An Outer Membrane Protein Involved in the Uptake of Glucose Is Essential for Cytophaga hutchinsonii Cellulose Utilization

Abstract

Cytophaga hutchinsonii specializes in cellulose digestion by employing a collection of novel cell-associated proteins. Here, we identified a novel gene locus, CHU_1276, that is essential for C. hutchinsonii cellulose utilization. Disruption of CHU_1276 in C. hutchinsonii resulted in complete deficiency in cellulose degradation, as well as compromised assimilation of cellobiose or glucose at a low concentration. Further analysis showed that CHU_1276 was an outer membrane protein that could be induced by cellulose and low concentrations of glucose. Transcriptional profiling revealed that CHU_1276 exerted a profound effect on the genome-wide response to both glucose and Avicel and that the mutant lacking CHU_1276 displayed expression profiles very different from those of the wild-type strain under different culture conditions. Specifically, comparison of their transcriptional responses to cellulose led to the identification of a gene set potentially regulated by CHU_1276. These results suggest that CHU_1276 plays an essential role in cellulose utilization, probably by coordinating the extracellular hydrolysis of cellulose substrate with the intracellular uptake of the hydrolysis product in C. hutchinsonii.

INTRODUCTION

Cytophaga hutchinsonii is a common cellulolytic soil bacterium that belongs to the phylum Bacteroidetes (1–4). A unique strategy is thought to be used by C. hutchinsonii to digest crystalline cellulose and to utilize filter paper as the sole carbon source. The mechanism, however, is largely unknown. Most other well-studied cellulolytic microorganisms apply one of two strategies, the extracellular-free-cellulase system or the cell surface-anchored multiprotein cellulosome, to achieve the efficient degradation of cellulose (5). C. hutchinsonii has been assumed to use a third but poorly understood strategy involving cell surface cellulases that produce cellulo-oligosaccharides that are directly transported into the periplasm for further digestion (6–8). Analysis of the genomic sequences of C. hutchinsonii revealed that all the annotated endoglucanases lack recognizable cellulose binding modules (CBMs), and no obvious homologs of cellobiohydrolases are present (3). Recently, endoglucanases that are processive and may act as functional equivalents of exocellulases, and cellulose binding proteins (CBP) on the outer membrane (OM), have been identified in C. hutchinsonii (9–11), though their exact roles in cellulose digestion remain to be determined.

Active import of oligosaccharides released by cell surface enzymes across the outer membrane has been proven to be crucial in the efficient utilization of polysaccharide substrates by many members of the phylum Bacteroidetes (12). This strategy has been well illustrated for starch utilization by the intestinal anaerobe Bacteroides thetaiotaomicron and involves a series of starch utilization system (Sus) proteins, specifically the cell surface lipoprotein SusD and the porin SusC, responsible for the binding and transport of oligosaccharides (13–16). For C. hutchinsonii, previous results indicated that, besides cellulose, glucose and cello-oligosaccharides can also be rapidly assimilated, and such reducing sugars do not accumulate in the culture supernatant when C. hutchinsonii is cultivated with cellulose as the sole carbon source (4, 7). Although two susC-like genes, each arranged in tandem with a susD-like gene, could be identified in the C. hutchinsonii genome, functional characterization revealed that the two sus-like pairs are not required for C. hutchinsonii cellulose utilization (17). Proteins involved in cellulo-oligosaccharide or even glucose transport and their potential relationship with cellulose utilization, therefore, still remain elusive.

In this study, a C. hutchinsonii mutant with a transposon-inactivated gene locus (CHU_1276) was found to be deficient in utilizing cellulose. The results demonstrate that CHU_1276 is involved in the efficient uptake of glucose and is essential for C. hutchinsonii cellulose utilization.

MATERIALS AND METHODS

Bacterial strains, plasmids, and culture conditions.

The strains and plasmids used in this study are listed in Table 1. Wild-type (WT) and mutant C. hutchinsonii strains were cultivated at 28°C in PY10 medium (1.0% peptone, 0.05% yeast extract, pH 7.3) supplemented with the indicated carbon sources, including glucose, cellobiose, Avicel, or filter paper. Escherichia coli was cultured in Luria-Bertani medium at 37°C supplemented with appropriate antibiotics at the following concentrations when needed: ampicillin, 100 μg/ml; kanamycin, 40 μg/ml; erythromycin, 60 μg/ml; and chloramphenicol, 15 μg/ml.

TABLE 1.

Strains and plasmids used in this study

Strain or plasmid	Description	Reference or source
Strains
C. hutchinsonii
ATCC 33406	Wild type	ATCC
T127	Transposon insertion mutation in CHU_1276; Emr	This study
T345	Transposon insertion mutation in CHU_1276; Emr	This study
T912	Transposon insertion mutation in CHU_1276; Emr	This study
T127 (1276)	Complemented strain with plasmid pCH1276; Emr Cmr	This study
E. coli
DH5α	λ− ϕ80dlacZΔM15 Δ(lacZYA-argF)U169 recA1 endA1 hsdR17(rK− mK−) supE44 thi-1 gyrA relA1	Life Technologies (Grand Island, NY, USA)
DH5α λpir	ϕ80lacZΔM15 ΔlacU169 recA1 endA1 hsdR17 supE44 thi-1 gyrA96 relA1 λpir	Rubin et al. (27)
Plasmids
pHimarEm1	Plasmid carrying HimarEm1; Kmr (Emr)	Braun et al. (19)
pCH03C	Complemented plasmid containing oriC region and cat resistance gene originating from pLYL03; Apr (Emr Cmr)	Zhou et al. (18)
pCH1276	2.5-kb CHU_1276 fragment cloned into pCH03C; Apr (Emr Cmr)	This study

^aAntibiotic resistance phenotypes: ampicillin, Ap^r; chloramphenicol, Cm^r; erythromycin, Em^r; kanamycin, Km^r. Unless otherwise indicated, the antibiotic resistance phenotypes are those expressed in E. coli. The antibiotic resistance phenotypes in parentheses are expressed in C. hutchinsonii.

Transposon mutagenesis, identification of sites of insertion, and Southern blot analysis.

The transposon-containing plasmid pHimarEm1 was introduced into wild-type C. hutchinsonii by electroporation, as described by Zhou et al. (18). Erythromycin-resistant colonies were transferred to a new agar plate by replica plating with a piece of sterile filter paper. Mutants defective in degradation of the filter paper were chosen for further study. Identification of the sites of HimarEm1 insertion was performed as described previously (18). Briefly, NsiI-digested genomic-DNA fragments were self-ligated and transformed into E. coli DH5α λpir. Clones carrying the HimarEm1 and adjacent chromosomal DNA sequences were selected on LB agar containing kanamycin, and the retrieved plasmids were sequenced with primer 615 (19). All the oligonucleotide primers used are listed in Table 2. For Southern blot analysis, chromosomal DNAs of the wild-type and mutant C. hutchinsonii strains were isolated, digested with EcoRV, and resolved by gel electrophoresis. DNA transferred to the nylon membrane was analyzed according to the manual for the DIG High Prime DNA labeling and detection starter kit I (Roche, Basel, Switzerland) (20). A 780-bp internal fragment of the erythromycin resistance gene amplified by primers ermFF and ermFR was labeled as the probe.

TABLE 2.

Primers used in this study

Primer	Sequencea (5′ to 3′)	Description
615	TCGGGTATCGCTCTTGAAGGG	Primer to sequence chromosomal DNA adjacent to IR2 of HimarEm1 (19)
ermFF	ATGACAAAAAAGAAATTGCCCGTTCG	Forward primer to amplify ermF
ermFR	AGGGACAACTTCCAGCATTTCC	Reverse primer to amplify ermF
RT1276F	TGGGTATCATTCGTGAGGACAG	Forward primer for confirming transcription of CHU_1276
RT1276R	ATTACGTTGGTAAGGACCTGCA	Reverse primer for confirming transcription of CHU_1276
RT1277F	TCTATGGTGAAGCTGGTAAGTC	Forward primer for confirming transcription of chu_1277
RT1277R	AATACCACCGATAAGGTCAAGG	Reverse primer for confirming transcription of CHU_1277
RT1278F	TTCATTTGGTACATCGGCT	Forward primer for confirming transcription of CHU_1278
RT1278R	AGCCAATATGGTTAACAGGAATT	Reverse primer for confirming transcription of CHU_1278
RT1279F	AGCCAGATTGCGGTGGTAGAAG	Forward primer for confirming transcription of CHU_1279
RT1279R	GTAACCCGGACCCACTCCTGTA	Reverse primer for confirming transcription of CHU_1279
P1276F	CGGGATCCATATTTTGTTACTTTTAGTATA	Forward primer to clone CHU_1276 promoter
1276R	GCTCTAGATTAGAAAAATATTTTTAATTCAACTG	Reverse primer to clone CHU_1276

^aRestriction enzyme sites in the primers are underlined.

RT-PCR and real-time quantitative RT-PCR.

Wild-type and mutant cells were grown to late-exponential phase, and 2 ml of the cultures was harvested by centrifugation at 6,000 × g at 4°C for 5 min. The pellet was washed once with double-distilled H₂O (ddH₂O) without RNase and stored at −80°C for subsequent RNA extraction. Total RNA was extracted by use of the TRIzol reagent (Invitrogen, Carlsbad, USA) according to the manufacturer's protocol and further digested using a Turbo DNA-free kit (Ambion, Carlsbad, CA, USA) to remove any genomic-DNA contamination. cDNA was synthesized using a Prime-Script RT reagent kit (TaKaRa, Shiga, Japan) according to the manufacturer's instructions. The generated cDNA was used as the template to determine the transcriptional status of genes downstream of CHU_1276. For real-time quantitative reverse transcriptase (qRT)-PCR, wild-type and mutant cells were pregrown to early exponential phase (optical density at 600 nm [OD₆₀₀] ≈ 0.3) in PY10 medium. The cell pellets of 150-ml cultures were transferred into equal volumes of PY10 medium supplied with the indicated carbon sources and incubated for different periods. The cells were then collected, washed once, and stored at −80°C for subsequent RNA extraction. Quantitative PCRs were performed on a Bio-Rad myIQ2 thermocycler (Bio-Rad, Hercules, CA, USA) using SYBR green supermix (TaKaRa, Shiga, Japan). Data analyses were performed using the relative quantitation/comparative threshold cycle (ΔΔC_T) method and were normalized to an endogenous control (16S rRNA gene), with expression on glucose as the reference. Three biological repeats were set for all assays.

Complementation of the CHU_1276 mutant.

The full-length CHU_1276 with its own promoter sequence was amplified from the C. hutchinsonii genome with primers P1276F and 1276R, digested with BamHI and XbaI, and inserted into pCH03C to obtain pCH1276. Plasmid pCH1276 was electroporated into the transposon-mutagenized mutant strain. Transformants were picked from PY10 agar containing both erythromycin and chloramphenicol and cultured in liquid PY10 medium. Plasmid DNA was extracted from the above-mentioned cultures with a plasmid minikit (Omega Biotech, Doraville, GA, USA) and transformed into E. coli. Plasmids isolated from E. coli transformants were then used to verify the integrity of pCH1276 in the complemented strain.

Growth analysis.

When glucose or cellobiose was used as the carbon source in PY10 medium, growth of C. hutchinsonii was monitored by measuring the optical density of the culture at a wavelength of 600 nm. When crystalline cellulose (Avicel PH101; Sigma, St. Louis, MO, USA) was used as the sole carbon source, cellular proteins were determined to reflect the growth status, as previously described (7). For growth assay on solid medium, cells grown to the exponential phase were collected, washed with PY10 containing no carbon sources, and resuspended in a series of dilutions. Equivalent amounts of WT and mutant cells were then spotted on PY10 plates with filter paper on top of the agar (0.6%).

Sugar consumption assay.

The wild-type and mutant strains were cultured at 28°C for 36 h in PY10 with 0.4% or 0.1% (wt/vol) glucose as the sole carbon source. The cells were then harvested by centrifugation at 6,000 × g for 10 min, washed with Na₂HPO₄-KH₂PO₄ buffer (50 mM, pH 6.8), and resuspended in the same buffer (6). The suspensions were mixed with equal volumes of buffer with glucose or cellobiose at a final concentration of 0.1% or 0.4% (wt/vol). The mixture was incubated at 28°C with shaking, and samples were taken and centrifuged at 10,000 × g for 10 min at different time points. Quantitative analysis of glucose and cellobiose in the supernatant by high-performance liquid chromatography (HPLC) was performed as follows. Samples were boiled and then incubated with bed resin TMD-8 (Sigma, St. Louis, MO, USA) overnight at 4°C and filtered with a 0.22-μm membrane to desalt. The salt-free supernatants were transferred to new tubes and stored at −20°C. The amounts of glucose and cellobiose in the supernatant at different times were determined with an LC-10AD HPLC (Shimadzu, Kyoto, Japan) equipped with a RID-10A refractive index detector and a Bio-Rad Aminex HPX-42A carbohydrate column (Bio-Rad, Hercules, CA, USA), as previously described (7). The column was maintained at 78°C and eluted with double-distilled water at a flow rate of 0.4 ml/min.

Enzymatic assays with intact cells.

C. hutchinsonii cells were grown in PY10 with glucose to mid-stationary phase and were collected at 6,000 × g and 4°C for 10 min. The pelleted cells were resuspended in PIPES as the intact cell samples, the cell densities of which were measured as described previously (7). The endoglucanase activity of the intact cells was determined essentially as previously described using sodium carboxymethyl cellulose (CMC-Na) (Sigma, St. Louis, MO, USA) as the substrate (7). β-Glucosidase activity was determined by measuring the amount of p-nitrophenol released, using p-nitrophenyl-β-d-glucopyranoside (PNPG) (Sigma, St. Louis, MO, USA) as the substrate at 20°C (21). One unit of carboxymethylcellulase (CMCase) activity was defined as the amount of enzyme releasing 1 μmol of reducing sugars per minute, and 1 unit of β-glucosidase activity corresponded to transformation of 1 μmol of PNPG per minute under the test conditions. All enzyme assays were performed in triplicate.

Isolation and identification of OMPs adsorbed to cellulose.

Preparation of the OMs and solubilization of OM proteins (OMPs) essentially followed the method of Zhou et al. (18). The solubilized OMPs were isolated and bound to cellulose as described by Zhou et al. (18). The adsorbed OMPs were eluted with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer and boiled for 10 min. The eluates were subjected to SDS-PAGE, followed by silver staining. The protein bands differentially stained between the WT and mutant strains were cut, analyzed by matrix-assisted laser desorption ionization–time of flight (MALDI-TOF) mass spectrometry, and used to search the Mascot PeptideMass Fingerprint data bank (Matrix Science).

Transcriptome and bioinformatic analysis.

Cells of WT and mutant strains were precultured on PY10 with 0.4% (wt/vol) glucose and then transferred to a fresh batch in an equal volume of PY10 medium containing either no carbon source, 0.4% glucose, or 0.1% (wt/vol) Avicel for 4 h. Total RNAs were extracted as described above. The RNA quantity (OD₂₆₀) and quality (OD₂₆₀/OD₂₈₀) were determined using a Nanodrop spectrophotometer (Quawell, San Jose, CA, USA). RNA sequencing (RNA-Seq) (quantification) was performed by staff at the Shanghai Hanyu Bio-Tech (Shanghai, China). Random primers, murine RNase inhibitor, and ProtoScript II reverse transcriptase were used to synthesize first-strand cDNA. The second strand was then synthesized with a second-strand synthesis enzyme mix. Short double-stranded cDNA fragments were purified with AMPure XP beads (Agencourt, Brea, CA, USA), resolved with an end repair reaction, and ligated to sequencing adapters. After purification by agarose gel electrophoresis, suitable fragments were enriched by PCR amplification using a universal PCR primer and an index (X) primer. Qubit (Thermo Fisher Scientific, MA, USA) quantification and a high-sensitivity DNA chip assay were performed to obtain the qualified cDNA library. The library sequencing was performed on a cBot Cluster Generation System using a TruSeq PE Cluster kit (Illumina, San Diego, CA, USA) and finally sequenced with an Illumina HiSeq 2500 (San Diego, CA, USA). Gene expression levels were calculated using the reads per kilobase per million reads (RPKM) method (22), thereby eliminating the effects of different gene lengths and sequencing levels so that the calculated gene expressions could be directly compared among samples. The log₂ ratio indicates the degree of differential expression between two samples and was the ratio of RPKM values for the treatment and control samples. The false-discovery rate (FDR) was used to determine the P value threshold in multiple tests (23). An FDR of ≤0.001 was used to judge the significance of values. Gene accession numbers were annotated according to the C. hutchinsonii genome assembly (http://www.ncbi.nlm.nih.gov/genome/?term=cytophaga+hutchinsonii), and ambiguous cases were annotated manually. The functional information for the transcripts was first obtained from the matching genes in the Swiss-Prot and TrEMBL databases using BLASTp with a threshold E value of <1e−5 and was then compared to the extended Gene Ontology (GO) database. The predicted proteins were also assigned to the Kyoto Encyclopedia of Genes and Genomes (KEGG) database (http://www.kegg.jp/kegg/pathway.html), with the top BLAST hit as annotation. Hierarchical-clustering analysis was performed by the Euclidean distance and complete linkage method with Cluster 3.0 software.

RESULTS

Identification of the gene locus CHU_1276, which is essential for cellulose utilization by C. hutchinsonii.

The hyperactive mariner transposon HimarEM1 was electrotransformed into C. hutchinsonii to screen for mutants deficient in cellulose utilization. Several mutants incapable of growing on the medium with filter paper as the sole carbon source were isolated from over 1,000 erythromycin-resistant colonies. Three mutants, T127, T345, and T912, were found to have the transposon inserted in the gene locus CHU_1276 (Fig. 1A and B). All three mutants exhibited the same phenotype, displaying a growth deficiency on filter paper, and T127 was thus selected for further study (Fig. 1C). The compromised growth of T127 on filter paper displayed by the mutant was largely corrected when the mutant was complemented with a plasmid expressing CHU_1276 (Fig. 1D), indicating that the deficiency in cellulose utilization by T127 was specifically caused by the inserted disruption of CHU_1276.

FIG 1.

Identification of CHU_1276, essential for C. hutchinsonii cellulose utilization by transposon mutagenesis. (A) Schematic illustration of the CHU_1276 locus and the surrounding genes. The insertion sites of HimarEm1 in the T127, T345, and T912 mutants are indicated by the arrowheads. (B) Schematic illustrations of the wild-type CHU_1276 locus interrupted by insertion of HimarEm1 in T127, T345, and T912. Restriction enzyme sites used for Southern blotting are marked. (C) Southern blot analysis of mutant strains probed with a 780-bp erythromycin resistance gene. M, DNA ladder (molecular mass marker). (D) Growth of wild-type and mutant strains on PY10 plates with filter paper as a carbon source. A series of dilutions of WT, T127, and T127 complemented with CHU_1276 cells were incubated for 9 days on filter paper.

The predicted protein encoded by CHU_1276 consists of 747 amino acids (aa) with a molecular mass of 82.2 kDa. Analysis of the predicted secondary structure of CHU_1276 suggests that it consists of 11.24% α-helix, 30.25% β-sheet, and 58.50% loop/coil (https://www.predictprotein.org/), with an N-terminal signal peptide of 23 amino acids. PSORTb analysis predicts that CHU_1276 may reside in the cytoplasmic or outer membrane (http://www.psort.org/psortb/). Structural prediction further predicts that a histone-like HU protein domain exists in its C-terminal region (aa 616 to 680) (http://toolkit.tuebingen.mpg.de/hhpred). The genes CHU_1277, CHU_1278, and CHU_1279 lie immediately downstream of CHU_1276 and were predicted to be transcribed from a single promoter located upstream of CHU_1276. None of the proteins encoded by these genes bears similarity in sequence to proteins of known function. To analyze how the interruption of CHU_1276 affected the transcription of the downstream genes, transcriptions of CHU_1277, CHU_1278, and CHU_1279 were analyzed by RT-PCR (see Fig. S1 in the supplemental material). The results showed that insertion in CHU_1276 did not eliminate the transcription of its immediate downstream genes but resulted in their significantly decreased expression, implying that they may form an operon. Specifically, a single band of 854 bp was detected using a forward primer located 619 bp from the stop codon in the first gene (CHU_1276) and a reverse primer in the second gene (CHU_1277) in the WT, but not in the mutant strain, using cDNA as the template (see Fig. S2B in the supplemental material). The transcription of CHU_1277 to CHU_1279 and cotranscription of CHU_1276 and CHU_1277 were, however, fully restored to the WT level when the mutant strain was complemented by CHU_1276 (see Fig. S1 and S2C in the supplemental material). Together, these results suggest that CHU_1276 and its adjacent genes downstream may form an operon but that the expression of CHU_1277, CHU_1278, and CHU_1279 may be further controlled by a fortuitous internal promoter within the operon that is responsive to the presence of CHU_1276.

Inactivation of CHU_1276 results in compromised growth on cellobiose, as well as on glucose, at low concentrations.

In accordance with its growth deficiency on filter paper, no significant growth was observed in liquid medium with Avicel as the sole carbon source (Fig. 2A). When cultured on cellobiose, the mutant displayed a significantly long lag in growth and did not pick up its full growth until after 72 h, although the final biomass yield was comparable to that of the WT strain (Fig. 2C). This extremely slow startup of growth was also observed for 0.1% (wt/vol) glucose, whereas growth of T127 was largely comparable to that of the WT in PY10 liquid medium with 0.4% (wt/vol) glucose (Fig. 2B). To further investigate whether the defect in the startup of growth was glucose concentration dependent, the growth curves of the WT and the T127 mutant were determined with different concentrations of glucose (Fig. 2D). Whereas the final biomass of both the wild-type and mutant strains decreased with decreasing concentrations of glucose, no significant lag phases were observed for the WT at all concentrations of glucose. In contrast, a glucose concentration-dependent lag was observed for T127, with the longest lag occurring with 0.1% (wt/vol) glucose. Together, these results indicated that CHU_1276 inactivation not only compromised the ability of C. hutchinsonii to assimilate cellulose, but also had an apparent effect on its efficient utilization of glucose and cellobiose.

FIG 2.

Phenotypic characteristics of wild-type and mutant strains of C. hutchinsonii in liquid PY10 with different carbon sources. (A) Growth curves of cells on Avicel. (B) Growth curves of cells on glucose. The solid and open symbols represent growth on higher (0.4% [wt/vol]) and lower (0.1% [wt/vol]) concentrations of glucose, respectively. (C) Growth curves of cells on cellobiose (0.4% [wt/vol]). (D) Growth curves of the WT (left) and the T127 mutant (right) on different concentrations of glucose. The data shown are the means of three independent experiments. The error bars show the standard deviations (SD) for the replicates.

CHU_1276 is involved in the efficient hydrolysis of cellobiose and the uptake of glucose by C. hutchinsonii.

To investigate any effect on the hydrolytic activities associated with C. hutchinsonii that may result from the absence of CHU_1276, the specific CMCase and β-glucosidase activities of the intact cells were determined. As shown in Fig. 3, whereas the CMCase activities of T127 were almost the same as those of the wild-type strain, the β-glucosidase activity was 21% lower than that of the WT. To further probe the possibility that this reduction in β-glucosidase activity in T127 might account for the growth phenotype on cellobiose, cellobiose consumption by the WT and the mutant strains in a resting system was determined (Fig. 4A). Whereas relatively rapid hydrolysis of cellobiose and a corresponding extracellular accumulation of glucose occurred in both wild-type and mutant cells, the consumption rate of the mutant cells was much lower than that of WT cells during the first few hours of incubation. Considering the similar growth phenotype of T127 on low concentrations of glucose, the uptake of glucose by WT and mutant cells was also determined (Fig. 4B and C). The results showed that the absence of CHU_1276 resulted in uptake of glucose at a low concentration (0.1% [wt/vol]) that was significantly slower than that of the WT, though the mutant displayed a rate similar to that of the WT at 0.4% (wt/vol) (Fig. 4B and C). However, a WT rate of uptake was restored after T127 cells were grown on 0.1% (wt/vol) glucose. It took only 3 h for cells cultured under these conditions to consume almost all the substrate compared with about 24 h for cells cultured with 0.4% (wt/vol) glucose (Fig. 4C, inset). To test whether mutant cells precultured on 0.1% (wt/vol) glucose restored the ability to initiate growth normally, we determined the growth curves of T127 cells inoculated from a 0.1% (wt/vol) glucose culture. As shown in Fig. 4D and E, although the final cell density of T127 was somewhat lower than that of the WT, hardly any difference was observed between the WT and T127 in picking up the growth with either 0.1% (wt/vol) glucose or 0.4% cellobiose. Of note, T127 cells precultured under these conditions were still incapable of utilizing cellulose (data not shown). Altogether, these data suggest that, besides its essential role in cellulose utilization, CHU_1276 plays an important role in facilitating the efficient hydrolysis of cellobiose and the uptake of glucose present at lower concentrations.

FIG 3.

Enzyme activity analysis of WT and T127 intact cells. (A) Specific CMCase activity. (B) β-Glucosidase activity. The error bars indicate standard errors for three replicates. The P value for the β-glucosidase activity was 0.017, indicating that the difference was significant.

FIG 4.

Cellobiose and glucose consumption by WT and T127. (A) HPLC analysis of cellobiose consumption (circles) and the corresponding accumulation of glucose (squares) by the resting cells of WT and T127 pregrown on 0.4% (wt/vol) glucose. The initial concentration of cellobiose was 0.2% (wt/vol). (B and C) Glucose uptake by WT and T127 cells by analyzing the residual glucose after the resting cells were incubated with glucose for the indicated times. WT and T127 cells were pregrown on 0.4% (wt/vol) glucose and then incubated with 0.4% (wt/vol) glucose (B) or pregrown on either 0.4% (squares) or 0.1% (circles) glucose and then incubated with 0.1% (wt/vol) glucose (C). The inset shows the initial 180 min of glucose consumption by cells pregrown on 0.1% (wt/vol) glucose. (D and E) Growth of the WT and T127 strains pregrown on 0.1% (wt/vol) glucose in PY10 with 0.1% (wt/vol) glucose (D) and 0.4% (wt/vol) cellobiose (E) as the sole carbon source was analyzed. The values are the means of three independent replicates.

Outer membrane proteins affected by the absence of CHU_1276.

To identify the differentially expressed proteins potentially resulting from the absence of CHU_1276, outer membrane proteins extracted from the WT and T127 cultured with 0.4% or 0.1% (wt/vol) glucose were bound to cellulose and were resolved by SDS-PAGE (Fig. 5). Apparently different profiles were observed for either WT or T127 cells when comparing their cellulose-bound OMPs from two different glucose concentrations (0.4% versus 0.1%). Whereas the profiles of the mutant and WT strains cultured on 0.4% (wt/vol) glucose were very similar, they were quite different on 0.1% (wt/vol) glucose. At least four protein bands were missing or reduced and three other proteins were gained in the mutant OM fractions compared with the WT. The bands that were different between the WT and mutant were cut and identified by mass spectroscopy. Figure S3 in the supplemental material demonstrates the identification of the indicated proteins. Among the identified proteins, CHU_1276 (band a) was detected only in the wild-type cells when cultivated on 0.1% (wt/vol) glucose, confirming that CHU_1276 is an outer membrane protein that is induced by a low concentration of glucose. CHU_1253 (band b) and CHU_0344 (band c) were otherwise detected only in T127 cultured on 0.1% glucose. CHU_1253 is a hypothetical protein that has relatively high similarity (51% identity) in sequence to FmdC, an outer membrane porin of Indibacter alkaliphilus belonging to Bacteroidetes. CHU_0344 is the main extracellular protein of C. hutchinsonii, with most of its orthologs containing PKD (polycystic kidney disease protein PKD1) and IG-like domains that may be involved in substrate adhesion (24).

FIG 5.

SDS-PAGE of the OMPs bound to cellulose. The asterisks indicate the different bands between WT and T127 under different culture conditions. Lanes: M, protein ladder as a molecular mass standard; 1 and 2, OMPs from WT and T127 cells, respectively, cultivated on 0.4% (wt/vol) glucose; 3 and 4, OMPs from WT and T127 cells, respectively, cultivated on 0.1% (wt/vol) glucose.

Transcriptional profiles of the wild-type and mutant strains.

To better understand the differences in the process by which the WT and T127 sense and respond to different carbon sources, the full genomic responses in both strains were assessed using RNA-Seq to profile genome-wide mRNA abundance when exposed to different culture conditions. Four hours after transfer to Avicel, 213 genes (5.6% of the total number of protein-encoding genes) that were significantly induced were identified, and 450 genes were found to be downregulated in WT cultures compared with those shifted to glucose conditions (≥2-fold change) (Fig. 6A). As an alternative reference condition, a total of 431 genes showed the same pattern of regulation observed on Avicel when WT cultures were shifted to no-carbon conditions (≥2-fold change) (Fig. 6A and C; see Data Set S1 in the supplemental material). The remaining 232 genes differentially expressed (59 upregulated and 173 downregulated) between Avicel and glucose conditions thus constituted the actual gene set with specific response to Avicel and were referred to as the “Avicel regulon” (Fig. 6D; see Data Set S2 in the supplemental material). A large proportion of hypothetical proteins (65%) were included in this regulon. As expected, CHU_1276 itself, as well as its downstream genes CHU_1277, CHU_1278, and CHU_1279, was among the upregulated regulon genes, and their induced expression was almost abrogated in the CHU_1276-interrupted mutant (Fig. 7A; see Fig. S4 in the supplemental material). Three of the four annotated β-glucosidase genes and two other genes encoding glycosidases of uncertain specificity were also found to be specifically upregulated by Avicel. Unexpectedly, all the annotated endoglucanase genes except CHU_2103 and CHU_1727 were also upregulated by the shift to no-carbon conditions without further induction on Avicel (Fig. 7B; see Data Set S3 in the supplemental material). Similar to the endoglucanases, the majority of proteins annotated as glycohydrolases that may be involved in hemicellulose digestion and glycosidases of uncertain specificity were primarily induced under no-carbon conditions (see Data Set S4 in the supplemental material), whereas glycohydrolases with predicted functions unrelated to cellulose or hemicellulose digestion respond to neither no carbon nor cellulose. Genes encoding a PemK-like growth inhibitor (CHU_0745), a starvation-inducible DNA-binding protein (CHU_2076), and a universal stress protein (CHU_1135), as well as quite a few two-component response regulator and transcription regulator genes, were downregulated under Avicel conditions (see Data Set S2 in the supplemental material). The involvement of these genes in cellulose utilization warrants further dissection.

FIG 6.

Differential transcriptional responses of WT and T127 strains to different carbon sources as analyzed by RNA-Seq. (A) Summary of genes differentially expressed in the WT after transfer of a glucose culture to Avicel versus no added carbon source. (B) Comparison of the differentially expressed genes between T127 and the WT on glucose and those in the WT after transfer of a glucose culture to no added carbon source. For each panel, genes with a log ratio of greater than or equal to 1 or less than or equal to −1 in each set were compared to identify those that were common or distinct among the sets, and the numbers refer to the number of genes in the set. (C) Hierarchical clustering of the GO transcripts with significantly different expression levels in the WT after transfer from glucose to Avicel or no added carbon source, as well as between the WT and T127 on glucose. The heat map was generated from hierarchical-cluster analysis of the 2,005 genes with significantly different expression levels under the above-mentioned conditions (>2-fold changes), with each row representing a different gene. (D) Numbers and functional-pathway categories of the Avicel regulon genes specifically responsive to cellulose in the WT strain.

FIG 7.

(A) Expression patterns of the gene locus CHU_1276 to CHU_1280 under different culture conditions in wild-type C. hutchinsonii and T127. The expression levels of the downstream genes in T127 when the cultures were shifted from a preculture on PY10 with 0.4% (wt/vol) glucose to 0.1% (wt/vol) Avicel compared to those in the WT under Avicel or no-carbon conditions are summarized. (B) Summary of the changes in the expression levels of proteins predicted as cellulases in C. hutchinsonii in response to different carbon sources. Cells of WT strains were precultured on PY10 with 0.4% (wt/vol) glucose and transferred to PY10 containing either no carbon source or 0.1% (wt/vol) Avicel for 4 h. Gene expression levels were determined by RNA-Seq.

The CHU_1276 mutant strain showed expression profiles apparently different from those of the WT, not only on Avicel, but also on glucose (Fig. 8A). A total of 1,508 genes were differentially expressed (≥2-fold change) compared to the WT on glucose. Of note, over half (54.7%) of these differentially expressed genes resulting from the absence of CHU_1276 on glucose showed a similar pattern of regulation in WT cells when they were shifted to no-carbon conditions, including six endoglucanase genes, as well as a significant number of membrane protein genes, sensor protein genes, and transcription regulator genes (Fig. 6B and C; see Data Set S5 in the supplemental material). This result implies that disruption of CHU_1276 affects the appropriate response to glucose. When the same mutant culture was shifted to Avicel, a large proportion of the Avicel regulon genes identified in the WT did not show the same change in T127 (see Data Set S2 in the supplemental material). Of the total of 1,054 genes that were differentially expressed compared to the WT on Avicel (see Data Set S6 in the supplemental material), a set of 428 genes (87 upregulated and 341 downregulated) were identified whose differential expression between the WT and the mutant strain was found to specifically result from the incubation on Avicel (Fig. 8B and C; see Data Set S7 in the supplemental material). They are assumed to represent the global effect of CHU_1276 on the transcriptional profile of C. hutchinsonii when it is cultivated on Avicel and reflect not only primary, but also secondary and tertiary effects.

FIG 8.

Transcriptional response of the T127 strain to cellulose compared to that of the WT. (A) Numbers of genes dysregulated, subdivided into genes repressed (white) and genes activated (black), due to the absence of CHU_1276 compared to the WT under three different culture conditions. (B) Identification of genes differentially expressed between the WT and the mutant strain that were specifically responsive to Avicel. The profiling data were filtered to obtain three sets of genes with a 2-fold or greater change between the WT and T127 under three different carbon conditions. The sets were then compared to identify those that were common or distinct among the sets. The numbers refer to the number of genes in the set. The full set of corresponding genes are listed in Data Set S5, and the 428 Avicel-specific genes are listed in Data Set S6 in the supplemental material. (C) Numbers and functional categories of pathways in T127-Avicel with expression levels significantly higher or lower than those in the WT specifically in response to Avicel.

DISCUSSION

Cellulose digestion by C. hutchinsonii occurs in a cell contact-dependent mode (4, 7, 25, 26), and C. hutchinsonii has been assumed to utilize cellulose with a strategy similar to that of B. thetaiotaomicron. This demonstrated strategy involves a series of Sus proteins, including SusD and SusC, which are thought to mediate the active transport of oligosaccharides across the outer membrane (12–16). C. hutchinsonii has two susC-like genes and two susD-like genes (3). Recent results, however, have revealed that the susC-like and susD-like genes are not required for efficient cellulose utilization or for growth of C. hutchinsonii on crystalline cellulose (17), indicating that the Bacteroidetes Sus paradigm for polysaccharide utilization may not apply to C. hutchinsonii cellulose utilization. C. hutchinsonii may thus adopt an alternative mode of utilization in which the cell surface enzymes of C. hutchinsonii digest cellulose extracellularly to cellobiose and glucose, eliminating the need for SusC-like and SusD-like proteins to import longer oligosaccharides. A recent study showed that extracted outer membrane proteins containing endoglucanase and β-glucosidase activities had the ability to hydrolyze cellulose to glucose in vitro (6). Interference with the efficient hydrolysis of cellobiose into glucose resulting from the absence of an outer membrane protein, CHU_1277, was further found to compromise the ability of C. hutchinsonii to assimilate cellulose (6). In the present study, we identified CHU_1276, a gene locus upstream of CHU_1277, whose disruption not only compromised the efficient hydrolysis of cellobiose, but also affected the uptake of glucose at a low concentration. The connection between the decrease in glucose uptake and the defect in cellulose utilization is not fully understood at present. However, a simple feedback inhibition could be postulated, where even the subtle accumulation of glucose due to the inefficient intracellular delivery of glucose during cellulose digestion may predispose C. hutchinsonii cells to adjust their cellulolytic capacity. The possibility that lower glucose feeding, together with the compromised cellobiose hydrolysis, may make the cells energetically less favorable for assimilation of cellulose cannot be excluded. CHU_1276 may itself directly mediate the uptake of glucose or exert an indirect effect on the transport by regulating the expression and functions of other porins. Evidence supporting the latter assumption comes from the fact that expression of two specific putative glucose transport permeases (CHU_1656 and CHU_1403) of the major facilitator superfamily were found to be affected by the absence of CHU_1276 on Avicel (see Data Sets S6 and S7 in the supplemental material). The recovered expression of these or other, unknown high-affinity transporters after being pregrown on low glucose, which are otherwise inefficiently induced in the absence of CHU_1276, may explain the restored normal growth of the mutant with 0.1% glucose. Notwithstanding this, one can speculate that CHU_1276 has additional roles in cellulose assimilation besides being involved in modulating the tightly coupled cello-oligosaccharide hydrolysis and import of the hydrolysates, considering that CHU_1276 mutant cells pregrown on 0.1% glucose were still unable to utilize cellulose. Based upon the previous observation that CHU_1277 plays an important role in assisting cell surface β-glucosidase to achieve sufficient activity for cellulose utilization (6), our findings provide further support for the notion that CHU_1276, as well as its downstream genes, including CHU_1277, represents the first identified gene cluster important in controlling C. hutchinsonii cellulose degradation. Moreover, CHU_1276 was found to exert a regulatory effect on the expression of its downstream genes. The exact functions of the individual proteins in the cluster and their potential interactions warrant further study to establish a definitive relationship between the hydrolysis of cello-oligosaccharides, including cellobiose, and the subsequent uptake of glucose and cellulose utilization.

Unlike most other well-studied cellulolytic microorganisms whose cellulolytic capacity as represented by an arsenal of hydrolytic enzymes is highly responsive to the presence of cellulose, the hydrolytic activities associated with C. hutchinsonii do not appear to be greatly induced upon incubation with cellulose (7). In this study we analyzed the whole transcriptomes of the C. hutchinsonii WT and the CHU_1276 mutant to identify the genes that are uniquely responsive to cellulose utilization. Several points could be made from these analyses. First, our transcriptional analysis revealed that the majority of the annotated endoglucanases were already upregulated when the cell culture was shifted to medium with no added carbon source but without further substantial induction on Avicel. Although an “Avicel regulon” specifically responsive to Avicel could be inferred in the WT, it is premature to say that this gene set could fully account for the establishment of the cellulolytic capability of C. hutchinsonii, considering that a significant number of genes (431), including 6 of the 9 annotated endoglucanase genes, were found to display the same pattern of regulation under no-carbon conditions as observed on Avicel. Whereas a two-stage mode of induction (derepressed and inductive phases) has been suggested for the cellulolytic fungus Trichoderma reesei based on the similarity of transcriptional profiles between Avicel and no-carbon conditions, the exact connection between the transcriptional response with no added carbon, a mimicry of a condition when C. hutchinsonii was first shifted to cellulose, and the full development of cellulolytic capacity needs further investigation. Second, when comparing the transcriptomes of the WT versus those of the CHU_1276 mutant, over 50% of the genes differentially expressed between the WT and the mutant strain on glucose were found to overlap those differentially regulated in the WT between no carbon source and glucose conditions. This result suggests that the absence of CHU_1276 exerts a profound effect on the response to and assimilation of glucose by C. hutchinsonii. Third, cellulose digestion by C. hutchinsonii occurs in a cell contact-dependent mode, and outer membrane proteins are thus considered to play important roles in both adhering to substrate cellulose and coordinating cellulose depolymerization and the uptake of hydrolytic products (6, 7, 18, 25, 26). Not surprisingly, a significant number of genes encoding outer membrane proteins and transporter proteins were among these differentially expressed genes affected by the absence of CHU_1276 (see Data Set S6 in the supplemental material). Our observation that quite different profiles of cellulose-bound outer membrane proteins were obtained for the mutant and WT strains further corroborates the above-described effect. Finally, a specific gene set was further identified in the absence of CHU_1276 upon incubation with Avicel. Although transcriptional profiling measures the consequences of the change of a regulator, and the transcriptional profile thus reflects secondary and even tertiary effects on gene expression, the identified gene set may be the regulated targets of CHU_1276 and thus account for the deficiency in cellulose utilization by C. hutchinsonii with inactivated CHU_1276. One could anticipate that systematic dissection of the functions of these differentially regulated genes would lead to more information on the pleiotropic effect exerted by CHU_1276. Overall, much remains to be learned about the physiology of the CHU_1276-mediated response and how C. hutchinsonii is capable of integrating information from multiple sources by utilizing specific outer membrane proteins and downstream systems to influence efficient cellulose utilization.