Next Article in Journal
Development of a Taxon-Specific Real-Time Polymerase Chain Reaction Method to Detect Trichoderma reesei Contaminations in Fermentation Products
Next Article in Special Issue
Ex Vivo Fermentation of Hay and Corn by Rumen Bacteria from Cattle and Sheep
Previous Article in Journal
Profile of PKS and NRPS Gene Clusters in the Genome of Streptomyces cellostaticus NBRC 12849T
Previous Article in Special Issue
Effects of Exogenous Oral Infusion of Volatile Fatty Acids on Ileal Microbiome Profiling and Epithelial Health in Goats
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Fermentation
Volume 9
Issue 11
10.3390/fermentation9110925
fermentation-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Fructan Catabolism by Rumen Microbiota of Cattle and Sheep

by
Jennifer R. Weinert-Nelson
1,
Isabelle A. Kagan
1,
Donald G. Ely
2,
Michael D. Flythe
1 and
Brittany E. Davis
1,*
1
Forage-Animal Production Research Unit, Agricultural Research Service, United States Department of Agriculture, Lexington, KY 40546, USA
2
Department of Animal and Food Sciences, University of Kentucky, Lexington, KY 40546, USA
*
Author to whom correspondence should be addressed.
Fermentation 2023, 9(11), 925; https://doi.org/10.3390/fermentation9110925
Submission received: 28 September 2023 / Revised: 19 October 2023 / Accepted: 20 October 2023 / Published: 24 October 2023
(This article belongs to the Special Issue In Vitro Fermentation, 3rd Edition)
Browse Figures
Versions Notes

Abstract

:
Fructans serve as the primary form of storage carbohydrate in cool-season grasses, but little is known about potential differences in ruminal fermentation of fructans between cattle and sheep. An ex vivo study was conducted to evaluate species differences in fructan catabolism. Buffered media containing ground orchardgrass (Dactylis glomerata L.) substrate was inoculated with uncultivated rumen microbiota obtained from cattle and sheep (n = 4 species−1). Fructan profiles were monitored over the incubation period (8 h; 39 °C) using high-performance anion-exchange chromatography coupled to pulsed amperometric detection (HPAEC-PAD). In both species, disappearance of long-chain fructans (degree of polymerization [DP] > 8) was evident by 2 h of incubation (p < 0.01), whereas short-chain fructans (DP 4–8) increased from 0 to 2 h prior to subsequent degradation (p < 0.01). However, the overall rate of long-chain fructan catabolism was greater in bovine versus ovine fermentations, particularly between 2 and 4 h (p < 0.01). Additionally, rapid utilization of short-chain fructans occurred from 2 to 4 h in bovine fermentations, but was delayed in ovine fermentations, with substantial degradation occurring only after 4 h of incubation (p < 0.01). These results indicate that rumen microbiota of cattle may have a greater capacity for fructan degradation.

1. Introduction

Fructans are fructose polymers that serve as the primary form of storage carbohydrate in a number of plant species, including cool-season grasses [1]. In the plant, stored fructan can be mobilized as an energy source to support active growth, while also playing a role in regulation of osmotic adjustments in response to abiotic stressors [2,3]. Fructan polymers can consist entirely of fructose residues connected by fructosyl-fructose linkages [4,5], but often include a glucose moiety (French and Waterhouse, 1993). Fructans are divided into structural classes (e.g., inulin, levan) based on position of the glucose unit and the type of linkage between fructose residues [6,7,8]. Fructans also vary in degree of polymerization (DP). The structural class and DP of fructans differ amongst both plant species and cultivars [9,10], as well as between tissues within a given species [11]. Seasonal effects on fructan concentrations and DP have also been reported [9,10,12].
Depending upon the above-noted factors as well as other variables including plant growth stage, geographic location, and environmental conditions, fructans have the potential to comprise a substantial proportion of the total non-structural carbohydrate content and overall plant biomass in cool-season grasses [13,14,15,16]. Therefore, in ruminant animals grazing pasture grasses or fed conserved forages (i.e., hay or silage), fructans may represent an important nutrient source. In the rumen, fermentation of fructans by the resident microflora produces short-chain fatty acids (SCFA) [17,18]. Once absorbed, these fermentation end-products can be utilized to meet energy requirements of the ruminant animal.
Despite the relevance of fructans in ruminant nutrition, research evaluating ruminal degradation of fructans is relatively limited. Overall, there are indications that DP may influence fermentation [19,20,21,22]. While specific bacteria may have differential capacities for degrading higher-DP long-chain fructans or lower-DP short-chain fructans [19,21], mixed bacteria harvested from equine digesta have been shown to degrade long-chain fructans prior to short-chain fructans [20]. More recently, Kagan et al. [22] found bovine rumen bacteria degraded long-chain fructan within the first two hours of fermentation, which was coupled with an increase in short-chain fructans, suggesting hydrolysis of long-chain fructans to short-chain fructans prior to subsequent degradation.
Degradation of fructans by ruminal bacteria of cattle [18,22] and sheep [17,23] has been evaluated separately, but few comparisons have been conducted to evaluate potential differences between these species. Weinert-Nelson et al. [24] did find lactate accumulation to be 40 to 70% greater when cool-season grass hay was incubated with bovine versus ovine rumen bacteria. Although neither fructanolytic bacteria and fructan fermentation were evaluated in that study, this difference was attributed to potential fermentation of fructans and other non-structural carbohydrates in the hay substrate given the minimal starch concentrations present in cool-season grasses [15,16,24]. To the authors’ knowledge, only one previous study has been conducted in which ruminal fructan fermentation has been directly compared between cattle and sheep, finding differences in the rate of fructan utilization [25]. However, this study utilized purified inulin as the plant-based fermentation substrate, and differences have been noted in fermentation of β(1 → 2)-linked inulin versus β(2 → 6)-linked fructans extracted from cool-season grass [18]. Additionally, no prior studies have characterized impacts of fructan DP on the fermentation of grass fructans between ruminant species. Therefore, the objective of the current study was to utilize a recently developed chromatographic method [22] to evaluate ex vivo fructan catabolism by ruminal bacteria of cattle and sheep during fermentation of orchardgrass (Dactylis glomerata L.) as a model cool-season grass.

2. Materials and Methods

2.1. Plant Material and Media Composition

Media was prepared based on a previously described protocol [26] modified as reported by Harlow et al. [27]. The medium was brought to an initial pH of 6.7 and autoclaved (121 °C, 103 kPa, 20 min) to remove O2. A carbonate buffer (4000 mg Na2CO3) was added after cooling under O2-free CO2. Medium (10 mL) was dispensed into Balch tubes containing 315 ± 0.06 mg plant material under anaerobic conditions before autoclaving for sterility. Plant material consisted of ground plant tissue (1-mm) from the orchardgrass cultivar ‘Prairie’, with sample collection and preparation as previously described by Kagan et al. [22]. Media reagents were purchased from Sigma-Aldrich (St. Louis, MO, USA).

2.2. Animals and Rumen Fluid Collection

All procedures and use of animals in this study were approved by the University of Kentucky Institutional Animal Care and Use Committee. Housing and animal care were consistent with the Guide to Care and Use of Agricultural Animals in Research and Teaching [28]. Four ram lambs [age = 134 ± 1 d; body weight (BW) = 44 ± 1 kg] and four Holstein steers (age = 212 ± 8 d; BW = 294 ± 4 kg) were selected from the University of Kentucky, Department of Animal and Food Sciences sheep flock and cattle herd at the C. Oran Little Research Center (Versailles, KY; geographic coordinates: 38°4′36″ N, 84°44′22″ W) as rumen fluid donors. All animals were fed the same cool-season grass hay diet containing predominantly tall fescue (Schedonorus arundinaceus [Schreb.] Dumort.) for 14 d prior to the experiment and had ad libitum access to conventional loose mineral and water. Chemical composition of tall fescue hay was as follows (100% DM basis): crude protein (CP) − 9.3%, acid detergent fiber (ADF; cellulose + lignin) − 42.9%, neutral detergent fiber (NDF; cellulose + hemicellulose + lignin) − 64.6%, non-fiber carbohydrate (NFC) − 16.8%, total digestible nutrients (TDN; estimated energy value) − 57.0%.
Rumen fluid obtained via oral intubation (500 mL) into the ventral sac of the rumen was transported to the laboratory in a sealed, insulated container within 1 h of collection. Rumen fluid (0.5 mL) was dispensed via tuberculin syringe into prepared anaerobic Balch tubes (as described above). Substrate-only and rumen fluid-only controls were also prepared. Tubes were incubated at 39 °C for 8 h. Aliquots were withdrawn using tuberculin syringes immediately after inoculation (time 0) and after 2, 4, and 8 h of incubation. Aliquots were also collected from rumen-fluid only and substrate-only controls at each timepoint. The pH was measured immediately (AccumetTM Research, AR10 pH meter; ThermoFisher Scientific, Waltham, MA, USA, with a Broadley-James® pH electrode; Broadley-James®, Irvine, CA, USA). Aliquots were then clarified by centrifugation (21,000× g, 2 min), with supernatants collected and stored at −20 °C for analyses of fructan profiles as well as mono and disaccharides. Aliquots from the 8-h timepoint were similarly collected and clarified to assess fermentation end-products (as described below).

2.3. Analysis of Fructans and Mono/Disaccharides

Clarified, frozen aliquots of supernatant from the incubations were thawed at 4 °C for 30 min and then allowed to finish thawing at ambient temperature. Aliquots were sonicated 5 min at 40 to 45 °C in a sonicating water bath (model 5800, Bransonics Corporation, Brookfield, CT, USA) to redissolve long-chain fructans, which can precipitate during freezing [29]. Sonicated samples were vortexed and then microcentrifuged (2 min; 13,000× g) to precipitate residual debris. Up to 1 mL supernatant was vacuum-filtered through a C18 endcapped solid-phase extraction column (Clean-Up, CEC18111, United Chemical Technologies, Bristol, PA, USA) packed with 100 mg sorbent and conditioned with 2 × 1 mL methanol followed by 3 × 1 mL water. Vacuum filtrates were stored at −20 °C until HPAEC analysis, at which time they were again thawed, sonicated 4 min at 40 to 45 °C, and diluted in water for chromatographic analysis (25-fold dilution for fructan profiling, and 50- to 100-fold dilution for analysis of mono and disaccharides).
Analysis was performed by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD) on an ICS-5000+ system (ThermoFisher Scientific, Waltham, MA, USA). Samples, controls, and standards (25 µL) were injected onto a ThermoFisher Scientific CarboPac PA200 column (5.5 µm particle size, 3 mm i.d. by 250 mm length) protected by a CarboPac PA200 guard column (3 mm i.d. by 50 mm length). Pulsed amperometric detection with a gold electrode [30] and quadruple-potential waveform [31] was employed. Eluents were 100 mM sodium hydroxide (solvent A; ThermoFisher Scientific, Waltham, MA, USA) and 1.0 M sodium acetate (Sigma-Aldrich, St. Louis, MO, USA) in 100 mM sodium hydroxide (solvent B). The gradient of Kagan et al. [12] was used, except that the gradient to 95% B/5% A and column wash at 95% B/5% A were changed to a gradient to and column wash at 100% B. The column and detector were kept at 28 °C, while the autosampler was at ambient temperature (generally 23 to 25 °C).

2.4. Quantification of Fructans and Mono/Disaccharides

Peak areas were determined with Chromeleon software (v. 7.2.10), using a minimum peak area of 0.08 nanocoulomb · min (nC · min). Standards were injected with every set of sample injections to check for peak retention times, and peaks were considered to be identical to standards if the retention times were within 0.07 min of each other. Purified orchardgrass fructan (DP ~7 to 70) and purified fructan of DP 4 to 12 from Sandberg’s bluegrass (Poa secunda J. Presl), as well as 1-kestose, 6-kestose, neokestose (6G-kestotriose), 1-nystose (1,1-kestotetraose), bifurcose ((1&6)-kestotetraose), and 6-nystose (6,6-kestotetraose), were a gift from P.A. Harrison (USDA Agricultural Research Service, Forage and Range Research Lab, Logan, UT, USA). Additional 1-nystose was purchased from Santa Cruz Biotechnology (Dallas, TX, USA). Short-chain and long-chain inulin (Orafti OPS and HP, respectively) were a gift from Beneo (Parsippany, NJ, USA). Chromatograms of rumen fluid injections were aligned with chromatograms of the corresponding orchardgrass-plus rumen fluid incubations. If peaks corresponding to standards were within 0.07 min of each other, the area of the peak from the rumen-only incubation was subtracted from the area of the peak from the corresponding orchardgrass-plus-rumen fluid incubation. Representative chromatograms from one rumen-fluid only control for each animal species can be found in Figure S1.
In order to estimate total fructan without hydrolysis, short-chain fructan was quantified by summing the area of peaks from 1-nystose (Figure S2a) to the last detectable peak in the series, corresponding approximately to the peak of DP 8 in the standard of short-chain fructans from P. secunda (Figure S2b). This range of peaks coincided roughly with the range of peaks in short-chain inulin (Figure S2c). Unknown peaks coeluting with peaks in the rumen fluid-only injections were excluded from the summation due to uncertainty about whether they were fructans or possibly other charged compounds. Peak area increased linearly with concentration, based on a standard curve of commercial short-chain inulin (y = 2.8919x + 0.1402, R2 = 0.9998). This method did not take into account the differences in response factors among fructans of different DP [32] (Table S1), but it allowed for an estimate of fructan concentrations within that region. Trisaccharides (1-kestose, 6-kestose, and neokestose or 6G-kestotriose), which eluted prior to 1-nystose, were not included in the quantification because that range included several peaks of uncertain identity (see Figure 1). Long-chain fructan was quantified by summing the area of peaks from above DP 8 (taken as 20.0 min) to 40 min or the end of the series of sharp peaks representing fructan (Figure S1b and Figure 1). As with short-chain fructan, peaks were excluded from the summation if they coeluted with peaks in the corresponding rumen fluid-only sample. Total peak area in the standard curve increased linearly with fructan concentration, based on a standard curve of commercial long-chain inulin (y = 1.2246x − 0.7401, R2 = 0.9984). The long-chain inulin ranged from about 20 to 38 min (Figure S1c). Orchardgrass fructan, used as a quantification standard by Kagan et al. [12], was available, but the long-chain inulin was chosen because it could be a generic standard for other grass species in the future.
Generally, fructans are considered as fructooligosaccharides (FOS) if the DP is 10 or less, and as fructans if the DP is above 10, although the DP above which FOS are considered fructans has been defined differently at times [8]. For the current study, due to the ranges encompassed by the standards, the FOS with a DP of 4 to 8 are referred to as short-chain fructans, and fructans with a higher DP are referred to as long-chain fructans.
Glucose and fructose were quantified with 9-point standard curves ranging from 0 to 30 µM monosaccharide. Sucrose was quantified with a 9-point standard curve ranging from 0 to 15 µM sucrose. Tri- and tetrasaccharides were quantified with 6- or 7-point standard curves ranging from 0 to 2 or 2.5 µM (See Table S1). Due to the fact that the above-described HPAEC-PAD method [12] was optimized for separation of fructans rather than for mono and disaccharides, partial co-elution of glucose and fructose coupled with variations in peak integration complicated accurate quantification of these individual analytes. Therefore, glucose, fructose, and sucrose were summed to give a total concentration of free sugars to compensate for the variability in individual concentrations.

2.5. Fermentation End-Product Analyses

Frozen aliquots were thawed and samples were transferred to a centrifugation filter unit (Amicon® Ultra-4 centrifugal filter unit, 30 kDa; EMD Millipore Corporation, Billerica, MA, USA) for clarification via centrifugation (1st step: 6000× g, 1 h, 4 °C; 2nd step: 21,000× g, 1 min, 4 °C). Lactate, succinate, acetate, propionate, butyrate, valerate, and isovalerate and/or methylbutyrate (IVMB) were quantified using HPLC (Summit HPLC; Dionex, Sunnyvale, CA, USA). Specifications of the HPLC system and analyses were as previously described by Weinert-Nelson et al. [24].

2.6. Chemical Composition of Orchardgrass Substrate

The chemical composition of the orchardgrass substrate material is shown in Table 1. In brief, crude protein was analyzed with a Leco FP-528 Nitrogen/Protein Analyzer (Leco Corporation, St. Joseph, MI, USA; Association of Official Analytical Chemists [AOAC] 990.03); ADF and NDF were analyzed with an ANKOM A200 Digestion Unit (ANKOM Technology, Macedon, NY, USA) (AOAC 973.18) [33,34]. Water-soluble carbohydrates (WSC) were extracted by shaking 100 mg tissue in 20 mL Millipore water for 1 h at ambient temperature [35]. Extracts were gravity-filtered through #4 Whatman (Buckinghamshire, UK) filters, brought to a final volume of 25 mL, and diluted 15-fold in water before being assayed by the phenol-sulfuric acid method of Dubois et al. [36], modified as described by Kagan et al. [35]. Ethanol-soluble carbohydrates (ESC) were extracted by shaking 100 mg tissue in 20 mL 80% ethanol for 4 h [37]. Extracts were filtered as described for WSC extracts and brought to 25 mL with 80% ethanol. Extracts were diluted 10-fold in water (final concentration of 8% ethanol) for storage at −20 °C until analysis [35]. Thawed extracts were diluted 1.5-fold in 8% ethanol (final dilution factor of 15) and assayed as for WSC extracts.

2.7. Statistical Analysis

All data were analyzed using PROC MIXED with the repeated statement in SAS (v. 9.4, SAS Inst. Inc., Cary, NC, USA). Animal was used as the experimental unit. Species, time, and the species × time interaction were set as fixed effects with animal as the random variable. The Kenward-Roger method was used to compute the denominator degrees of freedom for fixed effects and the repeated statement utilized the compound symmetry co-variance structure. Means were separated using the PDIFF option. Results were considered significant at p ≤ 0.05; trends were observed at p ≤ 0.10. Data are presented as least squares means ± SE.

3. Results

3.1. Orchardgrass Controls

Fructan and sugar profiles in the orchardgrass substrate incubated in the absence of rumen fluid at the start of the experiment (time 0) are characterized in Table 2 and Figure 1. Fructooligosaccharides consistently present included 6-kestose, bifurcose, and 6-nystose. Raffinose was often present, although in Figure 1, the peak was poorly resolved and too small to indicate on the chromatogram. Glucose, fructose, and sucrose were abundant in orchardgrass at the start of the incubations, and glucose and fructose partially coeluted.

3.2. Fructan Degradation

Representative chromatograms from one iteration with each animal species are shown in Figure 2. Long-chain fructan (DP > 8) concentrations and rates of disappearance were influenced by ruminant species and time (p ≤ 0.04; Table 3; Figure 2). The analyzed means are presented in Table 3, with rates of change shown in Figure 3a. Initial long-chain fructan concentrations were 414.9 ± 13.9 μg mL−1 regardless of species (p = 0.48). While there was a 3-fold more rapid degradation of long-chain fructan in ovine (–381.8 ± 83.6 μg h−1) than bovine (–131.8 ± 83.6 μg h−1) fermentations from 0–2 h of incubation (p = 0.05), the rate of fructan disappearance was greater in bovine fermentations from 2–4 h (bovine: –2011.0; ovine: –1569.6; SE: 84.4 μg h−1; p < 0.01; Figure 3a). No detectable long-chain fructans remained in bovine fermentations at the 4 h timepoint. In contrast, long-chain fructans were still present in ovine fermentations at 4 h, and there was continued degradation through 8 h (–131.7 ± 83.6 μg h−1), at which point long-chain fructans could no longer be detected.
Short-chain fructan (DP 4 to 8) concentrations and rates of appearance/disappearance were also influenced by species and time (p < 0.01; Table 3; Figure 3b). Initial short-chain fructan concentrations were 69.8 ± 16.2 μg mL−1 regardless of ruminant species (p = 0.27). Short-chain fructans increased from 0 to 2 h of incubation, with no difference between species in rate of appearance (bovine: +691.6; ovine: +892.7; SE 119.4 μg h−1; p = 0.29). Short-chain fructan concentrations began decreasing after 2 h in bovine fermentations (p < 0.01), but degradation was delayed in ovine fermentations, with a reduction in short-chain fructan concentrations occurring only after the 4 h time point (p < 0.01). Consequently, the rate of degradation was 26-fold more rapid in bovine (–873.7 ± 119.4 μg h−1) than ovine (–33.8 ± 119.4 μg h−1) fermentations from 2 to 4 h (p < 0.01), and the concentration of short-chain fructan at 4 h was reduced to 19.0 ± 21.5 μg mL−1. Degradation of short-chain fructans increased 19-fold from 4 to 8 h in ovine fermentations (–625.5 ± 120.5 μg h−1; p < 0.01). This increased degradation in ovine fermentations from 4 to 8 h matched the rates occurring in bovine fermentations early in the incubation during the 2 to 4 h period (p = 0.25). Only minimal short-chain fructans remained at 8 h in either bovine or ovine fermentations (p = 0.82).
The individual tetrasaccharides 1-nystose (DP 4) and bifurcose (DP 4) were influenced by species and time (p < 0.01; Table 3). Initial concentrations of 1-nystose and bifurcose were 2.6 ± 2.3 μg mL−1 and 13.8 ± 7.2 μg mL−1, respectively, regardless of species (p ≥ 0.72). Concentrations of 1-nystose and bifurcose at 4 h were greater in ovine (1-nystose: 30.3 ± 3.3 μg mL−1; bifurcose: 85.8 ± 9.6 μg mL−1) than bovine (1-nystose: 0.7 ± 3.3 μg mL−1; bifurcose: 2.1 ± 9.6 μg mL−1; p ≤ 0.01) fermentations but did not differ at other timepoints across the incubation (p ≥ 0.43). By 8 h, 1-nystose was undetectable in all bovine and ovine fermentations. Bifurcose was also undetectable in bovine fermentations at 8 h, but small amounts were variably present in ovine fermentations (4.8 ± 9.6 μg mL−1).
For all other short-chain fructans evaluated, there was no effect of species (p ≥ 0.40) or species by time interaction (p ≥ 0.06; Table 3). The fructans 6-kestose (DP 3), neokestose (DP 3), and 6-nystose (DP 4) did, however, vary by time (p ≤ 0.05). Both 6-kestose and 6-nystose were present in initial bovine and ovine fermentations (6-kestose: 6.5 ± 1.3 μg mL−1; 6-nystose: 8.0 ± 1.0 μg mL−1). Concentrations decreased after 4 h of incubation to near undetectable levels by 8 h (6-kestose: 0.4 ± 1.0 μg mL−1; 6-nystose: 1.1 ± 1.0 μg mL−1; p < 0.01). In contrast, neokestose was undetectable in initial fermentations for both species, with minimal accumulation by 2 h (0.3 ± 0.2 μg mL−1; p = 0.02) before again decreasing to undetectable concentrations by the end of the incubation at 8 h.
Free sugar concentrations (glucose, fructose, and sucrose combined) also did not differ by species (p < 0.85). While there was no species by time interaction (p = 0.11; Table 3), free sugars did differ by time (p < 0.01). Maximum concentrations of free sugars (1094.4 ± 50.4 μg mL−1) were found at 0 h, regardless of species (p < 0.01). Free sugar concentrations decreased by 2 h (p < 0.01) and again by 4 h (p < 0.01). Free sugars also tended to decrease by 8 h (p = 0.06), at which time free sugar concentrations had decreased by 83% from initial concentrations.

3.3. Fermentation pH

Fermentation pH was influenced by ruminant species and time (p < 0.01; Figure 4). Initial pH was 6.43 ± 0.03 regardless of species (p = 0.31), and pH also did not differ between species at 2 h (bovine: 6.57; ovine 6.56; SE: 0.03; p = 0.48). At 4 h, however, there was a species difference (p < 0.01), with pH below baseline levels in bovine fermentations (6.34 ± 0.03; p < 0.01), but not in ovine fermentations (6.43 ± 0.03; p = 0.54). In contrast, pH was lower in ovine (6.17 ± 0.02) than bovine (6.26 ± 0.03) fermentations at 8 h (p < 0.01).

3.4. Fermentation End-Products

Total SCFA concentration at 8 h (end of incubation period) differed by species, with total SCFA in bovine fermentations 19% greater than in ovine fermentations (p = 0.03; Table 4). The species difference in total SCFA was driven primarily by differences in acetate concentrations, which were 23% greater in bovine versus ovine fermentations (p < 0.01). There was no effect of species for other SCFA including succinate, lactate, propionate, and butyrate (p ≥ 0.12). Valerate and IVMB were below the limit of quantification in all samples.

4. Discussion

Sheep are commonly used as model ruminants in health, nutrition, and rumen fermentation research due to advantages related to overall animal size as well as length of production cycles. However, previous research has suggested that the rumen microbial communities of sheep and cattle may differ both in composition and function, including differences in the fermentation of non-structural carbohydrates [24,38,39]. Both sheep and cattle may be managed on high-forage diets including in pasture-based systems. Given that cool-season pasture grasses have the potential to accumulate substantial amounts of fructans as the primary form of storage carbohydrate [1,16], this study sought to characterize potential differences in ruminal fructan fermentation between these animal species.
Orchardgrass was utilized as a model cool-season grass substrate. Orchardgrass fructans have previously been characterized as primarily linear with β(2 → 6) linkages and a terminal glucose residue [40,41]. On a dry matter (DM) basis, the concentrations of fructan in the orchardgrass controls (13.4 ± 1.3 mg g−1) were within ranges reported in prior studies for orchardgrass harvested at a similar time of year (early May) within the same geographic region [12,42], and thus was a suitable substrate material for this ex vivo study. Presence of specific tetra- and trisaccharides including 1-nystose, 1-kestose, neokestose, and raffinose has also been documented in lyophilized orchardgrass tissue extracted on a larger scale [40,43]. In the current study, consistent detection of tetra- and trisaccharides in the orchardgrass controls was impacted by the relatively low concentrations of these analytes, the small amount of tissue used as substrate, and the dilution required to prevent overloading the column for the HPAEC analyses. However, peak detection improved as concentrations increased during incubation with rumen fluid.
Results of this study illustrated general similarities in ex vivo ruminal fermentation of orchardgrass fructans between cattle and sheep. In both ovine and bovine fermentations, degradation of long-chain fructans began during the initial 2h period, with the fastest rate of disappearance occurring between 2 and 4 h of incubation. In contrast, short-chain fructans accumulated in both species from 0 to 2 h, with subsequent degradation throughout the remainder of the incubation period. This was true both for overall short-chain fructans as well as for the tetrasaccharides bifurcose and 1-nystose. These results are in agreement with previous findings by Ince et al. [20] and Kagan et al. [22], both of which reported initial decreases in long-chain fructans coupled with concomitant increases in short-chain fructans in fermentations using equine foregut digesta and bovine rumen fluid, respectively, as the inoculum.
Despite these general similarities, this study also clearly revealed interspecies differences in fructan fermentation. While the rumen microbiota of both cattle and sheep were able to fully utilize fructans available in the orchardgrass substrate in the current study, the overall degradation of long-chain fructans in bovine fermentations proceeded more rapidly than in ovine fermentations. There was also a marked delay of short-chain fructan catabolism in ovine fermentations, with substantial degradation occurring only after 4 h of incubation. Catabolism of orchardgrass fructans by bovine fermentations in the current study proceeded at rates similar to those previously documented by Kagan et al. [22] and Hall and Weimer [18]. To the authors’ knowledge, no previous studies have evaluated catabolism of orchardgrass fructans by the rumen microbiota of sheep. Species differences in this study also align with those of Biggs and Hancock [25], who found that both inulin and bacterial levan were degraded 2–3 times more slowly when rumen fluid harvested from sheep was used as the inoculum in comparison with fermentations using bovine rumen fluid. Species differences in short-chain fructan catabolism in the present study were also reflected in patterns of appearance and disappearance of bifurcose and 1-nystose, but not in free sugar concentrations. A more frequent sampling interval and employing techniques to improve resolution of glucose, fructose, and sucrose peaks such as the method recently developed by Joyce et al. [44] could aid in characterizing fluctuations in these mono and disaccharides during catabolism of grass fructans. It should be noted that in bovine fermentations in the current study, the greatest degradation of both long- and short-chain fructans occurred during the 2–4 h period, with long-chain fructans non-detectable and short-chain fructans almost completely utilized by 4 h. It is therefore possible that the capacity of the bovine rumen microbiota to catabolize grass fructans exceeded the fructans provided by the orchardgrass used as the fermentative substrate in this experiment.
Species similarities and differences in catabolism of long- and short-chain fructans may provide information about the fructan hydrolytic enzymes that dominate in the bovine and ovine ruminal microbiota. Exohydrolases such as the β-fructofuranosidases produced by Butyrivibrio fibrisolvens (EC 3.2.1.26) or Lactobacillus plantarum (EC 3.2.1.80) hydrolyze the fructosyl linkages in fructan polymers, releasing terminal residues [23,45]. In contrast, endohydrolases such as the 2,6-β-D-fructan fructanohydrolase (E.C. 3.2.1.65) produced by Treponema saccharophilum and T. zioleckii can cleave at any point within the fructan chain [46,47]. Kagan et al. [22] suggested that the pattern of fructan catabolism in which initial decreases in long-chain fructan concentrations is coupled with increases in short-chain fructans was due to combined actions of microbial exo- and endohydrolases, and as described above, this overall pattern of degradation occurred in both species in the current study. Additionally, increases in concentrations of bifurcose and 1-nystose in both bovine and ovine fermentations suggest the hydrolysis of longer fructans, as both tetrasaccharides (the former branched, and the latter linear) were found by Chatterton et al. [40] in acid-hydrolyzed DP 5 fructan from orchardgrass. Fructans of DP > 5 in orchardgrass are of the levan type, with linear β(2 → 6)-linked fructosyl units and a terminal glucose [40]. The increase in bifurcose, a branched fructooligosaccharide, over time could have been due to hydrolysis of the branched DP 5 fructans by ruminal microbiota. The species differences in fructan degradation and utilization capacity may also have been driven by differences in fructanolytic enzymes produced by the rumen microbiota. The rapid disappearance of long-chain fructan in bovine fermentations between 2 and 4 h may indicate the presence of more active and/or ubiquitous endolevanases such as EC 3.2.1.65, which preferentially cleaves β(2 → 6)linked fructose residues [47]. This endolevanase may have been less predominant in ovine ruminal microbiota, given that the more gradual disappearance of long- and short-chain fructans in ovine fermentations resembled the gradual 2-h disappearance of long-chain fructan in timothy (Phleum pratense L.) incubated with a fructan exohydrolase from L. plantarum [45]. Given that catabolism of short-chain fructans also proceeded at a faster rate in bovine fermentations, exohydrolase activity may also differ between cattle and sheep. Differences in the enzymes present may also suggest differences in the dominant fructanolytic microorganisms of the bovine and ovine fermentations. While not evaluated as part of this study, further research is warranted to evaluate the extent to which potential differences in microbial community composition and microbial enzymes contribute to species differences in fructan utilization.
Species differences in fructan degradation were also reflected in fermentation pH. Catabolism of fructans was greater in bovine incubations from 2–4 h, and this corresponded with lower fermentation pH at 4 h. Accordingly, greater catabolism of fructans occurred in ovine incubations from 4–8 h, with a lower pH in ovine fermentations at 8 h. Although statistically significant, the magnitude of these species differences in pH were small, regardless of timepoint (≤0.10) and thus were likely not physiologically relevant. Fermentation pH remained above 6.0 throughout the incubation period for both species, well above the threshold where dysbiosis of the microbial community related to ruminal acidosis (pH < 5.0) or subacute acidosis (pH: 5.0–5.6) is known to occur [48,49,50] or at which viability of cellulolytic bacteria would be compromised (pH < 6.0) [51,52,53]. However, the minimal reductions and limited magnitude of species differences in fermentation pH in the current study are at least partly attributable to the heavily buffered nature of the medium used in the incubations, and it is therefore possible that ruminal pH differences during in vivo degradation of fructans may be more pronounced. Kasperowicz et al. [54] reported decreased post-prandial ruminal pH in sheep fed a diet including high-fructan grass hay versus a low-fructan control, with pH also remaining above 6.0 regardless of treatment. However, to the authors’ knowledge, no similar studies have been conducted in cattle. Additionally, even if changes in rumen pH in response to fermentation of fructans are not substantial, differences in fermentation end-product production and pH changes within microenvironments could impact overall microbial community stability and function.
While additional research would be required to confirm the species differences identified in the current study in vivo, these results suggest that fructan utilization capacity may be greater in cattle. This has the potential to impact total utilization of fructans. The principal end-product of microbial degradation of fructans is lactate [18,21], but production of comparatively small amounts of other SCFA has been reported when inulin was fermented by Streptococcus bovis [21] and when inulin or phlein were incubated with bovine rumen fluid [18]. Lactate produced during ruminal fermentation can then be catabolized by lactate-utilizing bacteria such as Megasphaera elsdenii and Selenomonas ruminantium, producing propionate, acetate, and butyrate [50,55]. Once absorbed, these fermentation end-products can be utilized to meet energy requirements of the ruminant animal [48,49,50]. Therefore, if cattle possess a superior ability to utilize dietary fructans, in turn, cattle would be able to harvest more energy from these dietary non-structural carbohydrates. A greater capacity for fructan utilization in cattle may potentially be related to historical differences in diet between species. Cuchillo-Hilario et al. [56] reported that sheep sought more nitrogen and less carbon when selectively grazing forages within a pasture sward, an association not observed in cattle grazing similar pastures. Additionally, Grant et al., [57] found that cattle maintained on pasture consumed more grasses and fewer forbs in comparison with sheep, which was attributed to a combination of differences in forage preference and selection ability due to anatomical differences in the size of mouthparts and mechanism of prehension. Cattle and sheep evolved as grazers [58], and it is possible that cattle and their rumen microbiota had greater exposure to dietary fructans over the course of this evolution due to greater consumption of grasses and that, in response, the rumen microbial community of cattle adapted to more efficiently utilize fructans present in the diet. In modern animal management systems, this adaptation would still confer an advantage to pasture-fed cattle and those maintained on other high-forage diets in which cool-season grasses would constitute a large proportion of the total diet, allowing cattle to better utilize grass fructans to support energy requirements for production.
Fermentation of the orchardgrass substrate in the current study produced metabolic products including lactate, acetate, propionate, and butyrate that were similar in concentration to previously reported values for ex vivo fermentation of tall fescue by bovine and ovine rumen bacteria [24]. In contrast to these prior studies, in which purified fructans were utilized as the fermentative substrate, the orchardgrass used in the current study was a complex substrate that provided other non-structural and structural carbohydrates as well as proteins that would have been subject to degradation by the rumen microbiota. Additionally, samples could only be collected at the end of the incubation period for SCFA analysis to ensure adequate volume during the fermentations, by which point all long-chain and almost all short-chain fructans had been completely utilized in both bovine and ovine incubations. It is possible that fluctuations in SCFA concentrations and differences between bovine and ovine fermentations could have occurred earlier in the incubations. Therefore, additional research is needed to more directly evaluate potential species differences in production of metabolic end-products from fermentation of long- and short-chain fructans. Despite the minimal differences in SCFA, chromatographic profiling of fructan catabolism in the current study conclusively demonstrated species differences in degradation of both long- and short-chain fructans and indicate that cattle may possess a greater capacity for utilization of grass fructans.

5. Conclusions

While there were general similarities in the pattern of fructan catabolism by the ruminal microbiota of cattle and sheep, the rate of fructan degradation varied dramatically. In both species, there was evidence of initial degradation of long-chain orchardgrass fructans to short-chain fructans. Short-chain fructans accumulated early in the incubation period (within the first two hours), prior to subsequent degradation in both bovine and ovine fermentations. However, the rate at which fructans were catabolized was more rapid in bovine fermentations, with almost all available grass fructans completely utilized after only four hours. In contrast, degradation of long-chain fructans was slower in ovine fermentations and there was a notable delay in catabolism of short-chain fructans, with fructans not fully utilized until the end of the incubation period. Based on these results, fructan utilization capacity appears to be greater in cattle. Future research should focus on better understanding potential differences in the composition and function of the fructanolytic guild in the rumen of cattle and sheep, particularly as it relates to the influence of fructan DP. The findings in this study also indicate that additional research is warranted to evaluate in vivo ruminal fructan utilization and to determine implications for nutritional management and production of cattle and sheep.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/fermentation9110925/s1. Table S1: Calibration equations used for mono/disaccharide and fructan standards; Figure S1: Representative chromatograms showing fructan profiles in rumen fluid-only controls following incubation with rumen fluid from one steer (a) or ram lamb (b). The areas of the isolated peaks present in controls were subtracted from the areas of peaks in the chromatograms of orchardgrass fermentations. Chromatograms were obtained by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD); Figure S2: Chromatograms of the standards used in this study, including (a) a mixture of glucose (G), fructose (F), sucrose (S), trisaccharides raffinose (R), 1-kestose (K), 6-kestose (6K), neokestose (nK), 1-nystose or nystose (N), bifurcose (Bf), and 6-nystose (6N); (b) a mixture of fructans from big bluegrass (Poa secunda J. Presl), DP 4 to 12 (numbers above corresponding peaks), and fructans from orchardgrass (Dactylis glomerata L.) of DP ranging from about 7 to 69, used to determine DP in samples; and (c) a mixture of short-chain (SC) and long-chain (LC) fructan from chicory, used to quantify short- and long-chain fructans. The 1-nystose (N) peak is indicated by an arrow. Chromatograms were obtained by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD).

Author Contributions

Conceptualization: J.R.W.-N., I.A.K., D.G.E. and B.E.D.; data curation: J.R.W.-N., I.A.K., M.D.F. and B.E.D.; formal analysis: J.R.W.-N., I.A.K. and B.E.D.; investigation: J.R.W.-N., I.A.K. and B.E.D.; methodology: J.R.W.-N., I.A.K., D.G.E., M.D.F. and B.E.D.; project administration: J.R.W.-N., I.A.K. and B.E.D.; resources: I.A.K., D.G.E., M.D.F. and B.E.D.; supervision: I.A.K., M.D.F. and B.E.D.; visualization: J.R.W.-N. and I.A.K.; writing—original draft: J.R.W.-N., I.A.K. and B.E.D.; writing—review and editing: J.R.W.-N., I.A.K., D.G.E., M.D.F. and B.E.D. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the USDA-ARS National Program 101, Food Animal Production Project (#: 5042-21000-004-00D). This research was supported in part by the ARS Administrator Funded Research Associate Program (J.R. Weinert-Nelson; Project #: 0101-88888-016-00D).

Institutional Review Board Statement

The animal study protocols were approved by the Institutional Animal Care and Use Committee at the University of Kentucky (protocols: #2020-3546, 6/2/2020; #2021-3772, 3/26/2021).

Data Availability Statement

All data generated and analyzed for this study are available from the authors upon request.

Acknowledgments

The authors acknowledge the technical assistance of Matthew Hamilton, LeeAnn Jacks, and Katherine Parsley, University of Kentucky Department of Animal and Food Sciences, and Tracy Hamilton, Brenda Coe, Jessica Ferrell, and Jacob Ibarra, USDA-Agricultural Research Service (ARS), Forage-Animal Production Research Unit. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Hendry, G.A.F.; Wallace, R.K. The origin, distribution, and evolutionary significance of fructans. In Science and Technology of Fructans; Suzuki, M., Chatterton, N.J., Eds.; CRC Press, Inc.: Boca Raton, FL, USA, 1993; pp. 119–139. [Google Scholar]
  2. Pollock, C.J. Tansley review no. 5 fructans and the metabolism of sucrose in vascular plants. New Phytol. 1986, 104, 1–24. [Google Scholar] [CrossRef]
  3. Livingston, D.P.; Hincha, D.K.; Heyer, A.G. Fructan and its relationship to abiotic stress tolerance in plants. Cell Mol. Life Sci. 2009, 66, 2007–2023. [Google Scholar] [CrossRef]
  4. Ernst, M.; Chatterton, N.J.; Harrison, P.A. Purification and characterization of a new fructan series from species of Asteraceae. New Phytol. 1996, 132, 63–66. [Google Scholar] [CrossRef]
  5. French, A.D.; Waterhouse, A.L. Chemical structure and characteristics. In Science and Technology of Fructans; Suzuki, M., Chatterton, N.J., Eds.; CRC Press, Inc.: Boca Raton, FL, USA, 1993; pp. 119–139. [Google Scholar]
  6. Vijn, I.; Smeekens, S. Fructan: More than a reserve carbohydrate? Plant Physiol. 1999, 120, 351–359. [Google Scholar] [CrossRef]
  7. Waterhouse, A.L.; Chatterton, N.J. Glossary of fructan terms. In Science and Technology of Fructans; Suzuki, M., Chatterton, N.J., Eds.; CRC Press, Inc.: Boca Raton, FL, USA, 1993; pp. 119–139. [Google Scholar]
  8. Benkeblia, N. Fructooligosaccharides and fructans analysis in plants and food crops. J. Chromatogr. A 2013, 1313, 54–61. [Google Scholar] [CrossRef] [PubMed]
  9. Pollock, C.J.; Jones, T. Seasonal patterns of fructan metabolism in forage grasses. New Phytol. 1979, 83, 9–15. [Google Scholar] [CrossRef]
  10. Suzuki, M. Fructans in forage grasses with varying degrees of coldhardiness. J. Plant Physiol. 1989, 134, 224–231. [Google Scholar] [CrossRef]
  11. Pavis, N.; Boucaud, J.; Prud’Homme, M.P. Fructans and fructan-metabolizing enzymes in leaves of Lolium perenne. New Phytol. 2001, 150, 97–109. [Google Scholar] [CrossRef]
  12. Kagan, I.A.; Kirch, B.H.; Thatcher, C.D.; Strickland, J.R.; Elvinger, F.; Teutsch, C.D.; Pleasant, S.R. Seasonal and diurnal variation in simple sugar and fructan composition of orchardgrass pasture and hay in the Piedmont region of the United States. J. Equine Vet. Sci. 2011, 31, 488–497. [Google Scholar] [CrossRef]
  13. Waite, R.; Boyd, J. The water-soluble carbohydrates of grasses: Part I. Changes occurring during the normal life-cycle. J. Sci. Food Agric. 1953, 4, 197–204. [Google Scholar] [CrossRef]
  14. Volaire, F.; Lelièvre, F. Production, persistence, and water-soluble carbohydrate accumulation in 21 contrasting populations of Dactylis glomerata L. subjected to severe drought in the south of France. Aust. J. Agric. Res. 1997, 48, 933–944. [Google Scholar] [CrossRef]
  15. Chatterton, N.J.; Harrison, P.A.; Bennett, J.H.; Asay, K.H. Carbohydrate partitioning in 185 accessions of Gramineae grown under warm and cool temperatures. J. Plant Physiol. 1989, 134, 169–179. [Google Scholar] [CrossRef]
  16. Jensen, K.B.; Harrison, P.; Chatterton, N.J.; Bushman, B.S.; Creech, J.E. Seasonal trends in nonstructural carbohydrates in cool-and warm-season grasses. Crop. Sci. 2014, 54, 2328–2340. [Google Scholar] [CrossRef]
  17. Thomas, G.J. Metabolism of the soluble carbohydrates of grasses in the rumen of the sheep. J. Agric. Sci. 1960, 54, 360–372. [Google Scholar] [CrossRef]
  18. Hall, M.B.; Weimer, P.J. Divergent utilization patterns of grass fructan, inulin, and other nonfiber carbohydrates by ruminal microbes. J. Dairy Sci. 2016, 99, 245–257. [Google Scholar] [CrossRef]
  19. Perrin, S.; Fougnies, C.; Grill, J.P.; Jacobs, H.; Schneider, F. Fermentation of chicory fructo-oligosaccharides in mixtures of different degrees of polymerization by three strains of bifidobacteria. Can. J. Microbiol. 2002, 48, 759–763. [Google Scholar] [CrossRef]
  20. Ince, J.C.; Longland, A.C.; Moore-Colyer, M.J.S.; Harris, P.A. In vitro degradation of grass fructan by equid gastrointestinal digesta. Grass Forage Sci. 2013, 69, 514–523. [Google Scholar] [CrossRef]
  21. Harlow, B.E.; Kagan, I.A.; Lawrence, L.M.; Flythe, M.D. Effects of inulin chain length on fermentation by equine fecal bacteria and Streptococcus bovis. J. Equine Vet. Sci. 2017, 48, 113–120. [Google Scholar] [CrossRef]
  22. Kagan, I.A.; Harlow, B.E.; Flythe, M.D. A chromatographic method to monitor fructan catabolism in two cool-season grasses fermented by mixed bovine ruminal microbiota. JSFA Rep. 2022, 2, 264–271. [Google Scholar] [CrossRef]
  23. Ziołecki, A.; Guczynska, W.; Wojciechowicz, M. Some rumen bacteria degrading fructan. Lett. Appl. Microbiol. 1992, 15, 244–247. [Google Scholar] [CrossRef]
  24. Weinert-Nelson, J.R.; Ely, D.G.; Flythe, M.D.; Hamilton, T.A.; Ferrell, J.L.; Davis, B.E. Ex vivo fermentation of hay and corn by rumen bacteria from cattle and sheep. Fermentation 2023, 101, 66–67. [Google Scholar]
  25. Biggs, D.R.; Hancock, K.R. In vitro digestion of bacterial and plant fructans and effects on ammonia accumulation in cow and sheep rumen fluids. J. Gen. Appl. Microbiol. 1998, 44, 167–171. [Google Scholar] [CrossRef] [PubMed]
  26. Stack, R.J.; Hungate, R.E.; Opsahl, W.P. Phenylacetic acid stimulation of cellulose digestion by Ruminococcus albus 8. Appl. Environ. Microbiol. 1983, 46, 539–544. [Google Scholar] [CrossRef] [PubMed]
  27. Harlow, B.E.; Flythe, M.D.; Aiken, G.E. Effect of biochanin A on corn grain (Zea mays) fermentation by bovine rumen amylolytic bacteria. J. Appl. Microbiol. 2017, 122, 870–880. [Google Scholar] [CrossRef]
  28. FASS. Guide for the Care and Use of Agricultural Animals in Research and Teaching; FASS Inc.: Champaign, IL, USA, 2020. [Google Scholar]
  29. Chatterton, N.J.; Harrison, P.A. Fructan oligomers in Poa ampla. New Phytol. 1997, 136, 3–10. [Google Scholar] [CrossRef]
  30. Rocklin, R.D.; Pohl, C.A. Determination of carbohydrates by anion exchange chromatography with pulsed amperometric detection. J. Liq. Chromatogr. 1983, 6, 1577–1590. [Google Scholar] [CrossRef]
  31. Rocklin, R.D.; Clarke, A.P.; Weitzhandler, M. Improved long-term reproducibility for pulsed amperometric detection of carbohydrates via a new quadruple-potential waveform. Anal. Chem. 1998, 70, 1496–1501. [Google Scholar] [CrossRef]
  32. Pöhnl, T.; Böttcher, C.; Schulz, H.; Stürtz, M.; Widder, S.; Carle, R.; Schweiggert, R.M. Comparison of high performance anion exchange chromatography with pulsed amperometric detection (HPAEC-PAD) and ultra-high performance liquid chromatography with evaporative light scattering (UHPLC-ELSD) for the analyses of fructooligosaccharides in onion (Allium cepa L.). J. Food Comp. Anal. 2017, 63, 48–156. [Google Scholar] [CrossRef]
  33. ANKOM Technology. Method 6: Acid Detergent Fiber in Feeds—Filter Bag Technique (for A200 and A2001); ANKOM Technology: Macedon, NY, USA, 2005; Available online: https://www.ankom.com/analytical-methods-support/fiber-analyzer-a200?f%5B0%5D=field_faq_group%3A89#:~:text=Acid-,Detergent,-Fiber%20Method%20(A200) (accessed on 26 July 2022).
  34. ANKOM Technology. Method 6: Neutral Detergent Fiber in Feeds—Filter Bag Technique (for A200 and A2001); ANKOM Technology: Macedon, NY, USA, 2006; Available online: https://www.ankom.com/analytical-methods-support/fiber-analyzer-a200?f%5B0%5D=field_faq_group%3A89#:~:text=View%20PDF-,Neutral,-Detergent%20Fiber%20Method (accessed on 26 July 2022).
  35. Kagan, I.A.; Lawrence, L.M.; Seman, D.H.; Prince, K.J.; Fowler, A.L.; Smith, S.R. Effects of sampling time, cultivar, and methodology on water- and ethanol-soluble carbohydrate profiles of three cool-season grasses. J. Equine Vet. Sci. 2018, 61, 99–107. [Google Scholar] [CrossRef]
  36. DuBois, M.; Gilles, K.A.; Hamilton, J.K.; Rebers, P.A.; Smith, F. Colorimetric method for determination of sugars and related substances. Anal. Chem. 1956, 28, 350–356. [Google Scholar] [CrossRef]
  37. Hall, M.B.; Hoover, W.H.; Jennings, J.P.; Webster, T.K.M. A method for partitioning neutral detergent-soluble carbohydrates. J. Sci. Food Agric. 1999, 79, 2079–2086. [Google Scholar] [CrossRef]
  38. Li, J.; Yan, H.; Chen, J.; Duan, C.; Guo, Y.; Liu, Y.; Zhang, Y.; Ji, S. Correlation of ruminal fermentation parameters and rumen bacterial community by comparing those of the goat, sheep, and cow in vitro. Fermentation 2022, 8, 427. [Google Scholar] [CrossRef]
  39. Henderson, G.; Cox, F.; Ganesh, S.; Jonker, A.; Young, W.; Janssen, P.H. Rumen microbial community composition varies with diet and host, but a core microbiome is found across a wide geographical range. Sci. Rep. 2015, 5, 14567. [Google Scholar] [CrossRef] [PubMed]
  40. Chatterton, N.J.; Harrison, P.A.; Thornley, W.R.; Bennett, J.H. Structures of fructan oligomers in orchardgrass (Dactylis glomerata L.). J. Plant Physiol. 1993, 142, 552–556. [Google Scholar] [CrossRef]
  41. Cairns, A.J.; Nash, R.; Machado de Carvalho, M.-A.; Sims, I.M. Characterization of the enzymatic polymerization of 2,6-linked fructan by leaf extracts from timothy grass (Phleum pratense). New Phytol. 1999, 142, 79–91. [Google Scholar] [CrossRef]
  42. Kagan, I.A.; Kirch, B.H.; Strickland, J.R. A chromatographic survey of methods for extracting long-chain grass fructans. Grass Forage Sci. 2011, 66, 434–448. [Google Scholar] [CrossRef]
  43. Chatterton, N.J.; Harrison, P.A.; Thornley, W.R.; Draper, E.A. Oligosaccharides in foliage of Agropyron, Bromus, Dactylis, Festuca, Lolium and Phleum. New Phytol. 1990, 114, 67–171. [Google Scholar] [CrossRef]
  44. Joyce, G.E.; Kagan, I.A.; Flythe, M.D.; Davis, B.E.; Schendel, R.R. Profiling of cool-season forage arabinoxylans via a validated HPAEC-PAD method. Front. Plant Sci. 2023, 14, 1116995. [Google Scholar] [CrossRef]
  45. Müller, M.; Seyfarth, W. Purification and substrate specificity of an extracellular fructanhydrolase from Lactobacillus paracasei ssp. paracasei P 4134. New Phytol. 1997, 136, 89–96. [Google Scholar] [CrossRef]
  46. Kasperowicz, A.; Michalowski, T. Assessment of the fructanolytic activities in the rumen bacterium Treponema saccharophilum strain S. J. Appl. Microbiol. 2002, 92, 140–146. [Google Scholar] [CrossRef]
  47. Kasperowicz, A.; Pristaš, P.; Piknová, M.; Javorský, P.; Guczyńska, W.; Michałovski, T.; Kwiatkowska, E. Fructanolytic and saccharolytic enzymes of Treponema zioleckii strain kT. Anaerobe 2010, 16, 387–392. [Google Scholar] [CrossRef] [PubMed]
  48. Owens, F.; Secrist, D.; Hill, W.; Gill, D. Acidosis in cattle: A review. J. Anim. Sci. 1998, 76, 275–286. [Google Scholar] [CrossRef] [PubMed]
  49. Krause, K.M.; Oetzel, G.R. Understanding and preventing subacute ruminal acidosis in dairy herds: A review. Anim. Feed Sci. Tech. 2006, 126, 215–236. [Google Scholar] [CrossRef]
  50. Nagaraja, T.; Titgemeyer, E. Ruminal acidosis in beef cattle: The current microbiological and nutritional outlook 1, 2. J. Dairy Sci. 2007, 90, E17–E38. [Google Scholar] [CrossRef]
  51. Russell, J.B.; Dombrowski, D. Effect of pH on the efficiency of growth by pure cultures of rumen bacteria in continuous culture. Appl. Environ. Microbiol. 1980, 39, 604–610. [Google Scholar] [CrossRef] [PubMed]
  52. Shi, Y.; Weimer, P. Response surface analysis of the effects of pH and dilution rate on Ruminococcus flavefaciens FD-1 in cellulose-fed continuous culture. Appl. Environ. Microbiol. 1992, 58, 2583–2591. [Google Scholar] [CrossRef] [PubMed]
  53. Weimer, P.J. Effects of dilution rate and pH on the ruminal cellulolytic bacterium Fibrobacter succinogenes S85 in cellulose-fed continuous culture. Arch. Microbiol. 1993, 160, 288. [Google Scholar] [CrossRef]
  54. Kasperowicz, A.; Stan-Głasek, K.; Kowalik, B.; Vandzurova, A.; Pristas, P.; Pająk, J.; Kwiatkowska, E.; Michałowski, T. Effect of dietary fructose polymers or sucrose on microbial fermentation, enzyme activity, ciliate concentration and diversity of bacterial flora in the rumen of rams. Anim. Feed Sci. Technol. 2014, 195, 38–46. [Google Scholar] [CrossRef]
  55. Fan, Y.; Xia, G.; Jin, Y.; Wang, H. Ambient pH regulates lactate catabolism pathway of the ruminal Megasphaera elsdenni BE2-2083 and Selenomonas ruminantium HD4. J. Appl. Microbiol. 2022, 132, 2661–2672. [Google Scholar] [CrossRef]
  56. Cuchillo-Hilario, M.; Wrage-Mönnig, N.; Isselstein, J. Forage selectivity by cattle and sheep co-grazing swards differing in plant species diversity. Grass Forag Sci. 2018, 73, 320–329. [Google Scholar] [CrossRef]
  57. Grant, S.A.; Suckling, D.E.; Smith, H.K.; Torvell, L.; Forbes, T.D.A.; Hodgson, J. Comparative studies of diet selection by sheep and cattle: The hill grasslands. J. Ecol. 1985, 73, 987–1004. [Google Scholar] [CrossRef]
  58. Rutter, S.M. Grazing preferences in sheep and cattle: Implications for production, the environment and animal welfare. Can. J. Anim. Sci. 2010, 90, 285–293. [Google Scholar] [CrossRef]
Fermentation 09 00925 g001
Figure 1. Chromatogram of orchardgrass substrate incubated in the absence of rumen fluid at the start of the experiment (time 0). Chromatogram was obtained by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD). Neokestose, 1-kestose, and 1-nystose were not detected. A putative raffinose peak was so small that it could not be shown on the figure. Glucose (G), fructose (F), sucrose (S), 6-kestose (6K), bifurcose (Bf), and 6-nystose (6N) are indicated by arrows. Regions quantified as short-chain (SC) or long-chain (LC) fructan are indicated by brackets.
Figure 1. Chromatogram of orchardgrass substrate incubated in the absence of rumen fluid at the start of the experiment (time 0). Chromatogram was obtained by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD). Neokestose, 1-kestose, and 1-nystose were not detected. A putative raffinose peak was so small that it could not be shown on the figure. Glucose (G), fructose (F), sucrose (S), 6-kestose (6K), bifurcose (Bf), and 6-nystose (6N) are indicated by arrows. Regions quantified as short-chain (SC) or long-chain (LC) fructan are indicated by brackets.
Fermentation 09 00925 g001
Fermentation 09 00925 g002
Figure 2. Chromatograms of orchardgrass fructan profiles following incubation with rumen fluid from one steer (a) or ram lamb (b) for 0, 2, 4, and 8 h (h). Chromatograms were obtained by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD). Glucose (G) and fructose (F), indicated by arrows on the 8-h chromatograms, partially coeluted at earlier timepoints, when the concentrations were higher. Sucrose (S) was not detected in this particular 4-h incubation from a ram lamb (see panel (b)). Short- and long-chain (SC and LC, respectively) fructan regions are indicated by brackets.
Figure 2. Chromatograms of orchardgrass fructan profiles following incubation with rumen fluid from one steer (a) or ram lamb (b) for 0, 2, 4, and 8 h (h). Chromatograms were obtained by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD). Glucose (G) and fructose (F), indicated by arrows on the 8-h chromatograms, partially coeluted at earlier timepoints, when the concentrations were higher. Sucrose (S) was not detected in this particular 4-h incubation from a ram lamb (see panel (b)). Short- and long-chain (SC and LC, respectively) fructan regions are indicated by brackets.
Fermentation 09 00925 g002
Fermentation 09 00925 g003
Figure 3. Rate of (a) long-chain and (b) short-chain fructan appearance and disappearance during ex vivo bovine and ovine orchardgrass fermentations. A positive rate of change indicates that fructans were accumulating during a given time period; a negative rate of change reflects disappearance of fructans during a given period. Data are presented as LSM ± SE. There was a species by time interaction for rate of change of both long- and short-chain fructans (p < 0.01), with asterisks indicating species differences within timepoints (p ≤ 0.05).
Figure 3. Rate of (a) long-chain and (b) short-chain fructan appearance and disappearance during ex vivo bovine and ovine orchardgrass fermentations. A positive rate of change indicates that fructans were accumulating during a given time period; a negative rate of change reflects disappearance of fructans during a given period. Data are presented as LSM ± SE. There was a species by time interaction for rate of change of both long- and short-chain fructans (p < 0.01), with asterisks indicating species differences within timepoints (p ≤ 0.05).
Fermentation 09 00925 g003
Fermentation 09 00925 g004
Figure 4. Fermentation pH during ex vivo incubations of orchardgrass with bovine and ovine rumen fluid. Data are presented as LSM ± SE. There was a species by time interaction for pH (p < 0.01), with asterisks indicating species differences within timepoints (p ≤ 0.05).
Figure 4. Fermentation pH during ex vivo incubations of orchardgrass with bovine and ovine rumen fluid. Data are presented as LSM ± SE. There was a species by time interaction for pH (p < 0.01), with asterisks indicating species differences within timepoints (p ≤ 0.05).
Fermentation 09 00925 g004
Table 1. Chemical composition of orchardgrass substrate.
Table 1. Chemical composition of orchardgrass substrate.
NutrientConcentration, g kg DM−1
Crude Protein143.5
Acid Detergent Fiber353.9
Neutral Detergent Fiber578.4
Water-Soluble Carbohydrate91.5
Ethanol-Soluble Carbohydrate74.1
Table 2. Fructan and sugar concentrations in orchardgrass controls.
Table 2. Fructan and sugar concentrations in orchardgrass controls.
Fructan/SugarConcentration, μg mL−1
Long-chain †444.1
Short-chain †30.0
Bifurcose11.3
1-Nystose0.0
6-Nystose7.6
1-Kestose0.0
6-Kestose3.4
Neokestose0.0
Raffinose7.5
Free Sugars ‡1,064.8
† Long-chain fructans: DP > 8, retention time ≥ 20 min; Short-chain fructans: DP 4–8, retention time: ~11.8 (1-nystose) to 19.99 min. ‡ Free sugars: glucose + fructose + sucrose.
Table 3. Fructan and sugar concentrations in bovine and ovine orchardgrass fermentations (LSM; n = 4).
Table 3. Fructan and sugar concentrations in bovine and ovine orchardgrass fermentations (LSM; n = 4).
Time, hp-Values
Fructan/Sugar, μg mL−10248SESpeciesTimeSpecies × Time
BovineOvineBovineOvineBovineOvineBovineOvine
Long-chain †408.2 α421.7 α383.1 x,α349.0 y,β0.0 b,β50.2 a,γ0.0 β0.0 δ16.80.44<0.010.04
Short-chain †53.7 β85.8 β185.4 b,α255.9 a,α19.0 b,β249.4 a,α4.7 β11.2 γ21.5<0.01<0.01<0.01
Bifurcose12.2 β15.4 γ40.1 α48.9 β2.1 β85.8 α0.0 β4.8 γ9.6<0.01<0.01<0.01
1-Nystose1.7 β3.4 γ14.1 α17.9 β0.7 a,β30.3 b,α0.0 β0.0 γ3.4<0.01<0.01<0.01
6-Nystose7.98.111.310.88.58.40.71.51.40.91<0.010.96
1-Kestose0.3 b8.9 a4.10.40.02.20.00.02.20.400.150.06
6-Kestose6.07.09.78.67.910.80.00.71.80.51<0.010.74
Neokestose0.000.00.70.30.00.50.00.00.20.780.050.20
Raffinose5.54.45.85.20.00.90.00.0 1.00.77<0.010.70
Free Sugars ‡1104.91084.0908.6750.283.4230.643.149.665.00.85<0.010.11
† Long-chain fructans: DP > 8, retention time ≥ 20 min; Short-chain fructans: DP 4–8, retention time: ~11.8 (1-nystose) to 19.99 min. ‡ Free sugars: glucose + fructose + sucrose. a,b Indicates differences between species within timepoints (p ≤ 0.05). x,y Indicates trends for differences between species within timepoints (p ≤ 0.05). α,β,γ,δ Indicates differences within species between timepoints (p ≤ 0.05).
Table 4. Short-chain fatty acid (SCFA) concentrations in bovine and ovine orchardgrass fermentations (LSM; n = 4).
Table 4. Short-chain fatty acid (SCFA) concentrations in bovine and ovine orchardgrass fermentations (LSM; n = 4).
SCFA, mmol L−1SpeciesStatistics
BovineOvineSEp-Value
Lactate21.0317.231.460.12
Succinate1.821.190.750.58
Acetate17.70 a14.42 b0.59<0.01
Propionate8.258.000.730.82
Butyrate1.911.780.110.44
Total SCFA50.70 a42.62 b2.050.03
a,b Indicates significant differences between species (p ≤ 0.05).
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Weinert-Nelson, J.R.; Kagan, I.A.; Ely, D.G.; Flythe, M.D.; Davis, B.E. Fructan Catabolism by Rumen Microbiota of Cattle and Sheep. Fermentation 2023, 9, 925. https://doi.org/10.3390/fermentation9110925

AMA Style

Weinert-Nelson JR, Kagan IA, Ely DG, Flythe MD, Davis BE. Fructan Catabolism by Rumen Microbiota of Cattle and Sheep. Fermentation. 2023; 9(11):925. https://doi.org/10.3390/fermentation9110925

Chicago/Turabian Style

Weinert-Nelson, Jennifer R., Isabelle A. Kagan, Donald G. Ely, Michael D. Flythe, and Brittany E. Davis. 2023. "Fructan Catabolism by Rumen Microbiota of Cattle and Sheep" Fermentation 9, no. 11: 925. https://doi.org/10.3390/fermentation9110925

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop