New mechanistic insight into the digestion of complex dietary fibre by rumen microbiota using combinatorial high-resolution glycomic and transcriptomic analyses

doi:10.21203/rs.3.rs-104578/v1

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New mechanistic insight into the digestion of complex dietary fibre by rumen microbiota using combinatorial high-resolution glycomic and transcriptomic analyses

https://doi.org/10.21203/rs.3.rs-104578/v1

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published 01 Dec, 2021

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Background

The rumen microbial community is considered the most efficient anaerobic digestive ecosystem known, yet less than half of the energy in low quality forages is actually metabolized. There is a knowledge gap regarding the specific factors that impede the ruminal digestion of plant cell walls or if rumen microbiota have the functional potential and activities to overcome these constraints. To address these issues, innovative experimental methods may provide a high-resolution understanding of the cell wall chemistries and higher-order structures that are resistant to microbial digestion and how they interact with the functional activities of the rumen microbial community.

Results

With this goal, we characterized the total tract indigestible residue (TTIR) from cattle fed a high-forage diet containing low-quality straw using two comparative glycomic approaches: ELISA-based glycome profiling and glycosidic linkage analysis. Using these techniques, we successfully detected numerous and diverse cell wall glycan epitopes in barley straw and TTIR and determined their relative abundance pre- and post-intestinal digestion. Of these, xyloglucans and heteroxylans were the most recalcitrant to digestion. Linkage analysis identified indigestible linkages consistent with the polysaccharide epitopes identified by ELISA-based glycome analysis. To determine if residual plant polysaccharides within TTIR could be metabolised, rumen microbiota from cannulated cattle fed barley straw were incubated with barley straw and TTIR in in vitro batch cultures. Transcript coding for carbohydrate-active enzymes (CAZymes) were identified and characterized for their contribution to cell wall digestion based on glycomic analyses, comparative gene expression profiles, and associated CAZyme families. High-resolution phylogenetic fingerprinting of these sequences revealed encoded enzymes with activities predicted to cleave the primary linkages within heteroxylan and arabinan.

Conclusion

This experimental platform provides unprecedented precision in the understanding of forage structure and digestibility, which can inform next-generation solutions to improve the growth of ruminants fed low quality forages and enhance the use of crop residues as a feedstock.

General Microbiology

glycoside hydrolase

CAZymes

rumen microbiome

glycome profiling

linkage analysis

transcriptome

differential gene expression

nutrient utilization

feed recalcitrance

dietary polysaccharides

Lignocellulosic biomass can be converted into a broad suite of fuels, chemicals, and biomaterials. The recalcitrance of cellulosic material and high cost of hydrolysis to simple sugars is a major barrier to the commercial viability of bioproducts developed from cellulosic biomass. With an ever-growing human population and more affluent societies, the global demand for food, meat and milk is projected to increase exponentially [1, 2]. Ruminant livestock are in a unique position to satisfy the growing demand for high quality protein, as ruminants can produce milk and meat via the microbial fermentation of cellulose-rich forages, crop residues, and food by-products. In this light, the rumen microbiota represents an underexploited repository of carbohydrate active enzymes (CAZymes) and microorganisms for applications in animal nutrition, biofuel, and bio-based chemical industries.

Globally, more than 73.9 million metric tons of crop residues are produced annually [3, 4]. Although these residues could serve as feed for ruminants, usually less than half of the biomass they contain is digested. Supplementing ruminant diets with exogenous enzymes has the potential to improve the utilization of crop residues for meat and milk production [5, 6, 7, 8]. However, current commercial enzyme products have been largely developed for biorefinery and bioprocessing applications and were not intended to function in the gastrointestinal tract of ruminants. Physiological conditions within the rumen are not favorable for some enzymes within cellulase mixtures [7, 8], with many of these products possessing enzyme activities that are redundant to those already produced by the rumen microbial community. Quantifying the digestibility of neutral detergent fiber (NDF) and acid detergent fiber (ADF) in ruminants provides only cursory insights into the factors that limit plant cell wall digestion in the rumen. Immunofluorescence microscopy coupled with cell wall specific monoclonal antibodies (mAbs) has proven to be a more informative method to investigate the polysaccharide composition of plant cell walls [9, 10], yet an informed and more tailored approach is needed to investigate ruminal fiber digestion. Such an approach should be anchored in a higher understanding of cell wall recalcitrance to ruminal enzymatic digestion and identification of rate limiting enzyme activities. Once rate-limiting enzymatic activities within the rumen are identified, CAZymes with synergistic activities could be used to enhance the digestion of fiber by rumen microbiota.

Here, we have developed a platform for coupling glycomic analyses to comparative metatranscriptomes of rumen microbiota in batch cultures with barley straw and total tract indigestible residue (TTIR) as substrates. We aimed to characterize recalcitrant cell wall structures and identify candidate enzyme activities with the potential to enhance the rumen digestion of barley straw. Using this approach, we identified indigestible cell wall moieties within barley straw, and characterized the linkages that contributed to recalcitrance. A suite of target genes was selected for downstream production of recombinant enzymes that could potentially overcome the constraints to the digestion of barley straw plant cell walls. Although this study focused on the digestion of barley straw, the experimental pipeline holds high promise for the improvement of lignocellulosic feedstock from diverse plant sources in support of the sustainable conversion of lignocellulose biomass into industrial feedstock.

Glycome profiling of barley straw and total tract indigestible residue

The sequential extraction of cell wall material leads to selective enrichment of various cell wall glycans into fractions based on the degree of integration into the plant cell wall [11]. Oxalate (AO) and carbonate (SC) fractions contain loosely bound pectic and arabinogalactan polysaccharides, while extraction with 1 M and 4 M KOH solubilizes hemicelluloses (e.g., xyloglucans and xylans) and tightly bound pectic polysaccharides. Chlorite (CH) treatment releases lignin and lignin-associated polysaccharides, while a subsequent 4 M KOH extraction after delignification recovers remaining recalcitrant hemicellulose and pectin fractions embedded within crystalline cellulose. Chemical fractionation of barley straw and TTIR followed by enzyme-linked immunosorbent assay (ELISA) identified differences in the glycome profile of these substrates (Fig. 1). One major difference between barley straw and TTIR is the amount of carbohydrate recovered in the oxalate (AO) extracts of these two materials; about two and half times as much carbohydrate was recovered in the AO extract from TTIR as compared with barley straw. Yet, with the exception of the de-esterified homogalacturonan epitopes, which were enriched in the TTIR AO extract in comparison with the AO of barley straw (Fig. 1; box 1), there was very little antibody reactive material in the AO extract of TTIR. This suggests that most of the carbohydrate extracted by AO is too small to bind to the ELISA plate, or too small to include the full epitopes recognized by the antibodies used in this study. In general, the AO and SC fractions of TTIR had minimal or lacked most xyloglucan (XG), xylan, type I rhamnogalacturonan (RG-I), and arabinogalactan (AG) moieties, suggesting that these carbohydrates were digested during passage through the ruminant digestive tract. In addition, there was a notable reduction in the AG-3 and AG-4 epitopes and in the lignin-associated MeGlcA-substituted xylans (Fig. 1, Box 7) in TTIR compared with barley straw. In contrast, the somewhat loosely associated MeGlcA-substituted xylans present in the SC extract appear resistant to degradation during passage through the bovine digestive tract (Fig. 1, Box 2).

In the remainder of the glycome profiles, there appear to be selective enrichments in the TTIR for particular epitopes in comparison with barley straw. For example, there was a relative increase in the abundance of Gal-XG epitopes in the 1M and 4 M KOH extracts of TTIR compared with barley straw (Fig. 1, Boxes 3, 4), as well an increased enrichment of the Xylan-3 and 2-Ara-substituted xylans (Fig. 1, Boxes 5, 6). With diverse 6-linked β-galactan epitopes (common in RG-I and AGPs), there appears to be an enrichment in these epitopes in the 4M KOH extract (Fig. 1, Box 9), but a decreased presence of these epitopes in the 1 M KOH extract (Fig. 1, Box 8) when compared with barley straw. There also appears to be enrichment in some RG-I epitopes in the 4 M KOH extract from TTIR (Fig. 1, Box 10) compared with the corresponding extract from barley straw. Lastly, there is an overall enrichment in the TTIR biomass for the most tightly bound glycans released in the 4 M KOH post-chlorite extraction (PC) compared with barley straw. It is important to note that CH and PC represent the most recalcitrant cell wall fractions. The persistent presence of cross-linked (glucurono) arabinoxylan, xyloglucan, and cellulose-embedded pectins (galactans, RG-I, and AG-2) in the KOH, CH and PC extracts of TTIR indicates their recalcitrance to microbial digestion within the ruminant digestive tract.

Glycosidic linkage analysis of barley straw and total tract indigestible residue

To further define the limits of plant cell wall digestibility by rumen microbiota, we undertook a glycosidic linkage analyses of barley straw and TTIR. The relative abundance of individual monosaccharide linkages in these fraction was determined (Additional File 1) and a ratio of linkage abundance in the barley straw control over the TTIR representing the overall digestibility was calculated (Fig. 2A). A number of linkages within the pectin-rich fraction (EDTA + Na₂CO₃); 2,4-Xylp, t-GalAp, 4-GalAp, 3,6-Galp, 3-Araf, 5-Araf, 2,3,5-Araf, 4-Manp, and 3-Glcp linkages were more readily digested by rumen microbiota. Linkages from the hemicellulose fraction (4 M KOH), t-Araf, t-Xylp, 2-Xylp, 4-Xylp, 2,4-Xylp, 3,4-Xylp, t-GalAp, 4-GalAp, 4-Manp, and 3-Glcp, were freely digested. As expected, the final recalcitrant cellulosic fraction exhibited the lowest level of digestibility for most linkages in all fractions (Fig. 2A).

In order to evaluate the recalcitrance of barley straw to digestion by the rumen microbiota and identify rate-limiting enzymatic activities, the ten most abundant indigestible linkages were calculated as a ratio of abundance in TTIR over that of barley straw, and ranked highest to lowest (Fig. 2B). In the AIR fraction, t-Xylp was found to be highly abundant (1.6 ± 0.6), whereas 3,4-Xylp (5.4 ± 0.4), t-Manp (2.4 ± 0.7), and 2-Araf (2.6 ± 0.9) were the most indigestible linkages identified within the EDTA + Na₂CO₃, 4 M KOH, and cellulosic residue fractions, respectively.

Polysaccharide composition was estimated by assigning the linkage composition data to classes of polysaccharides according to Pettolino et al. [12] (Fig. 2C, Table 1, Additional File 1b). Differences in polysaccharide composition were observed between barley straw and TTIR samples. TTIR contained more arabinan (AB), HX and XG than barley straw in AIR and the final cellulosic residual fractions. The EDTA + Na₂CO₃ fraction of TTIR showed higher HX and XG content. AG-2 content within the TTIR 4 M KOH fraction was observed to be higher as compared to barley straw.

Table 1

Prediction of polysaccharide composition (%) based on glycosidic linkage data.
Polysaccharides¹	AIR		EDTA + Na₂CO₃		4M KOH		Residue
Polysaccharides¹	Barley	TTIR	Barley	TTIR	Barley	TTIR	Barley	TTIR
Arabinan	1.9	2.1	6.3	5.3	2.1	2.5	0.2	0.4
Heteroxylan (HX)	30.8	33.8	17.1	32.4	79.6	76.2	4.7	7.9
Type I arabinogalactan (AG-I)	1.2	1.5	2.9	3.7	0.3	0.4	t.r.	0.1
Type II arabinogalactan (AG-II)	2.2	2.2	12.1	11.6	2.9	4.3	0.3	0.6
Xyloglucan (XG)	5.1	5.5	5.5	8.2	9.0	9.8	2.3	3.0
Mixed linkage glucan (MLG)	3.3	1.9	23.7	21.0	4.2	4.8	1.2	1.4
Cellulose	49.7	47.2	n.a.	n.a.	n.a.	n.a.	84.4	78.3
Homogalacturonan (HG)	0.2	0.3	7.0	2.4	0.4	0.3	t.r.	t.r.
Heteromannan (HM)	0.9	0.8	9.6	5.7	0.5	0.5	0.5	0.8
Unassigned (UA)	4.8	4.7	15.8	9.8	1.1	1.1	6.1	7.4
Note: “t.r.” means trace amount (mol%<0.1%). “n.a.” means linkage not assigned to polysaccharide. No 4-Glcp linkage was assigned to cellulose as EDTA + Na₂CO₃, and 4 M KOH solutions do not extract this fraction [60].
¹ The estimation of polysaccharide was according to Pettolino et al. 2012 [12].

RNA-seq output and de novo assembly

RNA-seq and differential gene expression analyses were conducted to study the composition of rumen microbial communities in vitro, and to compare microbial CAZyme expression profiles of rumen microbes cultured on TTIR or barley straw. Rumen microbial samples were batch-cultured in triplicate in vitro for each substrate (TTIR and barley straw), as reported previously [13], and transcriptomics analyses were performed. A total of 364,583,860 reads were used for quantitative RNA-Seq analysis. The number of reads per sample ranged from 56.95 million to 66.20 million. The transcriptome was assembled de novo using the Trinity assembler [14] and led to 1,998,343 distinct transcripts with a median transcript length of 336 (bp) and N50 of 533 (bp) (Additional File 2).

Microbial taxonomic classification based on putative mRNA.

RNA-seq raw reads were submitted as input to the Kaiju [15] webserver for taxonomical assignment at the whole transcriptome level. The results of the taxonomic classification with Kaiju were further processed and visualized with Krona [16]. Bacteria represented the majority (88%) of rumen transcripts (Table 2). Phylum Firmicutes, Bacteroidetes, Actinobacteria, Fibrobacteres, Proteobacteria, and Spriochaetae were at a relative abundance above 1%. At the family level, Lachnospiraceae (9%) Ruminococcaceae (7%), Spirochaetaceae (5%), Prevotellaceae (4%), Clostridiaceae (4%), Fibrobacteraceae (2%) were most abundant. The transcripts associated with Eukaryotes accounted for 8% of the total transcripts with Ascomycota, Basidiomycota, Neocallimastigomycota, and Intramacronucleata being major phyla (Table 2).

Table 2

Taxonomic analyses of microbiota transcriptome
Domain	Phylum	Family
Bacteria (88%)	Firmicutes (41%)	Lachnospiraceae (9%)
		Ruminococcaceae (7%)
		Clostridiaceae (4%)
		Unclassified Clostridiales (4%)
		Paenibacillaceae (1%)
	Bacteroidetes (17%)	Prevotellaceae (4%)
		Bacteroidaceae (2%)
		Unclassified Bacteroidales (2%)
		Rikenellaceae (1%)
		Porphyromonadaceae (1%)
		Flavobacteriaceae (1%)
	Actinobacteria (9%)	Streptomycetaceae (3%)
	Fibrobacteres (2%)	Fibrobacteraceae (2%)
	Proteobacteria (12%)	Pseudomonadaceae (0.6%)
	Spriochaetae (5%)	Spirochaetaceae (5%)
	Lentisphaerae (0.9%)	Unclassified Lentisphaerae (0.9%)
Eukaryota (8%)	Opisthokonta (64%)	Ascomycota (32%)
		Neocallimastigomycota (6%)
		Basidiomycota (18%)
	Alveolata (9%)	Intramacronucleata (2%)

Note: Taxonomical affiliations of whole transcriptomic data was obtained from taxonomic analyses of mRNA by Kaiju [15]. Relative abundance (%) at the domain level were calculated as a percentage of total mRNA transcripts, whereas relative abundance at the phyla and family level are calculated as a percentage of mRNA transcripts within the affiliated domain. Only those phyla and families with relative abundance higher than 0.6% are reported.

CAZyme expression under in vitro batch culture

To investigate the functional activities of these rumen samples, the RNA-seq raw reads for both barley straw and TTIR cultures were assembled into a total of 1,993,243 transcripts with an average length of 492 bp (Additional File 2) and parsed through BLASTx and the CAZy database (v2017-07-20) [17]. Several transcripts were annotated as CAZymes in the following families: 126 glycoside hydrolase (GH), 27 polysaccharide lyase (PL), and 16 carbohydrate esterase (CE) (Additional File 3). The overall transcriptome contained all the major GH families putatively involved in cellulose and hemicellulose digestion. Substrates for individual GH, PL, and CE families were assigned according to Couger, et al. [18]. A total of 13 GH families possibly involved in cellulose degradation were identified, collectively accounting for ~ 18% of total CAZymes (Fig. 3A, Additional File 4). Major cellulose-targeting GH families included GH5, GH9, GH45, GH48, and GH124 endoglucanases; GH6 and GH48 cellobiohydrolases; and GH1 and GH3 β-glucosidases. Similarly, transcripts targeting hemicellulose degradation included GH10, GH11, GH43, GH5, GH26, and GH8; debranching enzymes belonging to GH2, GH3, GH43, GH51, GH95 and GH43 and acetyl xylan esterase of CE1 family. Out of 39 hemicellulose targeting GH families, only 12 had a relative abundance greater than 1% (accounting for ~ 34% of CAZyme expression, Fig. 3B). Twelve GH, 6 PL, and 3 CE putatively involved in pectin degradation were identified, including polygalacturonases (GH28), rhamnosidases (GH78), galactanases (GH53), pectin lyases (PL1) and pectin acetylesterases (CE12) (Fig. 3C). In addition, transcripts encoding enzymes that degrade starch (pertaining to GH13, GH31, and GH77 families) were also observed (Fig. 3D, Additional File 4).

Transcripts predicted to be involved in cellulose, hemicellulose, pectin and starch digestion (GH + PL + CE) were taxonomically classified to gain insight into the predominant species involved in cell wall degradation. The majority of the cellulose-targeting transcripts were associated with the Firmicutes (42%; Ruminococcus, 9%, Lachnospiraceae, 5%), Bacteroidetes (23%; Prevotella, 7%, Bacteroides, 3%), and Fibrobacteres (4%; Fibrobacter 3%) (Fig. 4A, Additional File 5). Fungi contributed ~ 4% of transcripts that were predicted to target cellulose digestion (Fig. 4A). Similarly, Firmicutes contributed ~ 38% (Clostridium 6%, Ruminococcus 5%) and Bacteroidetes 26% (Prevotella 7%) to the pool of transcripts targeting hemicellulose (Fig. 4, Additional File 5). Likewise, transcripts for families putatively involved in dismantling pectin were extensively assigned to bacteria and primarily to Firmicutes (39%; Ruminococcus, 5%) and Bacteroidetes (24%; Prevotella, 6%). In contrast, starch-degrading transcripts mostly originated from Bacteroidetes (34%; with Prevotella contributing 11% of total starch digesting transcripts). In total, the majority of Firmicutes and Bacteroidetes-assigned CAZyme transcripts targeted hemicellulose and cellulose digestion (Fig. 4B), with Bacteroidetes associated with the transcripts targeting starch digestion. Out of the total Fibrobacter CAZyme transcripts, 59%, 30%, and 11% putatively targeted hemicellulose, cellulose, and pectin, respectively (Fig. 4B).

Gene expression at CAZyme family level

Metatranscriptomic data was used to evaluate the contribution of specific CAZyme families to cell wall degradation. Thus, for those CAZyme family transcripts known to mediate the deconstruction of major cell wall constituents, total expression levels of transcripts for the individual CAZyme family was calculated as a precentage of total expression levels of all CAZyme transcripts (Additional File 6). Endoglucanase and β-glucosidase (GH5, GH8, and GH9) transcripts were higher (p > 0.001) in TTIR cultures (Additional File 6A), while barley straw cultures had a higher abundance of cellobiohydrolase (GH6, GH48) transcripts (Additional File 6B). Interestingly, both GH6 and GH48 families contributed towards the cellobiohydrolase pool for barley straw degradation, whereas GH48 was the only major family of cellobiohydrolases associated with TTIR. Similar transcript abundance was seen for xylan degradation families (GH10, GH11, GH43, and GH39) for TTIR and barley straw (Additional File 6D). Putative pectin-degrading GH families showed higher expression on barley straw with GH78 as the major CAZyme family, whereas GH28 and GH53 were the major pectin-associated GH families expressed with TTIR (Additional File 6F).

Differential gene expression at individual transcript level

Identifying transcripts that catalyze the cleavage of carbohydrate linkages that limit ruminal digestion could provide insight into improving the digestibility of barley straw. Therefore, we focused on differentially expressed transcripts between barley straw and TTIR that are classified as CAZymes. Differential expression analyses identified a total of 88 glycoside hydrolases (GH) that were upregulated when rumen microbiota were cultured with TTIR, while in the presence of barley straw, 130 GH transcripts were (p < 0.01) upregulated (Table 3, Additional File 7). In TTIR samples, 7 PL and 9 CE transcripts were upregulated (p < 0.01), whereas 9 PL transcripts and 21 CE transcripts were upregulated (p < 0.01) with barley straw. At the family level (the total collection of differentially expressed transcripts within a given CAZy family), higher expression was observed in the presence of TTIR of CAZy families known to be putatively involved in degradation of cellulose (GH5), xylan (CE15, GH67), pectin (GH105, PL11) and mannan (GH26), but were not found to be statistically significant due to the high variation in expression levels across replicate batch cultures (Fig. 5, Additional File 7).

Table 3

Total number of upregulated CAZyme transcripts on barley straw and TTIR substrates.
Upregulated Transcript Enzyme Class	Number of transcripts
Upregulated Transcript Enzyme Class	BS	TTIR
Glycoside hydrolase (GH)
(p < 0.01)	130	88
(p < 0.1)	26	3
Polysaccharide lyase (PL)
(p < 0.01)	5	7
(p < 0.1)	0	1
Carbohydrate esterase (CE)
(p < 0.01)	21	9
(p < 0.1)	6	3

To increase the resolution of functional annotation, the nucleotide sequences for differentially expressed (DE) transcripts were re-submitted to the recent version of the dbCAN2 server [19] and manually curated for CAZyme annotation. To distinguish dominant individual DE transcripts, the transcriptional expression pattern for DE transcripts between TTIR and barley straw cultures was studied for CAZy families of interest known to target XG, HX, and pectin (Fig. 6). Transcript TR363754|c1_g3 (GH11), TR744494|c3_g1 (GH16), and TR463944|c0_g2 (CE15) contributed significantly towards TTIR overexpressed DE transcripts as their relative abundance was lower in the presence of barley straw. Whereas other transcripts, such as TR400632|c0_g1 (GH11), TR849235|c0_g1 (GH11), and TR300921|c2_g2 (GH43), were observed to be upregulated in barley straw cultures as compared to TTIR cultures.

Prediction of enzyme activities for rumen DE transcript CAZymes

The differentially expressed transcripts represent uncharacterized rumen microbial CAZyme sequences. As HX and AB were identified as key recalcitrant components by glycome and linkage analyses, statistically significant DE transcripts from relevant polyspecific families GH11, GH43, and CE15 were selected for high-resolution functional prediction with SACCHARIS [20]. Phylogenetic trees generated for biochemically characterized members of GH 11 (Fig. 7A) indicated that TTIR overexpressed transcripts (TR363754|c1_g3, TR668965|c0_g1, TR196282|c0_g1, TR190685|c0_g1, and TR83509|c0_g1) partitioned into distantly-related clades, as compared to those overexpressed in barley straw (TR476599|c0_g1, TR849235|c0_g1, TR327227|c0_g1, TR291133|c0_g1, TR400632|c0_g1, and TR1014827|c1_g1). All GH11 transcripts are predicted to function as endo-β-1,4-xylanases, the dominant enzymatic activity for the family. Similarly, in GH43 (Fig. 7B), TTIR-overexpressed transcript TR712918|c0_g1 was classified as an endo-α-1,5-L-arabinanase, while barley straw-overexpressed transcripts TR400300|c0_g2 and TR451123|c0_g1 were predicted to be putative arabinofuranosidases. Interestingly, TR941931|c2_g1 and TR300921|c2_g2 were both overexpressed in barley straw and were predicted to be endo-α-1,5-L-arabinanases. All three TTIR-overexpressed CE15 transcripts were classified as acetyl xylan esterases (Fig. 7C).

Developing efficient technologies to enhance the conversion of low-quality forages into available energy within the rumen is pivotal for sustainable dairy and beef production [21]. Potentially, such technologies can reduce feed costs and improve the competitiveness of cattle management, while reducing the environmental footprint of meat and milk production [22]. Supplementing ruminant diets with exogenous enzymes has the potential to enhance feed efficiency [7, 8]. However, current commercial enzymes have been tailored for bio-refinery and bioprocessing applications and as such are not optimized for the physiological conditions of the rumen [23, 24, 25]. Likewise, commercial enzymes often contain redundant enzyme activities already encoded within the rumen microbiota. Understanding the rate-limiting steps in ruminal fiber digestion from the perspective of plant cell wall chemistry and endogenous catalytic potential is critical for the successful design of technologies based on enzyme addition, programming of the microbiome, and breeding plant cell walls that are more amenable to digestion by microorganisms within the ruminant digestive tract. Here we present an informed experimental platform (Fig. 8) that couples high-resolution cell wall structural analysis with targeted meta-transcriptomic analysis of rumen microbiota so as to expand the present understanding of the factors that limit plant cell wall degradation.

Straw residues that escape ruminal total tract digestion (TTIR) can provide valuable insight into nutrient utilization by rumen and lower-tract microbes and into plant cell wall components that confer recalcitrance to the hydrolysis of straw. Presented here is the first report using ELISA-based glycome profiling to study the progression of fiber utilization and recalcitrance during total tract digestion of fiber in ruminants. This methodology has been extensively used to characterize plant cell walls including those of genetically modified plants, and microbially and chemically treated plant biomass [9, 10, 26]. Glycome profiling provided us with an unprecedented ability to decipher the architecture of native barley straw and its residual fraction that is recalcitrant to digestion in cattle (TTIR) as it can detect diverse structural features within plant cell wall polysaccharide classes (Fig. 1). For example, a comprehensive suite of more than 25 mAbs that recognize different substructures, including each of the major side-chains of xyloglucans was applied, while another set of approximately 30 mAbs that recognize diverse xylan epitopes including Ara, GlcA and MeGlcA substitutions as well as various lengths of backbone residues [27] were used to assay xylans (Fig. 1). This not only allowed us to identify recalcitrant polymers with high precision, but also to differentiate polysaccharide subclasses that were removed during passage through the ruminant digestive tract or that exhibit high recalcitrance and were enriched in the TTIR. For instance, 4-O-Me-GlcA xylan epitopes that are lignin bound, as well as those xylans loosely associated with the cell walls in barley straw were removed during passage through the digestive tract (Fig. 1). Alternatively, 2-Ara-substituted xylans, as well as xylan epitopes recognized by the Xylan-2 and Xylan-3 groups of antibodies, were enriched in TTIR, suggesting that these epitopes directly contribute to recalcitrance. Relative proportions of xyloglucan and xylan epitopes in TTIR remained largely unchanged from the proportions of these epitopes in native barley straw (Fig. 1). However, it was clear that glycome profiling alone could not fully describe the glycan composition of TTIR. For example, there was a significant fraction of glycans that were loosely associated with the TTIR (AO extract), yet, with the exception of de-esterified HG epitopes (DP > 4), there was no antibody binding to this extracted fraction from TTIR. This lack of antibody binding may be attributed to the fact that short-chain oligosaccharides released during digestion fail to bind to ELISA plates [10]. Alternatively, while the antibody collection used here recognizes a large and diverse set of plant cell wall glycan epitopes, there may be other epitopes native to barley straw or generated during passage through the ruminant digestive tract that are not recognized by the current antibody library. Therefore, to confirm glycome profiling results we quantified individual glycosidic linkages using partially methylated acetyl alditol derivatization, which measures all glycan linkages present. The linkage data showed that the TTIR was enriched in HX, XG, and AB when compared with barley straw at the whole cell wall level (Fig. 2, Table 1), suggesting that these components were more recalcitrant than other components within this biomass. Fractionation of the walls showed that the EDTA + carbonate extract was strongly enriched in HX and XG (Fig. 2, Table 1), which was in contrast to the results from glycome profiling, where the equivalent extract (AO) only showed enhanced HG content (Fig. 1). This may reflect the composition of the large fraction of cell wall glycans that are loosely associated with TTIR, but which were not assayable in the glycome profiling ELISAs for the reasons mentioned above. Consistent abundance of HX and XG within the whole cell wall and the EDTA/carbonate and KOH extracts of TTIR suggest that these epitopes contribute to the recalcitrance of barley straw within the ruminant digestive tract. Apart from cell wall composition, linkage analysis also provided useful insight into the predominant bonds within identified recalcitrant polysaccharides. The results presented in this study could prove to be a valuable resource for crop breeders to develop feed stocks that exhibit increased digestibility in ruminants.

Identification of predominant recalcitrant bonds coupled to transcriptomic and differential expression analyses, provided two complementary approaches to identify candidate linkages to target for improved plant cell wall digestion. To identify potential enzymes tailored for the digestion of recalcitrant linkages, TTIR and barley straw were used as substrates to culture rumen microbial communities in batch cultures. Incubation with TTIR provided an unique experimental approach that enabled the rumen microbial digestion of recalcitrant plant cell walls to be studied. This is the first such attempt where recalcitrant residues like TTIR have been used to study rumen community composition and CAZyme expression. Nutritional value and effective fermentation of TTIR was confirmed by ELISA results and through comparison of total gas production using barley straw and TTIR as substrates in in vitro cultures (Additional File 8). Firmicutes (Lachnospiraceae, Ruminococcaceae, Clostridiaceae), Bacteroidetes (Prevotellaceae, Bacteroidaceae and unclassified Bacteroidiales), Actinobacteria, Fibrobacteres, Proteobacteria and Spirochaetae were found to be dominant bacterial phyla at the transcriptome level (Table 2). These results are in agreement with previously reported metatranscriptomic analysis, such as Dai et al. [28], that reported Firmicutes, Bacteroidetes, Spirochaetes, Proteobacteria, Actinobacteria and Fibrobacteres are the most abundant bacterial phyla. Likewise, Comtet-Marre et al. [29] found that bacteria represented 77.5% of rumen ssu rRNA reads with Firmicutes, Bacteroidetes, Fibrobacteres, Proteobacteria, Spirochaetae and Lentisphaerae as predominant phyla. Although the present study found a similar core microbiome, there was variation in the abundance of taxonomic families as compared to previous literature [28, 29]. This variation may be attributed to the shorter incubation time (48 h), and absence of host effects in closed in vitro batch culture system as compared to in vivo animal-based studies. Shorter incubation times favor rapidly growing bacteria over the slower growing fungal and protozoal members of ruminal microbial communities. In contrast to the host environment where fermentation by-products like VFA and ammonia are absorbed across the intestinal epithelium, closed batch culture systems accumulate by-products, which can alter the fermentation process towards certain types of microbial communities. Furthermore, host genetics have also been reported to influence microbial populations [30], as a region on chromosome 6 in cattle has recently been shown to be associated with Actinobacteria, Euryarchaeota and Fibrobacteres densities [31]. While whole animal experiments can not specifically study the digestion of TTIR, in vitro batch cultures provided us with the level of control over experimental variables required to investigate those factors that limit plant cell wall digestion.

Gene expression analyses of rumen microbial communities cultured on barley straw and TTIR demonstrated substrate-specific modes of digestion, as evidenced by the expression levels of CAZy family transcripts. At the family level, higher expression of putative endo-glucanases GH5, GH9, and GH8 on TTIR indicated an abundance of metabolizable cellulose within TTIR, as confirmed in the linkage analyses. Furthermore, GH48 cellobiohydrolases were the dominant exo-glucanases expressed during the digestion of TTIR by rumen microbes. In contrast, GH6 and GH48 were the predominant families associated with the microbial digestion of barley straw (Additional File 6B). Both GH6 and GH48 are known to efficiently hydrolyze the crystalline regions of cellulose [28, 32]; while cellobiohydrolases of family GH48 act on the reducing ends of cellulose chains, and GH6 attack non-reducing ends, generating cellobiose [33]. The high expression levels of these exo-glucanases suggest there is a greater abundance of amorphous cellulose at the reducing ends of micro-fibrils in TTIR as compared to barley straw, as result of its exposure to CAZymes during passage through the ruminant digestive tract. The level of expression of GH43 hemicellulases in both TTIR and barley straw was higher than other hemicellulose-targeting activities (Additional File 6E). Among the 501 bacteria in the Hungate catalogue, the GH43 family was reported as the most abundant hemicellulase [34]. Metagenomic [35–37] and metatransciptomic [28, 29, 38–40] studies have suggested that members of the GH43 family are the principal hemicellulases within the rumen. GH43 encode a range of debranching enzymes, including arabinofuranosidases, xylanases, galactanases, arabinanases and β-xylosidases, which aid in the degradation of arabinoxylan [41]. Indeed, glycomics revealed that arabinoxylan was enriched in TTIR, highlighting its resistance to ruminal digestion. Additionally, abundance of branched linkages like 3,4-Xylp, t-GlcAp, 2,3,4-Xylp, t-Araf in TTIR also confirm high resistance of heteroxylan to degradation by rumen microbiota (Fig. 2). Metatranscriptomic analyses reflected the efforts of the rumen microbiota to degrade this polysaccharide, with the GH10 family being the dominant family of xylanases (Additional File 6D). This family has been previously reported to digest heteroxylan and release xylotriose with a methylglucuronic acid (MeGA) moiety at the non reducing end which is hydrolyzed by members of the GH5 family of xylanases [42]. The GH5 xylanases have been shown to be active against glucuronoxylan and MeGA substituents on the xylan backbone [42]. Pectin polysaccharides differed between barley straw and TTIR (Figs. 1 and 2). Accordingly, expression levels of pectin-targeting CAZy families differed between rumen microbiota incubated with TTIR vs barley straw (Additional File 6F). Extracted rumen microbial communities cultured on barley straw expressed more GH78s, a family that contains α-l-rhamnosidases that act on rhamnogalacturonan and arabinogalactan-protein linkages [43]. In contrast, with TTIR there was a higher expression of GH28, a family that encodes polygalacturonases and rhamnogalacturonases that hydrolyse the α-1,4 galacturonate linkage in HG and RG-I respectively; and GH53s, which encode for β-1,4-galactanases [44].

Metatranscriptomics of barley straw and TTIR cultures provided a broad picture of the enzymatic potential of the rumen in terms of CAZy families. However, to gain more insight into the enzymatic processes of ruminal digestion, differential expression analyses, a more nuanced approach with in-depth analyses was used. Based on differential expression, several GH families putatively involved in cellulose, hemicellulose, and pectin digestion were identified (Fig. 5). Informed by glycomics, the relative contribution of dominant DE transcripts was determined for candidate CAZyme families with a focus on those activities important for digestion of recalcitrant plant cell walls (Fig. 6). Since candidate transcripts were expressed by the microbial community in the presence of TTIR, it is highly likely that these transcripts coded for enzymes targeted at inaccessible or non-hydrolyzable linkages within recalcitrant polymers of TTIR. As identified transcripts represent uncharacterized microbial genes, phylogenetic analysis using SACCHARIS was employed in an attempt to postulate enzyme function and specificity for DE transcripts from the selected families GH5, GH11, GH43, and CE15 important for XG, HX, and AB saccharification. No significant difference was seen between barley straw and TTIR in the relative contribution of GH5 and GH9 family DE transcripts (Fig. 6), suggesting that rumen microbes may lack specialized xyloglucanases for the utilization of XG, despite its abundance within TTIR. Thus, xyloglucanases may represent a rate-limiting enzyme activity in the rumen, and exogenous supplementation of xyloglucanases may promote straw digestion within this environment. Although no xyloglucanases were identified, substrate-specific gene expression targeted towards HX and AB digestion was evident from phylogenetic trees of GH11, GH43 and CE15 DE transcripts, whereby the expression of specialized xylanases and arabinanases were induced within the rumen microbial community for the digestion of TTIR. Interestingly, GH11 transcripts that were overexpressed in TTIR cultures clustered distantly from the barley straw overexpressed GH11 transcripts, and some of the early diverging sequences, are GH11-like, and may represent members of a new family (Fig. 7A). The divergence of TTIR-overexpressed transcripts also suggests that changes in expression levels is a result of changes of the predominant bacterial species, differing between TTIR and barley straw microbes for utilizing recalcitrant HX. Furthermore, the TTIR-overexpressed GH11 transcript TR83509|c0_g1 was observed to contain an unique arrangement with four tandem GH11-like domains (Fig. 7A), possibly the first report of a novel repetitive xylanase architecture. The phylogenetic analyses of the overexpressed GH43 transcripts revealed that both TTIR (TR712918|c0_g1) and barley straw (TR941931|c2_g1 and TR300921|c2_g2) clustered together with endo-α-1,5-l-arabinanases (Fig. 7B), suggesting differential regulation of arabinanase activity in rumen microbiota cultured on barley straw vs TTIR. The recalcitrance of arabinan to microbial digestion was also reflected by linkage analysis and glycome profiling (Figs. 1 and 2; Table 1). Transcripts TR400300|c0_g2 and TR451123|c0_g1 were overexpressed in incubations with barley straw and clustered with arabinofuranosidases, refective of the cross-linked nature of HX in barley straw (Fig. 7B). Likewise, the three overexpressed CE15 transcripts with TTIR were embedded among acetyl xylan esterase activity, further reflecting the recalcitrant nature of HX. The results shown in this study also identified 5-linked Araf within arabinan, 3- linked Araf within arabinoxylan and Xyl-3AC and Xyl-2AC within acetyl-xylan as linkages that resisted microbial digestion (Fig. 7). These results suggest that rumen microorganisms are quite efficient in utilizing loosely bound pectin and hemicellulose, but are limited in their ability to digest cross-linked core pectin and hemicellulose.

Previous metatranscriptomic studies have been mostly limited to GH family levels with limited or absent information regarding substrate specificity of identified transcripts [28, 29, 38–40]. In this study, transcript-level SACCHARIS analyses in combination with glycome profiling and linkage analysis suggested that substrate-specific CAZyme transcript expression is inducible. The TTIR-overexpressed transcripts and putative activities may identify those CAZymes with the greatest potential to improve the rate and extent of recalcitrant plant cell wall residues by the rumen microbial consortia. Identified target CAZymes of rumen origin are likely to stand a better chance of success when applied as additive enzymes as they are optimized for the complex physiological conditions within the rumen. Future recombinant production and characterization of these candidate CAZymes and their screening is needed to ascertain their synergy to the natural compliment of CAZymes produced by rumen microbiota. In addition, the barley straw overexpressed transcripts and their closely clustered CAZymes signifies important enzyme activities for effective barley straw digestion and hold high intellectual value for their wide application to industrial sectors such as bioenergy, and green chemistry. Here we have shown that integrating high-resolution analytical methods, such as cell wall structural characterization, in vitro batch culture, RNA-sequencing and DE analysis (Fig. 8), can be successfully used to identify recalcitrant polymers and resistant linkages within feeds, as well as inform the biological processes involved in ruminal digestion of polysaccharides.

A comparative study involving changes in cell wall glycan composition, glycosidic linkages, and the relative extractability of polysaccharides within TTIR and barley straw provided an unprecedented level of understanding of nutrient utilization by rumen microbes and helped to delineate the role of matrix polysaccharides in cell wall recalcitrance towards microbial enzymatic digestion. Adopting TTIR as a substrate for batch culture of rumen samples and the application of next generation sequencing to study gene expression by the rumen microbial community, successfully identified a number of CAZyme genes displaying potential for downstream enzyme product development and degradation of recalcitrant residues. Furthermore, incubation of rumen microbial communities on TTIR has proven a powerful approach for enzyme discovery. In this regard, characterization of the small oligosaccharide fraction associated with TTIR may allow targeted searches from other organisms for hydrolytic enzymes capable of digesting these oligosaccharides, whose addition may enhance the rumen digestion of recalcitrant forages. Taken together, the results presented in this study suggest that the cell wall layered structure hinders enzymatic accessibility to recalcitrant cell wall components like HX and XG. Considering the rumen is one of the most efficient biomass digestive systems known, the results of this study hold high value for bioproduct and bioprocess advances with lignocellulosic feedstocks. The integration of high-resolution glycomics with targeted metatranscriptomics analyses is an innovative experimental platform that is expandable to other feed-host systems and industrial fermentation processes as it holds vast promise for addressing rate-limiting reactions in digestive ecosystems.

Experimental setup and collection of samples.

All animal procedures and protocols used in this study were reviewed and approved by the Lethbridge Research and Development Center Animal Care Committee (ACC number 1501) in accordance with the guidelines of Canadian Council on Animal Care (CCAC, 2009). Overview of the experimental setup is graphically represented in Figure 8.

In vitro batch culture was used to culture mix rumen microbes on barley straw and total tract indigestible residues (TTIR) as substrates, as described previously [12]. TTIR was obtained from faecal samples collected from heifers (six) fed barley straw-based diets containing 70:30 forage-to-concentrate on a dry matter (DM) basis [45]. Collected faecal samples were pooled and extensively washed (6-7 times) with 50 mM citrate buffer to remove soluble particles. Given the high digestibility of the concentrate, it was assumed that the majority of TTIR originated from barley straw. The fecal residue obtained was dried for 72 h in an oven at 40 °C and defined as TTIR for use in batch culture experiments. Barley straw was dried at 40°C and ground to pass through a 1 mm screen.

In vitro batch cultures were set up in 3 replicate serum vials. Barley straw and TTIR were weighed (0.7g/bag) into acetone-washed and pre-weighted filter bags (F57 ANKOM bag, Ankom Technology Corp., Macedon, NY) that were then heat sealed. Individual bags were then placed in 125 mL serum vials. Rumen fluid from 4-5 different sites within rumen was collected from four ruminally cannulated Angus x Hereford heifers fed 50% grass hay, 30% barley straw, 15% corn dried distillers grains plus solubles and 5% mineral/vitamin supplement (DM basis) [13]. Collected rumen fluid was strained through cheesecloth and equal volumes from each cow were combined. The inoculum was prepared by mixing rumen fluid 1:4 with mineral buffer [13]. Inoculum (65 mL) was transferred to each vial under a stream of O₂-free CO₂. Vials were sealed with rubber stoppers and placed in a rotary shaker (120 rpm) in an incubator at 39 °C. Gas pressure in each vial was measured at 3, 6, 12, 24 and 48 h of incubation by inserting a 23 gauge (0.6 mm) needle attached to a pressure transducer (model PX4200-015GI, Omega Engineering, Inc., Laval, QC, Canada) and used to estimate gas production according to Romero-Pérez and Beauchemin (2018) [46]: Gas volume = 4.7047 × (gas pressure) + 0.0512 × (gas pressure²).

RNA extraction and sequencing

Total RNA from the solid bound ruminal microbes were extracted as described previously [47]. Briefly, the solid contents within nylon bags from in-vitro batch cultures were recovered upon completion of 48 h of incubation and manually ground to a fine powder using a mortal and pestle for 5 min in liquid nitrogen. Ground samples (~ 200 mg) were placed in 2 mL microfuge tubes and each was mixed with 1.5 mL of TRIzol reagent. The samples were thawed, incubated at room temperature for 5 min, and subsequently the RNA was extracted using the acid guanidinium-phenol-chloroform (AGPC) method [48]. Total RNA was further purified with MEGAclear kit according to manufacturer instructions (Applied Biosystems/Ambion). The RNA concentration and integrity were estimated using an Agilent 2100 bioanalyzer (Agilent Technologies, Mississauga, Ontario, Canada) and RNA 6000 Nano kit (Agilent Technologies) according to the manufacturer recommendations. RNA sequencing was conducted on rRNA depleted library using the HiSeq 4000 PE100 platform at McGill University-Genome Quebec Innovation center.

Data analysis and bioinformatics

RNA-seq raw reads were submitted as input to the Kaiju [15] webserver for taxonomical assignment at the whole transcriptome level. Kaiju was run in greedy mode with a minimum match length of 11 and a minimum match score of 55, with 5 mismatches allowed. The results of the taxonomic classification with Kaiju were visualized with Krona [16].

The raw sequenced reads were further processed using MUGQIC RNA-Seq De Novo assembly and differential analysis pipeline [49]. Raw sequenced reads were trimmed and clipped for sequencing adapters, low quality and short sequences were filtered using Trimmomatic [50], normalized and assembled using Trinity normalization utility inspired by the Diginorm algorithm [51]. The sequenced reads were assembled by the de novo transcriptomic assembly program Trinity. Transcripts from Trinity were aligned against the uniprot_sprot.trinotate_v2.0 protein database using the BLASTx program. The Trinotate suite was used for homology searches to known sequence data (BLAST+/SwissProt/Uniref90), protein domain identification (HMMER/PFAM), protein signal peptides and transmembrane domain prediction (signal/tmHMM), and for comparison to currently curated annotation databases (EMBL Uniprot eggnog/GO Pathways databases). The derived functional annotation data were integrated into SQLite database to prepare the annotation report. To identify putative carbohydrate active enzymes, all sequences were profiled against the CAZy database [16]. Gene abundance was estimated using RSEM and differentially expressed transcripts between substrates were identified using DEseq [52] and edgeR [53] package from Bioconductor. Sequence assemblies were deposited in GenBank under BioProject PRJNA673210.

For DE transcript functional analyses, DE transcript nucleotide sequences were re-submitted to the dbCAN2 meta server [19], manually curated, and categorized by CAZy family. The amino acid sequences from the dbCAN-annotated CAZymes were submitted as input to SACCHARIS for phylogenetic analyses [20]. Sequences and accession numbers of characterized GH11, GH43, and CE15 enzymes were extracted from the CAZy database [17], and ProtTest [54] was used for best-fit model selection using the sequence alignment. FastTree [55] was used to generate the phylogenetic trees which were then annotated using iTOL [56].

Glycome profiling of barley straw and total tract indigestible residue

Alcohol insoluble residues from barley straw and TTIR samples were sequentially extracted using 50 mM ammonium oxalate, 50 mM sodium carbonate [with 0.5% (w/v) NaBH₄], 1 M KOH [with 1% (w/v) NaBH₄], 4 M KOH [with 1% (w/v) NaBH₄], sodium chlorite solution (acidified by glacial acetic acid), and 4M KOH [with 1% (w/v) NaBH₄] resulting in five wall extracts designated AO, SC, 1M KOH, 4M KOH, CH, and 4M KOHPC, respectivelyy [57]. These plant cell wall extracts were probed with a collection of 154 cell wall glycan-directed mAbs [58] using an enzyme-linked ELISA, and binding intensities from this experiment presented as a heat map as described previously [57].

Linkage analysis of barley straw and total tract indigestible residue

Alcohol insoluble residues (AIRs) were prepared according to Wood et al. [59]. The dried AIRs were de-starched by incubation with porcine pancreatic α-amylase in 10 mM Tris-maleate buffer (pH 6.9) following Pettolino et al. [11]. The de-starched AIRs were further fractionated based on the literature [60-63] with modifications. Briefly, the cell walls (~55 mg) were treated with 10 mL of 0.25 M sodium borodeuteride (NaBD₄, 99 atom % D, Alfa Aesar) deionized water solution at 4°C for 24 h, followed by quenching the excess reductant by dropwise addition of 10% (v/v) acetic acid and adjusting pH to neutral. The suspensions were then centrifuged (3000 × g, 0.5 h), and resulting supernatants were pooled with three deionized? water washes of the pellets (with centrifugation between each wash), and a pooled solution (Solution 1) was stored at 4 °C. The pellets were then subjected to sequential extraction of polysaccharides using 15 mL of each of the following solutions: 50 mM EDTA (pH 6.5), 50 mM Na₂CO₃ (containing 25 mM NaBD₄) , and 4 M KOH (containing 25 mM NaBD₄) over 24 h at room temperature with gentle magnetic stirring. After extraction, soluble fractions were separated from residues by centrifugation (3000 × g, 0.5 h), followed by neutralizing the supernatants, washing the residues three times with deionized water (with centrifugation between washes), and pooling all washes into corresponding fractions. The EDTA and Na₂CO₃ extracts and the aforementioned Solution 1 were pooled (designated F_EDTA+Na2CO3). The 4 M KOH fraction and the final residue left after strong alkaline extraction were designated F_4MKOHand F_Residue, respectively. All fractions were dialyzed against deionized water and lyophilized. For linkage analysis of the whole cell walls and the isolated fractions, uronic acids in the samples (~2 mg) were first converted to their 6,6-dideuterio neutral sugars using carbodiimide activation at pH 4.75 followed by NaBD₄ reduction at pH 7.0. Samples were then dialysized against deionized water, and freeze-dried. The carboxyl-reduced samples were converted to their partially methylated alditol acetate (PMAA) derivatives by permethylation with iodomethane and sodium hydroxide in dimethyl sulfoxide [64,65] with 2 M trifluoroacetic acid at 121°C for 1.5 h, reduction with NaBD₄, and peracetylation with acetic anhydride [12]. Deutero-methylation using iodomethane-d3 (≥99.5 atom % D, Sigma) was applied to the whole cell wall and F_4MKOH fractions in order to identify and quantify the linkages from endogenously O-methylated sugars (e.g., 4-O-methyl glucuronic acids) [66-68]. The PMAAs were then subjected to GC-MS and GC-FID analyses on an Agilent 7890A-5977B GC-MS/FID system (Agilent Technologies, Santa Clara, CA). The PMAAs were separated using a medium polarity Supelco SP-2380 capillary column (30 m × 0.25 mm × 0.20 μm, Sigma-Aldrich) with a constant column outlet helium flow rate of 0.8 mL/min. Sample solutions were injected at an inlet temperature of 250°C with a split ratio of 10:1. The oven temperature was programmed to start at 120 °C (hold 4 min) followed by increasing at 8 °C/min to 175 °C, 0.5 °C/min to 183 °C (hold 8 min), 0.5 °C/min to 195 °C, 4 °C/min to 210 °C, and 20 °C/min to 255 °C (hold 8 min). The transfer line temperature was kept at 280 °C. The mass spectrometer was operated in electron ionization (EI), full-scan mode (ionization energy, 70 eV; source temperature, 230 °C; quad temperature, 150 °C). The FID detector was operated at 300 °C (H₂ flow, 30 mL/min; air flow, 400 mL/min; N₂ makeup flow, 25 mL/min). The PMAAs were then identified by comparing their MS fragmentation patterns with those of reference derivatives and the literature [69], and quantified based on the previously reported FID response factors calculated using the effective carbon number concept [70]. Glyosidic linkages were assigned to relevant cell wall polysaccharides, and the relative compositions of each type of polysaccharides were then estimated by summing up corresponding linkage compositions [12,71]. Six separate experiments were conducted for the whole cell walls and the F_4MKOHfractions, of which half were used for deuterio-methylation analyses and the other half for methylation analyses. The F_Residuefractions were subjected to six separate experiments (methylation analysis only). Three separate methylation analyses were conducted for the F_EDTA+Na2CO3fractions.

AIR: alcohol insoluble residue, ADF: acid detergent fiber, AG: arabinogalactan, AO: ammonium oxalate, CAZyme: carbohydrate active enzymes, CE: carbohydrate esterases, CH: chlorite, ELISA: enzyme-linked immunosorbent assay, GH: glycosyl hydrolase, HG: homogalacturonan, HX: hetero xylan, mAbs: monoclonal antibodies, NDF: neutral detergent fiber, PC: post-chlorite, PL: pectin lyases, RG: rhamnogalacturonan, SC: sodium carbonate, TTIR: total tract indigestible residue, XG: xyloglucan.

Ethics approval and consent to participate

Consent for publication

No consent was required for this publication

Availability of data and material

Sequenced transcripts have been deposited into the NCBI SRA database BioProject accession number: PRJNA673210.

Competing interests

Authors declare no competing interest.

Funding

This work was funded by Alberta Agriculture and Forestry, Canada: project ID 2018F182R, and Agriculture and Agri-Food Canada: project ID: J-001589. Glycome profiling of biomass samples was supported by the Center for Bioenergy Innovation (Oak Ridge National Laboratory), US Department of Energy (DOE) Bioenergy Research Center supported by the Office of Biological and Environmental Research in the DOE Office of Science.

Authors' contributions

XX, AKB, DRJ, SV, and MGH contributed for experiment and data analysis for cell wall characterization. AKB, ROP performed in vitro batch culture, RNA extraction, transcriptome sequence analysis. KEL and MR contributed towards data processing, statistical support and graphical representation of data. AKB, DWA, and TAM were funded and conceived the study. AKB, DWA and TAM wrote the manuscript and all authors reviewed and approved the manuscript.

Acknowledgements

The authors thank the Lethbridge Research and Development Centre (AAFC) staff for their technical and animal care assistance.

Authors' information (optional)

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Additional file 1: (A) The relative abundance of the glycosidic linkages as determined by gas chromatography – mass spectrometry/flame ionization detection (GC-MS/FID) of partially methylated acetylated alditol (PMAA) derivatives. (B) Assignment of linkages to the polysaccharides according to Pettolino et al. (2012) [12].

Additional file 2: Trinity de novo assembly metrics.

Additional file 3: Identified CAZyme transcripts based on gene annotation results. Transcripts from Trinity were aligned against the uniprot_sprot.trinotate_v2.0 protein database using the BLASTx program. The Trinotate suite was used for homology searches to known sequence data (BLAST+/SwissProt/Uniref90), protein domain identification (HMMER/PFAM), protein signal peptides and transmembrane domain prediction (signal/tmHMM), and for comparison to currently curated annotation databases (EMBL Uniprot eggnog/GO Pathways databases). To identify putative carbohydrate active enzymes, all sequences were profiled against the CAZy database [16].

Additional file 4: The relative abundance of CAZy family and relative contribution of each family towards total CAZyme expression. The relative abundance is defined as the proportion of transcripts for a given family in the total of all CAZyme (GH, CE, and PL only) transcripts. Percentage of expression for transcripts for GH families putatively out of the total CAZyme (GH, CE, and PL only) expression was quantified. FPKM values were normalized by library size (as calculated by DEseq).

Additional file 5: Taxonomic affiliation of transcripts according to Kaiju analysis.

Additional file 6: Relative contribution of GH family transcripts for in vitro BS and TTIR-supplemented bacterial cultures.

Additional file 7: Predicting CAZyme activities from individual differentially expressed (DE) transcripts by dbCAN2.

Additional file 8. Cumulative gas production by rumen microbiota when incubated with BS and TTIR under in vitro batch culture.

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New mechanistic insight into the digestion of complex dietary fibre by rumen microbiota using combinatorial high-resolution glycomic and transcriptomic analyses

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Abstract

Background

Results

Conclusion

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Background

Results

Glycome profiling of barley straw and total tract indigestible residue

Glycosidic linkage analysis of barley straw and total tract indigestible residue

RNA-seq output and de novo assembly

CAZyme expression under in vitro batch culture

Gene expression at CAZyme family level

Differential gene expression at individual transcript level

Prediction of enzyme activities for rumen DE transcript CAZymes

Discussion

Conclusion

Methods

Abbreviations

Declarations

References

Additional files

Supplementary Files

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