRa Bio, Kusatsu, Shiga, Japan). PCR products for screening were generated with SpeedSTAR HS DNA polymerase (TaKaRa Bio). Plasmid was transformed into NEB 10-beta Competent E. coli by heat shock at 42°C. PCR products, plasmid DNA, and genomic DNA were purified with kits from Stratagene and Zymo Research. Prior to transformation into C. bescii, pGR027 was methylated with M.CbeI, as described previously (64); methylation was confirmed by lack of digestion in the presence of HaeIII restriction endonuclease. C. bescii competent cells were prepared as described previously and transformed by electroporation with a Bio-Rad Gene Pulser (65). Electroporation was performed using 1 μg of methylated plasmid DNA with voltage of 1.8 kV, resistance of 400 Ω, and capacitance of 25 μF. After electroporation, the cells were promptly transferred to recovery media, preheated to 70°C. At 0, 45, 90, and 180 min postelectroporation, 5-ml samples were transferred to 50 ml of selective medium containing 50 μg/ml kanamycin and incubated at 75°C with shaking at 150 rpm until growth was observed (24 to 72 h). Cultures with growth were screened for plasmid integration by PCR and colony purified by three rounds of plating and selection to eliminate merodiploidy before counterselection for second crossover (plasmid loss). Counterselection was accomplished on solid medium using modified DSM 516 medium with 8 mM 5-floroorotic acid (5-FOA) and 20 μM uracil. Clean deletion of Athe_1803 was verified by Sanger sequencing (GeneWiz, South Plainfield, NJ) (see Fig. S2).

Phenotypic characterization of msmK knockout strain.

Unless otherwise specified, C. bescii was cultured under anaerobic conditions (80% [vol/vol] nitrogen and 20% [vol/vol] carbon dioxide headspace) at 75°C in modified DSM 516 medium with plate shaking at 150 rpm, as described previously (65). To overcome uracil auxotorophy in modified strains, all media contained 20 μM uracil. Media either contained soluble substrates at a concentration of 25 mM or insoluble substrate loadings of 5 g/liter. Xylose, crystalline cellulose (Avicel), pectin from apple, and beechwood xylan were obtained from Sigma (St. Louis, MO). Glucose was obtained from Amresco (Solon, OH). Cellobiose was obtained from MP Biomedicals (Santa Ana, CA). Prior to medium preparation, pectin and xylan were washed to remove contaminating soluble sugars that could interfere with growth experiments. Pectin and xylan were suspended in cold 70% ethanol, soaked for 30 min, and pelleted by centrifugation at 10,000 × g for 10 min. The wash procedure was repeated three times before substrates were dried overnight under airflow in a fume hood. For colony purification during strain construction, solid growth medium and cultures were prepared using modified DSM 516 medium with 3% agar, as described previously (65). Solid medium cultures were grown at 70°C for 48 h under argon atmosphere.

Growth studies were performed for up to 30 h in biological triplicate in 100-ml serum vials with 50 ml of culture sealed with butyl rubber stoppers. Before beginning growth experiments, MACB1018 (ΔpyrE) and MACB1080 (ΔpyrE ΔmsmK) were revived from glycerol stocks in modified DSM 516 medium with fructose and passaged once before inoculating experimental cultures containing varied sugar substrates. Experimental cultures were inoculated to a cell density of 3.3 × 10⁶ cells/ml. C. bescii competent cells of MACB1018 (ΔpyrE) were grown to a final optical density at 680 nm of 0.06 to 0.07 at 70°C in 1-liter medium bottles containing 500 ml of low-osmolarity defined medium with amino acids mix (LOD-AA) under argon atmosphere and sealed with butyl rubber stoppers, as described previously (65, 66). Cells were counted on a Petroff-Hausser counting chamber (Hausser Scientific, Horsham, PA). Cultures were counted undiluted or diluted either 5-fold or 10-fold in 1× C. bescii base salts, depending on culture density. For protein estimations, the Bradford method was utilized in 96-well plate format with bovine serum albumin standards (Bio-Rad, Hercules, CA).

Transcriptomics. (i) Cell growth and harvest.

Complex media for the cultivation of C. bescii DSM 6725, containing 0.5 g liter⁻¹ yeast extract, was prepared as described previously (43). Five different growth substrates (glucose, cellobiose, crystalline cellulose [Avicel], xylose, and birchwood xylan; 5 g liter⁻¹ [final concentration]) were picked to represent five different conditions for differential transcriptomic analyses. Growth experiments were performed at 78°C as closed anoxic cultures (400-ml volume, shaken at 150 rpm) as described previously (7). Cells were harvested in the mid- to late-exponential-growth phase (0.5 × 10⁸ to 1.5 × 10⁸ cells ml⁻¹) by immediately (<1 min) bringing the culture to room temperature through pumping through a glass cooling coil bathed in an ice-water slurry, as previously described (67), a procedure which also led to the removal of most of the insoluble substrate. Cells were separated from the supernatant by centrifugation at 6,000 × g for 5 min and immediately shock-frozen in liquid nitrogen.

(ii) RNA isolation.

Frozen cell pellets collected from 200 ml samples drawn from each fermenter were resuspended in 2 ml of TRIzol (Invitrogen, CA) and lysed by sonication at 40% power (9 W) on a Misonix 3000 sonicator (Farmingdale, NY). Samples were sonicated for a total of 15 s (3 × 5 s pulse, followed by 1 min off). Chloroform was added, and the sample was mixed and centrifuged. The aqueous layer was removed and mixed 1:1 with 80% ethanol. The mixture was then processed on an RNeasy column (Qiagen, Hilden, Germany) according to the manufacturer’s protocol and the on-column DNase digestion. RNA was eluted off the column in 35 μl of RNase-free H₂O. The RNA was quantified using a NanoDrop 1000 instrument (Thermo Scientific, Waltham, MA), and the RNA quality was assessed using an Agilent 2100 bioanalyzer (Agilent Technologies, Santa Clara, CA) and a Nano6000 chip kit. Purified total RNA was depleted of rRNA using a Ribo-Zero rRNA removal kit for Gram-positive bacteria (Epicentre-Illumina, Madison, WI) according to the manufacturer’s protocol. The depleted sample was concentrated on an RNA Clean & Concentrator-5 (Zymo Research, Irvine, CA) according to the manufacturer’s protocol.

(iii) RNA-Seq library preparation.

Depleted RNA was used to prepare RNA seq libraries using the Epicentre ScriptSeq mRNA-Seq Library preparation kit (Epicentre-Illumina) following the manufacturer’s protocol. Agencount AMPure beads (Beckman Coulter, Indianapolis, IN) were used to purify the cDNA, and unique indexes were added during 13 cycles of library amplification. The final RNA-Seq libraries were purified with Agencount AMPure beads (Beckman Coulter) and quantified with a Qubit fluorometer (Life Technologies, Carlsbad, CA). The library quality was assessed on an Agilent 2100 Bioanalyzer (Agilent) using a DNA 7500 DNA Chip (Agilent). Samples were pooled and diluted, and two paired end sequencing runs (2 × 76 bp) were completed on an Illumina MiSeq instrument (Illumina) using the standard protocol and V3 chemistry.

(iv) RNA-Seq data analysis.

The processing of raw reads was performed based on our previous workflow (68). The raw reads were first assessed for sequencing quality using SolexaQA++ tool kits (69). We performed quality trimming of the raw reads using BWA dynamic trimming algorithm in the SolexaQA++ tool kits to obtain the high-quality reads (phred score > 25 and length after trimming > 25 bp). The high-quality reads were aligned to the C. bescii genome using a high-performance aligner Stampy software (70, 71) with default parameters to generate Bam files. The raw count table was generated from the bam files using BEDTools (72). The raw count table was then used for statistical analysis to identify differential gene expression using the Voom method (73, 74).

(v) 5′ RNA-Seq for TSS determination.

Total RNA from C. bescii cultures grown on glucose, cellobiose, Avicel, xylose, and xylan were also used as starting material for TSS enrichment. Briefly, 4 μg of total RNA was treated with Ribo-Zero magnetic kit for bacteria (Epicentre) to remove rRNA, and the rRNA-depleted material was purified using Zymo RNA Clean & Concentrator (Irvine, CA) and eluted with 10 μl of RNase-free water. Depleted RNA was quantified on a NanoDrop 1000 (Thermo Scientific) and visualized on an Agilent 2100 bioanalyzer instrument (Agilent Technologies). The 5′ terminator exonuclease (Epicentre) was used to digest RNAs that contained a 5′-monophosphate end in a reaction at 30°C for 30 min that also contained an RNase inhibitor (New England Biolabs, Ipswich, MA). Samples were subsequently purified using RNAClean SPRI beads (Agencourt, Beverly, MA) at 1.8× and eluted with 18 μl of RNase-free dH₂O. The exonuclease-treated RNA was then used in a reaction with RNA 5′-pyrophosphohydrolase (RppH) to remove pyrophosphate (New England Biolabs, Ipswich, MA). The reaction proceeded with RNase inhibitor at 37°C for 60 min, and the samples were purified with Agencourt RNAClean SPRI beads (1.8×) and eluted with 11 μl of RNase-free water. The samples were visualized on an Agilent 2100 bioanalyzer instrument. Samples were then ligated with an Illumina 5′ adaptor (5′-ACACUCUUUCCCUACACGACGCUCUUCCGAUCU-3′) using T4 RNA ligase at 37°C for 60 min, followed by Agencourt RNAClean SPRI bead (1.8×) cleanup. An Illumina 3′ adaptor (5′-GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCTNNNNN*N-3′) was then introduced during first-strand synthesis with SuperScript III (Invitrogen) (25°C for 5 min, 50°C for 50 min, 70°C for 15 min, and holding at 4°C), followed by AMpure SPRI bead (Agencourt) purification (1.8×) and elution in 24 μl of nuclease-free water. Samples were indexed and enriched by PCR (95°C for 3 min; 15 cycles of 98°C for 20 s, 55°C for 30 s, and 72°C for 30 s; 72°C for 7 min; and then holding at 4°C) with KAPA HiFi. To prepare samples for Illumina sequencing, DNA was separated on a 2% 1× TAE ultrapure agarose gel, and gel slices were cut from 300 to 700 bp and purified with a Qiagen Minelute column. Final libraries were quantified with a Qubit fluorometer (Life Technologies), and the library quality was assessed on a Bioanalyzer DNA 7500 DNA chip (Agilent). Samples were then pooled and diluted. Pooled barcoded libraries were sequenced in one direction for 151 bases on an Illumina MiSeq instrument.

In total, five 5′ RNA-Seq data were generated for C. bescii grown on different carbon sources. The 5′TSS-generated raw data sets were quality filtered (Illumina CASAVA 1.8) and further analyzed using Rockhopper version 2.03 (75). High-quality reads were aligned to the C. bescii genome using Rockhopper with default parameters, except for the minimum expression level used to determine 5′ UTRs was set to 0.1. The 5′ UTRs were analyzed in the integrated IGV genome browser for each candidate CU operon in C. bescii to identify TSS position (relative to the translational start of the first gene in operon) and 5′ UTR sequence (see Data Set S2). The identified 5′TSSs were visualized on multiple alignments of orthologous upstream regions of CU genes (see Fig. S1) to locate their candidate −35 and −10 promoter elements that correspond to consensus sequences TTGACA and TATAAT, respectively.

Data availability.

All RNA-Seq and 5′ RNA-Seq raw sequencing and processed data are available in the Gene Expression Omnibus (GSE163475).