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Article

Invasive Pigweed (Amaranthus spinosus) as a Potential Source of Plant Secondary Metabolites to Mitigate Enteric Methane Emissions in Beef Cattle †

by
Wilmer Cuervo
1,*,
Mariana Larrauri
2,3,4,
Camila Gomez-Lopez
4 and
Nicolas DiLorenzo
4
1
Animal and Veterinary Sciences Department, Clemson University, Clemson, SC 29634, USA
2
Facultad de Ciencias Agropecuarias, Universidad Nacional de Córdoba, Córdoba 5000, Argentina
3
CONICET-Argentina, Buenos Aires 1686, Argentina
4
North Florida Research and Education Center, Animal Sciences Department, University of Florida, Marianna, FL 32446, USA
*
Author to whom correspondence should be addressed.
†
This article is a revised and expanded version of an abstract entitled “Utilizing Invasive Pigweed (Amaranthus Spinosus) as a Novel Methane Mitigation Strategy in Beef Cattle Feed: A Sustainable Approach”, which was presented at Wilmer Cuervo et al., Louisville, KY, USA at the Southern Section from the ASAS 2023.
Grasses 2025, 4(2), 14; https://doi.org/10.3390/grasses4020014
Submission received: 14 January 2025 / Revised: 20 February 2025 / Accepted: 13 March 2025 / Published: 10 April 2025
(This article belongs to the Special Issue The Role of Forage in Sustainable Agriculture)
Versions Notes

Abstract

:
Global beef demand will rise by 40 million tons in 30 years, increasing methane (CH4) emissions. Pigweed (Amaranthus spinosus), an invasive weed abundant in southeastern U.S. pastures, may mitigate CH4. Yet, its potential as a feed additive remains unexplored. The aim of this study was to evaluate the influence of pigweed and its extracts on ruminal fermentation and CH4 production. For Exp 1, ruminal fluid from three American Aberdeen steers was incubated with 0, 2.5%, 5%, or 10% of diet dry matter (DM) of roots, stems, leaves, seeds, or the whole pigweed plant (WHO). In Exp 2, extracts from the leaves and WHO were incubated under the same conditions. For the first experiment, 2.5% of the roots, 5% of the leaves, and 10% of the WHO decreased acetate and butyrate concentrations (p < 0.01). In contrast, the WHO, leaves, and seeds at 2.5% of DM increased propionate concentration (p = 0.05). Increasing levels of WHO, leaves, and seeds quadratically reduced CH4 (p < 0.001). The addition of 2.5 and 5% of leaves and WHO reduced in vitro CH4 production per gr of organic matter fermented (p < 0.01). In Exp 2, based on their CH4 mitigation, the leaves and WHO were extracted, and their phenol (3.2 and 1.1 mg/g of DM, respectively) and flavonoid (19.7 and 1.9 mg/g of DM, respectively) contents were determined. Extracts from WHO (2.5%) decreased acetate and CH4 (p < 0.05), while 5% inclusion decreased gas production and increased ruminal pH (p < 0.03). Leaf extracts (2.5%) increased propionate and reduced acetate: propionate (p < 0.05). The leaves and WHO extracts did not affect IVOMD at either inclusion level (p > 0.4). Extracts at 5% from WHO were more effective than that from leaves in reducing CH4 (27% vs. 4%). The evidence suggests that the inclusion of 2.5 to 5% of WHO extracts shifts ruminal fermentation towards propionate-producing impairing methanogenesis, representing a sustainable strategy to mitigate CH4. This hypothesis must be further assessed under in vivo supplementation of the extracts to beef cattle.

1. Introduction

Projections for global population growth indicate that an unprecedented rise in food production will be needed during the next few decades [1]. The beef industry worldwide, particularly within the U. S., will determine the consumption of protein [2]. Estimates as high as an additional 40 million tons of beef are projected by 2050 [3]. A great portion of the increased demand in beef products will be fulfilled with animals raised in U.S. cow–calf operations, which are based largely on pasture systems [4]. Although effective in exploiting the forage resource, grazing animals exhibit a greater enteric methane (CH4) emission than during any other productive stage of the beef life cycle [5,6,7]. Methane is a very potent greenhouse gas; its global warming potential is 28 times that of carbon dioxide (CO2) over a 100-year time frame [8]. Given that enteric CH4 emissions from cattle represent one of the largest sources of greenhouse gases in agriculture, the projected increase in the beef industry places additional pressure on finding viable, effective, and sustainable CH4 mitigation strategies [9]. According to a recent study [10], one of the most efficient strategies to mitigate CH4 emissions in grazing systems is the inclusion of tanniferous forages. Tannins are a group of phenolic compounds for which inclusion in the diet of beef cattle has been linked to a reduction in CH4 [11]. Similar to phenols, flavonoids, alkaloids, and saponins are metabolic compounds produced in the various pathways of secondary metabolites and, for that reason, are defined as plant secondary metabolites (PSMs) [12]. While Saponins may disrupt sterols in protozoal membrane and affect membrane transport proteins (i.e., calcium and sodium channels) [13,14], tannins may bind adhesin proteins from a cell envelope in Archaea, impeding methanogen-protozoa complex formation and decreasing hydrogen transfer [15]. On the other hand, flavonoids negatively impact Gram-positive bacteria, inhibiting the synthesis of their cell wall and nucleic acids [16]. Interestingly, these secondary compounds are widely present in several invasive weeds [17], some of which are constantly present within the pastures grazed by beef cattle in the U.S. Of these species, pigweed (Amaranthus spinosus) is one of the most troublesome and aggressive that competitively expands across grasslands, reducing biomass total forage value and quantity [18]. Interestingly, pigweed is exceptionally rich in PSM, including polyphenols and flavonoids [19], which have received attention in ruminant research for their potential role in mitigating methanogenesis in the rumen.
Pigweed is one of the most invasive weeds in the pastures grazed by beef cattle in cow–calf operations located across the southeast U.S. [20]. However, the presence of pigweed might represent a unique opportunity for leveraging this invasive species as part of a CH4 mitigation strategy. Even more, as an invasive species, using pigweed as a feed supplement could offer a dual advantage in terms of an alternative control strategy and potentially diminishing the environmental footprint associated with beef production. Nevertheless, to the best of our knowledge, no studies have addressed the effects of controlled intake of pigweed invading grazing paddocks, and none have studied extracts of PSMs from spiny pigweed to explore their effects on CH4 production from beef cattle. There is a gap in the knowledge in identifying the specific effects of PSMs present in aggressive weeds like pigweed on ruminal function and particularly the potential of pigweed extracts as a feed additive for ruminants. Therefore, the objective of this study was to investigate the effect of the in vitro addition of morphological portions of pigweed on ruminal fermentation parameters, in vitro organic matter digestibility (IVOMD), and CH4 production. Furthermore, we tested the effects of including extracts from selected portions on the same variables to establish the specific method of action of PSMs present in pigweed when added to ruminal conditions from a beef steer under grazing conditions. We hypothesize that the addition of pigweed will promote ruminal fermentation changes leading to a reduction in CH4 production. Likewise, we hypothesized that the potential effects on ruminal parameters would vary according to the evaluated morphological portion and the level of inclusion of the extracts.

2. Materials and Methods

2.1. Location and Experimental Animals

Regarding the use of experimental animals, the Institutional Animal Care and Use Committee of the University of Florida (protocol number 00005681) reviewed and approved of all procedures for the experiments conducted at the North Florida Research and Education Center (NFREC), Marianna, FL. This study was divided into two experiments, in which sampling and collection were performed at the Feed Efficiency facilities of the NFREC using 3 ruminally cannulated Angus steers (875 ± 24 kg of body weight; BW; and 10 years old). A total of 12 plants of Amaranthus spinosus (28–35 d of regrowth) were harvested by digging around each plant to obtain the whole morphological portions, from the roots to the seeds (Table 1). The plants were harvested in late spring to be processed and incubated in a total of 6 in vitro incubations, performed from October 2023 to July 2024.

2.2. General Description of Experiments

2.2.1. Experiment 1—In Vitro Batch Culture of Morphological Parts of Pigweed

In Exp 1, three separate in vitro incubations were performed to evaluate the effect of the individual addition of four morphological portions from Pigweed (roots, stems, leaves, seeds) and the whole pigweed plant (WHO). The separated portions, WHO, and samples from the forage consumed by the cannulated steers—bermudagrass (Cynodon spp.)—were dried (55 °C—48 h), grounded through a 2 mm sieve in a Wiley-type Mill (Arthur H. Thomas Co., Philadelphia, PA, USA), and stored for further incubation (see Section 2.3). Each morphological portion from the pigweed plant was incubated at four levels of inclusion: 0, 5, 10, and 20% of dry matter incubate (DMI). Several ruminal fermentation parameters were determined, as were IVOMD and CH4 production. Cannulated steers (3 per incubation) were grazing bermudagrass for 7 d prior to the collection and served as donors of ruminal fluid (RF), which was deposited directly into 125 mL glass serum bottles. Thus, 0, 5, 10, and 20% of the incubated DM from the diet (Bermudagrass) were replaced by the correspondent amount of each morphological portion. Glass bottles were incubated in triplicate per animal and per treatment under independent batch culture incubations. Each morphological portion at each evaluated dosage was incubated in individual serum bottles. After 24 h of incubation, ruminal pH and gas production were measured, with samples collected for later determination of ammonia nitrogen (NH3-N), CH4 concentration, and volatile fatty acids (VFAs). Similarly, using plastic scintillation tubes, IVOMD was determined following the two-stage technique described by [21]. All procedures will be further described in detail in Section 2.3 and Section 2.4.

2.2.2. Experiment 2—Batch Culture of Extracts from Pigweed

In the second experiment, the same cannulated steers were included as donors; RF was extracted, diluted, and incubated in 125 mL glass bottles containing basal diet (Table 1) with extracts from specific parts of the pigweed. Based on the results from Experiment 1, extracts from the leaves and WHO were isolated and purified (as described below on Section 2.4. See Preparation of pigweed ethanolic extracts and yield) and added at 2.5 and 5% of DMI. After incubation, the ruminal parameters were determined at 24 h and 48 h (as explained before), including ruminal pH, VFA, NH3-N, IVOMD, and CH4. In addition, the extraction yield from each morphological component and antioxidant activity from extracts were determined. The step by steps of pigweed extraction and antioxidant activity methodologies are described in Section 2.4.

2.3. Ruminal Collection and Batch Culture Incubations

2.3.1. Collection of Ruminal Fluid and Procession for Batch Culture

Ruminal fluid from ruminally cannulated steers were obtained at 07:00 am, collecting from the medium ventral sac of the rumen as a representative sample of ruminal content. Fluid was strained through 4 layers of cheesecloth until RF overflowed a 750 mL pre-warmed (39 °C) thermos to guarantee anaerobic conditions until further processing on the sample in the in vitro batch culture incubations. Later, in the laboratory, under constant CO2 flushing, RF from the experimental animals was mixed and filtered (2 layers of cheesecloth), and pH was recorded using a manual pH meter (Five easy plus FP20, Mettler Toledo, Columbia, MD, USA). Then, the RF was diluted (3:1 v:v) with pre-warmed McDougall’s buffer, and 50 mL of the mixed sample was placed in a 125 mL glass bottle without diet (blank), containing 0.7 g of the dried and ground diet (bermudagrass), stated as the control (CON), or the diet plus the treatment. For Exp 1, three bottles per combination of morphological part (root, stem, leaf, seed, and WHO) and inclusion level (0, 2.5%, 5%, and 10% of DMI) were incubated at 39 °C under agitation (60 rpm) for 24 h using a SSI3 SHEL –LAB/shaking incubator (SHELDON Manufacturer Cornelius, OR, USA). Ruminal pH, VFA, NH3-N, CH4, and gas production were determined after 24 h.

2.3.2. Gas Production and Methane Concentration

After 24 h of incubation, fermentation in 125 mL glass bottles was stopped by placing them for 10 min on an ice bath, gas pressure was determined by using a digital pressure gauge (No 23V730 Grainger International Inc., Lake forest, IL, 60045, USA), and a gas sample was collected from the headspace of the bottle using a 14-gauge needle connected to a 2-way valve on a syringe plugged into a vacuumed and sealed 5 mL sterile glass vial (Med lab Pompano Beach, FL, USA). Gas bottles were stored at room temperature for further CH4 determination. The methane concentration in the gas samples was analyzed by gas chromatography (Agilent 7820A GC; Agilent Technologies, Palo Alto, CA, USA) using flame ionization and a capillary column (Plot Fused Silica 25 m by 0.32 mm, Coating Molsieve 5A, Varian CP7536; Varian Inc., Lake Forest, CA, USA). To ensure proper determination, certified standards were used to regulate the gas chromatograph for CH4 (4 mg/L, Scott Marrin Inc., Riverside, CA, USA). The temperatures for the injector, column, and detector were 80 °C, 160 °C, and 200 °C, respectively. Nitrogen (N2) was used as the carrier gas with a flow rate of 3 mL/min. The detector makeup flow was 21.1 mL/min, and the split ratio for the injected CH4 sample was 5:1.

2.4. Laboratory Analysis

2.4.1. Volatile Fatty Acid Concentration

After gas sample collection, the 125 mL glass bottles were unsealed, and 50 mL of the diluted RF was acidified (1:100 of 20% v:v H2SO4 solution) and stored at −20 °C for VFA determination in liquid–liquid solvent extraction using ethyl acetate [22]. The thawed samples were centrifuged (10,000× g, 15 min, 4 °C), and 2 mL of the supernatant was mixed with 4 mL of the internal standard of metaphosphoric (25% w/v): crotonic (2 g/L) acid solution. After overnight frizzing, the samples were thawed and centrifuged (10,000× g, 15 min, 4 °C), and the supernatant was transferred into 12 × 75 mm borosilicate glass culture tubes (Fisherbrand; Thermo Fisher Scientific Inc., Waltham, MA, USA) and mixed (1:2) with ethyl acetate. After vortexing (2 s), the tubes were rested (5 min) to allow the separation of the ethyl acetate; then, the top layer fraction was pipetted out into clear glass vials (9 mm; Fisherbrand; Thermo Fisher Scientific Inc., Waltham, MA, USA). The samples of VFA were analyzed using a gas chromatograph (Agilent 7820A GC, Agilent Technologies) with a flame ionization detector and a capillary column (CP-WAX 58 FFAP 25 m × 0.53 mm, Varian CP7767; Varian Inc.). The column temperature was maintained at 110 °C, and the injector and detector temperatures were 200 and 220 °C, respectively.

2.4.2. Ammonia (NH3-N) Concentration

Ammonia–N was determined by using the phenol-hypochlorite reaction, according to [23]. Briefly, thawed RF samples were centrifuged (10,000× g, 15 min, 4 °C), and 20 μL of supernatant was pipetted into 12 × 75 mm borosilicate glass culture tubes (Fisherbrand; Thermo Fisher Scientific Inc., Waltham, MA, USA) containing 1 mL of Phenol reagent. The tubes were vortexed, and 0.8 mL of a hypochlorite solution was added, vortexed again, covered with glass marbles, and placed in a water bath at 95 °C for 5 min. The only adaptation from the original method was that the final absorbance reading was carried out on 200 μL at 665 nm in a 96-well plate reader (DU 500: Beckman Coulter Inc., Palo Alto, CA, USA).

2.4.3. In Vitro Organic Matter Digestibility

At the same time as the batch culture, 0.7 g of the substrate (diet plus all the treatments) and inoculum (diluted RF) was incubated in 200 mL plastic scintillation tubes with rubber stoppers to analyze IVOMD. At 24 h, 6 mL of 20% (v:v) HCl solution was added to each tube, and after a slight swirl, 2 mL of a 5% pepsin suspension was added; then, the tubes were returned to the incubator for another 24 h. After this period, the samples were filtered by a vacuum (Whatman No. 541 ashless; Whatman International Ltd., Maidstone, UK), rinsed with deionized water, dried (65 °C for 48 h), and then burned (550 °C, 6 h) to determine the IVOMD based on gravimetric calculations, as follows.
IVOMD (%) = [(incubated organic matter − residual organic matter)/incubated organic matter] × 100.

2.4.4. Preparation of Pigweed Ethanolic Extracts and Yield

The pigweed was collected from NFREC, Marianna, FL, USA. The pigweed was dried in an oven at 65 °C for 48 h. The morphological parts of the pigweed were separated in order to obtain the leaves and WHO and were grounded (2 mm sieve) to a powder using a grinder. The polyphenols from the leaves and WHO were extracted by a solid-liquid method using ethanol-water (70:30 v:v). Ten grams of each fraction (Leaves or WHO) was mixed with 250 mL of extraction solvent and placed in a shaking water bath (Avanti J-E, Beckman Coulter Inc., Palo Alto, CA, USA) at 60 °C, 80 rpm for 40 min. After that, the extracts were filtrated (Whatman 3) and dried in a rotary evaporator at 55 °C (RE-501, VEVOR, USA). The aqueous part of the leaves and WHO extracts were lyophilized to dryness. The extracts were kept in caramel flask at −18 °C until use [24].
The extraction yield of pigweed extracts was calculated according to the following formula [25]:
Extraction yield = (g dry extracted matter)/(g dry pigweed morphological part).

2.4.5. Total Phenol and Flavonoid Content

The total phenolic compounds in the pigweed leaves and WHO extracts were determined using the Folin–Ciocalteu method [25]. The absorbance of samples was measured in a spectrophotometer at 760 nm (Fisherbrand UV/VIS AccuSkan GO Spectro-photometer, Thermo Fisher Scientific Inc., Hampton, NH, USA). Gallic acid (GAE, Thermo Fisher Scientific, Bridgewater, NJ, USA) was used as standard. Total phenolic content was expressed as mg of gallic acid equivalent (GAE)/g dry matter (DM) extract.
Total flavonoids were determined according to the AlCl3 method [26] measuring absorbance at 367 nm. The calibration curve was prepared using quercetin (QE, Thermo Fisher Scientific, Bridgewater, NJ, USA). The results were expressed in mg quercetin equivalent (QE)/g DM extract.

2.4.6. DPPH Radical Scavenging Activity

The radical scavenging activity of the pigweed leaves and WHO extracts was deter-mined using diphenyl picryl hydrazyl radical (DPPH) according to [25]. Different aliquots of the samples on methanol (100 μg/mL) were added to a 1.5 mL DPPH methanolic solution (20 μg/mL). The absorbance of samples was measured at 517 nm. The inhibitory concentration 50% (IC50) was calculated from the curve obtained by plotting inhibition percentage of vs. final extract concentrations [27].

2.5. Calculations and Statistical Analysis

Data from Exp 1 were analyzed as a randomized complete block design following a 5 × 4 factorial arrangement of treatments (5 portions × 4 inclusion levels), using the MIXED procedure of SAS version 9.4 (SAS Institute Inc., Cary, NC, USA). Each of the independent incubations was considered as a replicate. For Exp 1, the model included the fixed effects of the morphological portion (root, stem, leaf, seed, and WHO), the inclusion level (0, 2.5%, 5%, and 10%), and their interaction when significant. Polynomial orthogonal contrasts were calculated to determine the linear, quadratic, or cubic effects of the increasing addition of pigweed morphological portions. A value from the 0% inclusion level (control) was considered for building and analyzing the contrasts. Significant differences were accepted if p < 0.05, and tendencies were considered when 0.05 < p < 0.1.
For Exp 2, the data were analyzed as a randomized complete block design following a 2 × 3 factorial arrangement of treatments (2 extract sources × 3 inclusion levels), using the PROC MIXED of SAS version 9.4. The model included the fixed effects of the extracts (from the leaves and WHO), inclusion level (0, 2.5%, 5%), and their interaction when significant. Polynomial orthogonal contrasts were calculated to determine the linear or quadratic effects of the addition of the pigweed extracts. For both experiments, incubation was included as a random effect and the experimental unit corresponded to the average of the three bottles within each incubation. The means from the three incubations were compared using Tukey’s test.

3. Results

3.1. Ruminal Fermentation After the Addition of Morphological Parts from Pigweed

The proportion of stem (59.6 ± 6.2% of DM), leaves (19.9 ± 7% of DM), seeds (13.5 ± 1.9% of DM), and roots (2.2 ± 1% of DM) in the WHO is included in Table 1. Regarding the chemical composition, crude protein (CP) of leaves was elevated (29.4% of DM) compared to WHO (20.2% of DM) and particularly compared to the bermudagrass basal diet (9.1% of DM). In contrast, the estimated organic matter (OM) content for WHO was reduced (79.3% of DM) compared to the leaves (84.8% of DM) and basal diet (83% of DM). The content of neutral and acid detergent fiber (NDF and ADF) for WHO (48.1% and 28.8%, respectively) and the leaves of pigweed (41.2% and 16.7%, respectively) were notably lower when compared to the incubated diet (72.7% and 36.2%, respectively).
As shown in Table 2 and Table 3, acetate was affected by the incubated portion (p = 0.05), and the acetate–propionate ratio (A:P) was influenced by the inclusion level (p < 0.001). Except for A:P, significant portion × inclusion interactions were observed across all fermentation parameters (p < 0.01). The inclusion of 2.5% of incubated DM of all portions, except roots, increased the total VFA (p < 0.01), branched chain VFA (BCVFA), propionate, and butyrate concentrations (p < 0.001). Increasing levels of all portions, except roots, resulted in a marked decrease in the concentration of these fatty acids, particularly when WHO was incubated (p < 0.001). The incubation of stems led to a greater acetate concentration (p < 0.001), whereas WHO reduced its concentration (p < 0.001). A linear reduction in acetate molar proportion (p < 0.001) was observed after increasing the inclusion of all portions (Table 2), with the reduction being more pronounced at higher levels (5 and 10%, Table 2) of leaves (p < 0.001). As shown in Table 3, propionate molar proportion exhibited a quadratic response, with an increase at 2.5% inclusion followed by a linear decrease at greater inclusion levels. No differences in propionate molar proportion were detected among morphological portions (p = 0.29). As shown in Table 2, a significant interaction (p < 0.001) revealed a linear reduction in butyrate proportion with increasing inclusions of WHO, stems, and roots (p < 0.001). By contrast, a quadratic increase in butyrate molar proportion was observed with increasing the seed inclusion levels, with the effect being more pronounced at 2.5% and 5% inclusion of leaves (Table 3). Similar trends were noted for BCVFA molar proportion to those observed for butyrate. While the inclusion of leaves promoted a greater BCVFA molar proportion, WHO inclusion had the opposite effect (p < 0.001).
As shown in Table 4, while the inclusion of leaf and roots depressed NH3-N concentration (p < 0.001), the stems enhanced it when incubated in ruminal fluid (p < 0.01). A quadratic reduction in NH3-N was observed at increasing levels of the evaluated morphological portions of the pigweed plant. In contrast, a quadratic increase in ruminal pH was observed with increasing inclusion levels, with a pronounced increment at 2.5% (p < 0.001). Higher levels of WHO (5%, and 10%) resulted in elevated ruminal pH (p < 0.01). Such an increment was not observed for the rest of the portions (p < 0.001). The gas production tended to be reduced by the addition of seeds and WHO (p = 0.09), while the addition of stems increased gas production (p = 0.02). Likewise, IVOMD was linearly reduced after increasing levels of pigweed (p < 0.001), particularly when the seeds, leaves, and WHO were incubated (p < 0.001). A portion × inclusion interaction (p < 0.01) revealed an identical trend for CH4 intensity (mmol/g of fermented OM) at elevated inclusion of the leaves and WHO (p < 0.01). Despite CH4 concentration (mg/mL) not being affected by the main factors (p > 0.15) nor their interaction (p = 0.11), CH4 produced by every g of incubated diet exhibited an identical behavior observed for CH4 intensity.

3.2. Solid–Liquid Extraction with Solvents

As shown in Table 5, the extraction yield for the selected portions was notably greater for the seeds (155.23 mg/g) than for the WHO (131.19 mg/g), leaves (118.22 mg/g), and roots (135.58 mg/g). The seeds and leaves showed higher phenol contents (3.5 and 3.15 mg/g, respectively) compared to the rest of the parts (<1.2 mg/g). Similarly, a greater flavonoid content was observed for the seeds (3.7 mg/g) and leaves (19.7 mg/g) than for the rest of the plant components (<2 mg/g). The antioxidant activity, expressed as the concentration of the antioxidant molecule (µg/mL) required to reduce the initial concentration of DPPH radicals by 50% (DDPH EC50 µg/mL), was statistically greater for the seeds (133,12 µg/mL) than for the leaf (169.8 µg/mL), WHO (1262.5 µg/mL), stem (1639 µg/mL), and root (4164 µg/mL) extracts (p < 0.05)

3.3. Ruminal Fermentation After the Addition of Extracts from Pigweed

The chemical composition of the incubated forage and the selected portions to be extracted are shown in Table 1. No portion × inclusion interaction was detected for any of the evaluated fermentation parameters (p > 0.05). Table 6 shows how the addition of extracts from the WHO resulted in a greater ruminal pH (6.57 vs. 6.53) and NH3-N concentration (8.03 vs. 8.7 mM) than extracts from the leaves (p < 0.03). In contrast, extracts from the leaves led to a greater concentration of propionate (p = 0.04). The A:P was particularly reduced when 5% of pigweed extracts (either WHO or leaves) were incubated (p = 0.02). Gas production (mL/g OM fermented) was significatively lower when WHO extracts were added (p = 0.01), particularly under 5% inclusion (47.07 mL/g OMF) compared to 2.5% (56.49 mL/g OMF; p = 0.01). Methane concentration (mM) and CH4 intensity (mM/g of OM fermented) were diminished under WHO extract inclusion (p < 0.03) and this reduction seemed to be more pronounced when pigweed extracts were included at 2.5% compared to 5% of incubated DM. Thus, the CH4 concentration reduction was greater (27%) under the inclusion of 5% of WHO extracts compared to leaf extracts at the same rate (4%), as shown in Table 6. Neither of the analyzed extracts at the evaluated doses affected IVOMD (p > 0.4).

4. Discussion

4.1. Chemical Composition Effects on Ruminal Fermentation

The observed results from both experiments in the study suggest that pigweed (Amaranthus spinosus) as a direct-fed additive or its extracts have significant potential to mitigate enteric CH4 emissions in beef cattle. To the best of our knowledge, the intentional inclusion of pigweed biomass material or its extracts to a beef cattle diet have never been evaluated under in vivo nor in vitro conditions. Recent reports from grazing animals consuming pigweed include toxicity issues such as myocardial degeneration, renal disease and nitrate intoxication [28], renal edema and hemorrhage, dramatic increased in aminotransferases, diarrhea, and eventually the death of the animal [29]. Regarding the chemical composition, ADF was considerably lower in the WHO (34%) and leaves (43%) compared to bermudagrass added to the culture bottle, which could partially contribute to a lower CH4 in vitro production, which has been previously observed when ADF was reduced in the incubated diet [30]. The discrepancy between the reconstructed and measured WHO CP values could be attributed to differences in sample selection. The sample for the WHO to determine N was likely obtained from a single pigweed plant rather than a mix representing the average morphological composition (19.8% leaves, 13.5% roots, 2.2% seeds, and 59.6% stems). In contrast, the CP values for individual portions were determined from pooled samples. If the selected plant had a higher proportion of leaves and seeds and fewer stems and roots, this could have resulted in an overestimated whole-plant CP value.
The higher extraction yields from the seeds and WHO confirm the potential of these options to be scaled to an alternative additive based on pigweed. Interestingly, extracts from the seeds and leaves exhibited the greatest contents of phenols and flavonoids, and antioxidant activity of all the evaluated extracts, coinciding with previous observations by [31] reporting an increase in arsenic detoxification of urine and feces from dairy cattle. Those authors attributed this detoxification to an enhancement of the enzymatic and non-enzymatic antioxidant metabolic processes mediated by Amaranthus spinosus preventing cellular oxidative damage. Likewise, previous reports [32] agreed with this mechanism after using pigweed leaf extracts, showing similar proportions between phenol (2.81 in GAE, g/100 g) and flavonoid (18.4 QE, g/100 g) contents compared to the present study (3.19 mg of GAE and 19.67 mg of QE/g of DM, respectively). Moreover, that author reported higher antioxidant activity (80 µg/mL) than that observed here (164 µg/mL). Such activity coincided with a potent antibiotic effect against Streptococcus and Escherichia coli, comparable to the action of Ampicillin. This could partially explain the shift in ruminal fermentation patterns mediated by the addition of seed and leaf extracts.

4.2. Effects on Ruminal Fermentation

In line with the reduced fiber detected in the samples after increasing the inclusion (2.5% to 10% of DMI) level of pigweed portions, a linear reduction in A:P was observed. This observation agreed with that of [33]: a reduction in A:P after incubating decreasing levels of fiber in the diet. The elevated BCVFA, propionate, butyrate, and total VFA production observed after the inclusion of 2.5% of all pigweed portions (except roots) might suggest that greater inclusions could impair microbial metabolism, leading to changes in the major VFA synthetic pathways [34]. This agrees with previous reports of elevated polyphenol and flavonoid concentrations in extracts from tea and dragon fruit peel, which were associated with the reduction in total VFA [35,36]. Indeed, when the inclusion was greater (10%), the addition of WHO decreased VFA, confirming the suggested effect. Interestingly, the specific effects on particular VFAs varied with the incubated portion. Thus, stems increased acetate concentration, which could be associated with the elevated NDF concentration and reduced CP in these portions, agreeing with the observations of [34], which reported an increase in acetate concentration (18%) after incubating diets with increasing levels of NDF and ADF from forages. Interestingly, the incubation of the diet with WHO resulted in a reduction in acetate concentration, suggesting that a particular combination of portions (excluding stems and roots) mediated a change in ruminal populations, leading to a reduction in this VFA. Considering that stems and roots represent nearly 73%, one can hypothesize that PSM activity in seeds and leaves is notably potent in mediating the observed shifts in VFA. Previously, ref. [37] reported reduced fiber (< 18%) and elevated soluble carbohydrate content (27%) in the aerial parts of pigweeds. These traits has been associated with a reduction in acetate concentration in ruminal fluid [38,39]. Here, the observed linear reduction in acetate molar proportion after increasing levels of leaves confirms the suggested effect. Considering that changes in acetate molar proportion were observed even at low doses of inclusion (2.5 and 5%), it could be proposed that besides the greater soluble carbohydrate concentration, PSM (particularly phenols and flavonoids) in the leaves mediated the changes in acetate. Such shifts in acetate might suggest a modulation of ruminal microbiome by PSMs in the leaves.
Despite no differences in propionate being detected among the incubated portions, the quadratic increase in this VFA suggested how propionate producer microbes seemed to be stimulated with low doses (2.5%) but affected by greater inclusions (10%). This agrees with the findings in [35], in which the authors associated the observed increase in propionate concentration after polyphenol (from tea) addition (4% of DMI) to an enhanced expression of genes associated with Ruminococcaceae, a family linked to propionate production. A cubic trend showed how increasing levels of WHO, stems, and roots reduced butyrate. This unclear behavior after increasing levels of polyphenols was observed by [15] after adding condensed tannins from Lotus pedunculatus at 10 mg/100 g of DM inoculated. The greater BCVFA concentration after the inclusion of pigweed leaves coincided with the results from previous studies evaluating that dragon fruit peel extracts elevated the content of polyphenols (particularly flavonoids) added to RF from deer under in vitro conditions. Such an increase coincided with an increased population of cellulolytic bacteria and greater NDF digestibility [36]. Coinciding with the observed here, previous studies [40] reported that the addition of 50 mg/g of synthetic glucoside and tannic acid (flavonoids similar to those present in pigweed leaves) resulted in a notable reduction in NH3-N. These and other authors [41] associated this response with the inhibitory effects of phenols on the adherence and proteolytic activity of ruminal microbes to protein. Besides the effects on VFA, ruminal pH exhibited a quadratic increase, particularly after 5% and 10% after the addition of WHO. These effects are in agreement with previous observations revealing that polyphenol addition significatively reduces the relative abundance of Streptococcus, as a relevant population acidifying ruminal environment. It has been hypothesized that polyphenols affect the cell membrane fluidity of these microbes [42].
The extracts from WHO mediated a reduction in ruminal pH and an increase in ammonia concentration, suggesting additional effects on populations linked with reducing ruminal pH. This result agreed with the reports by [43] about the effect of polyphenols (condensed tannins) on reducing the activity and abundance of Gram-positive bacteria (i.e., Actinobacteria), which have been associated with ruminal pH drops [44]. Considering the previously reported antibiotic effect of extracts from pigweed leaves on microbes (i.e., Streptococcus, E. Coli) [32] and since changes on ruminal fermentation were specific for acetate, propionate, A:P, and BCVFA, one could suggest that the antibiotic mechanism mediated by phenols and flavonoids affects microbial populations differentially, resulting in different changes in metabolic products (i.e., VFA and NH3).

4.3. Methane Reduction Potential

Increasing levels of seeds, leaves, and WHO reduced gas production and CH4 intensity (mM/g of OMF) but tended to decrease at higher doses (10%). This response agreed with that in [45], the authors of which reported dose-dependent negative effects of PSMs on microbial populations. Therefore, maintaining inclusion levels below 5% appears critical to balancing CH4 mitigation with fermentation efficiency. Simultaneously, at higher inclusion levels (10%), reductions in VFA production, CH4 intensity, and IVOMD might suggest a potential inhibition of overall microbial activity. This theory has been previously proposed by [46], who reported the inhibition of Archaea species relative abundance after the addition of 20 g/kg of pure flavonoids, specifically the mix of naringin and hesperidin. Similar to the present study, these authors did not report a negative impact on IVOMD when low doses of the evaluated extracts were incubated. The quadratic reduction in CH4 intensity with increasing inclusion of leaves and WHO aligns with that in studies showing the effectiveness of polyphenols in suppressing methanogenesis [47]. Similarly, the extracts from WHO at 2.5% were particularly effective, promoting CH4 reductions of over 10% without affecting IVOMD. These results suggest that pigweed extracts are as effective as other well-documented additives like condensed tannins [11].
When analyzing the high inclusion level of extracts in the incubation (5% of incubated DM), CH4 reduction was greater with the WHO than leaf extracts, suggesting that other morphological portions like seeds might contain PSMs with greater anti-methanogenic activity or that such CH4 mitigation is mediated by different metabolites. Considering this was an exploratory study, only two portions showing potential to mitigate CH4 production, enhancing propionate and not affecting IVOMD, were selected to obtain the extracts (WHO and leaves). Remarkably, neither doses (2.5 and 5%) nor extracts (leaves or WHO) affected IVOMD, suggesting that the effect mediated by phenols and flavonoids present in the extracts were particularly directed against microbial communities producing acetate and CH4, but not to all microbes associated with fermentation processes. A direct effect of polyphenols (condensed tannins) on ruminal Archaea had been previously reported [15]. Considering that the reduction in CH4 did not concur with a reduction in IVOMD after the addition of WHO and leaf extracts, we could suggest that phenols and flavonoids from pigweed extracts might also exert a direct and specific effect on methanogens. Nonetheless, this effect should be further evaluated using specific primers and ideally meta-transcriptomic analyses. In this sense, previous studies [40] showed that the in vitro incubation of 50 mg/g of flavonoid (Gallocatechin) and phenolic (tannic acid) compounds resulted in a simultaneous reduction in ruminal protozoa and CH4 production. Thus, future studies involving the use of these extracts must assess the count, diversity, and viability of protozoa.
The results from incubating bermudagrass with the extracts suggested that an additional portion other than the leaves (most likely the seeds) might be mediating the observed CH4 reduction. This agrees with previous observations by [48], who reported that the seeds and leaves from Amaranthus are used in Ethiopia to treat amoeba infection in humans due to their chemical components acting like toxins against intestinal microorganisms. These observations are consistent with the capacity of polyphenols to modulate microbial populations, shifting VFA dynamics towards propionate-producing bacteria over methanogens [49]. Likewise, a recent study showed that such a shift in populations decreased hydrogen availability for methanogenesis, leading to lower CH4 synthesis and production [50]. The addition of low levels (2.5% and 5% of incubated DM) of leaves, seeds, and WHO were particularly effective in reducing CH4 production, suggesting that their polyphenolic and flavonoid contents (as determined in Exp 2) are linked to these effects. Similar findings have been reported for tannin-rich feeds like quebracho (Schinopsis balansae) [51].
Overall, the inclusion of pigweed leaves, seeds, and the whole plant, as well as the extracts from leaves and the whole plant influenced ruminal fermentation, VFA profiles, and CH4 production without compromising IVOMD. These findings align with previous studies on polyphenol-rich plants such as sainfoin and quebracho, which have been shown to decrease CH4 emissions via shifting fermentation pathways toward propionate or decreasing acetate to propionate [52,53]. When comparing the effects mediated by pigweed morphological portions and their extracts, the observed results evidenced an important trade-off. While extracts provide a concentrated source of active compounds, their preparation is labor-intensive and costly. Considering that the low inclusion of WHO mitigated CH4 while promoting ruminal fermentation towards propionate, it is valuable to explore processing technologies (e.g., silage, baleage, drying and grinding, or pelleting) to scale up the potential use of pigweed biomass in grazing systems. Likewise, since direct grazing represents a barrier to the use of pigweed and animals will avoid pigweed-infested areas [54], obtaining extracts from this invasive weed might represent a sustainable approach for grazing systems. However, further research is needed to optimize extraction techniques and assess the feasibility of using pigweed as a commercial feed additive. Besides the observed ruminal fermentation responses, the dual role of pigweed as an aggressive invasive plant and as a CH4 mitigator might represent a unique opportunity to enhance pasture sustainability. To date, only two controlled studies have evaluated the effects of pigweed invasion in grazing paddocks on beef steers’ performance [18,55]. These studies found no significant impact on performance but highlighted the negative effects of spines in mature plants, which hinder the grazing process. By exploring potential management strategies to allow pigweed within pastures in a controlled fashion, producers could reduce the environmental footprint of beef production while managing pastureland more effectively. Similar strategies have been proposed for other invasive species with high PSM content [55].
The evident effects of extracts from WHO on A:P and CH4 mitigation that were not observed in leaves’ extracts open the door for further examination of specific extracts (including tannins) from the seeds and potentially from the roots. Nevertheless, when thinking about scaling up these types of alternative supplements, extracting polyphenols and flavonoids from WHO appears to be a more realistic opportunity. While this study provides compelling evidence for the efficacy of pigweed in reducing CH4 emissions, several limitations should be addressed. First, the incubation with extracts from pigweed was conducted in vitro, and the observed results should be further evaluated under in vivo conditions. Second, the specific phenols and flavonoids responsible for the observed effects were not identified. Thus, future research should focus on in vivo trials and compound identification using more specific techniques (i.e., ultra-performance liquid chromatography) to optimize the application of pigweed and its extracts as a feed additive.

5. Conclusions

This study demonstrates the potential of pigweed (Amaranthus spinosus) as an alternative additive to mitigate enteric CH4 emissions in beef cattle while modulating rumen fermentation. The present study demonstrated that the addition of pigweed leaves, seeds, and whole plant at 5% of incubated DM promoted ruminal fermentation changes, leading to a reduction in CH4 production. Similarly, the observed effects on acetate, propionate, and CH4 production varied according to the evaluated extracts. Among the morphological portions evaluated, leaves and the whole plant inclusion exhibited the greatest reductions in CH4 yield, particularly at 5% of DM inclusion. Simultaneous to such mitigation, shifts in VFA favoring propionate production and diminishing A:P were observed. The observed changes coincided with the detection of elevated concentrations of phenols and flavonoids, and a notable increase in antioxidant activity in the seeds, leaves, and whole plant of pigweed. The use of pigweed as a feed additive not only offers a CH4 mitigation strategy but also provides an innovative way to utilize an invasive weed prevalent in grazing paddocks across the southeast of US, addressing both environmental and agricultural challenges. Further research is needed to validate these findings under in vivo experimental settings and to explore the long-term implications of pigweed extract supplementation on animal performance and environmental sustainability.

Author Contributions

Conceptualization, W.C., M.L. and N.D.; methodology, W.C., C.G.-L., M.L. and N.D.; software, W.C. and M.L.; validation, W.C., M.L. and N.D.; formal analysis, W.C. and M.L.; investigation, W.C. and M.L.; resources, W.C.; data curation, W.C., C.G.-L., M.L. and N.D.; writing—original draft preparation, W.C.; writing—review and editing, W.C., C.G.-L., M.L. and N.D.; visualization, C.G.-L.; supervision, W.C. and N.D.; project administration, W.C. and N.D.; funding acquisition, W.C. and N.D. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by VoLo Foundation through the VISTA Award 2023.

Institutional Review Board Statement

This study was approved by the Institutional Animal Care and Use Committee at the University of Florida (Protocol IACUC No 202200000038).

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this paper are available from the corresponding author on request.

Acknowledgments

The authors acknowledge the support of Georgia Dubeux, Araceli Maderal, Ignacio Fernandez-Marenchino, and Federico Tarnonsky for their collaboration during the field and lab work. The authors also acknowledge the support of Luana Danta Queiroz for his help with the information about pigweed encroachment and DiLorenzo’s lab team for supporting the batch culture incubations and sample processing. Likewise, the authors acknowledge support from VoLo foundation.

Conflicts of Interest

The funders had no role in the design of this study; in the collection, analyses, or interpretation of the data; in the writing of the manuscript; or in the decision to publish the results.

References

  1. (FAO)-UN. How to Feed the World in 2050; Food and Agricultural Organization of the United Nations: Rome, Italy, 2009. [Google Scholar]
  2. Drouillard, J.S. Current situation and future trends for beef production in the United States of America—A review. Asian-Australas. J. Anim. Sci. 2018, 31, 1007–1016. [Google Scholar] [CrossRef] [PubMed]
  3. (FAO) FaAOotUN. FAOSTAT; Food and Agricultural Organization of the United Nations: Rome, Italy, 2021. [Google Scholar]
  4. Greenwood, P.L. Review: An overview of beef production from pasture and feedlot globally, as demand for beef and the need for sustainable practices increase. Animal 2021, 15, 100295. [Google Scholar] [CrossRef]
  5. Rotz, C.A.; Isenberg, B.J.; Stackhouse-Lawson, K.R.; Pollak, E.J. A simulation-based approach for evaluating and comparing the environmental footprints of beef production systems. J. Anim. Sci. 2013, 91, 5427–5437. [Google Scholar] [CrossRef] [PubMed]
  6. Rotz, C.A.; Asem-Hiablie, S.; Dillon, J.; Bonifacio, H. Cradle-to-farm gate environmental footprints of beef cattle production in Kansas, Oklahoma, and Texas. J. Anim. Sci. 2015, 93, 2509–2519. [Google Scholar] [CrossRef]
  7. Place, S.E.; McCabe, C.J.; Mitloehner, F.M. Symposium review: Defining a pathway to climate neutrality for US dairy cattle production. J. Dairy Sci. 2022, 105, 8558–8568. [Google Scholar] [CrossRef] [PubMed]
  8. Miller, S.M.; Wofsy, S.C.; Michalak, A.M.; Kort, E.A.; Andrews, A.E.; Biraud, S.C.; Dlugokencky, E.J.; Eluszkiewicz, J.; Fischer, M.L.; Janssens-Maenhout, G.; et al. Anthropogenic emissions of methane in the United States. Proc. Natl. Acad. Sci. USA 2013, 110, 20018–20022. [Google Scholar] [CrossRef]
  9. Smith, P.E.; Kelly, A.K.; Kenny, D.A.; Waters, S.M. Enteric methane research and mitigation strategies for pastoral-based beef cattle production systems. Front. Vet. Sci. 2022, 9, 958340. [Google Scholar] [CrossRef]
  10. Arndt, C.; Hristov, A.N.; Price, W.J.; McClelland, S.C.; Pelaez, A.M.; Cueva, S.F.; Oh, J.; Dijkstra, J.; Bannink, A.; Bayat, A.R.; et al. Full adoption of the most effective strategies to mitigate methane emissions by ruminants can help meet the 1.5 C target by 2030 but not 2050. Proc. Natl. Acad. Sci. USA 2022, 119, e2111294119. [Google Scholar] [CrossRef]
  11. Grainger, C.; Clarke, T.; Auldist, M.J.; Beauchemin, K.A.; McGinn, S.M.; Waghorn, G.C.; Eckard, R.J. Potential use of Acacia mearnsii condensed tannins to reduce methane emissions and nitrogen excretion from grazing dairy cows. Can. J. Anim. Sci. 2009, 89, 241–251. [Google Scholar] [CrossRef]
  12. Chen, M.-S. Inducible direct plant defense against insect herbivores: A review. Insect Sci. 2008, 15, 101–114. [Google Scholar] [CrossRef]
  13. Wallace, R.J.; McEwan, N.R.; McIntosh, F.M.; Teferedegne, B.; Newbold, C.J. Natural products as manipulators of rumen fermentation. Asian Australas. J. Anim. Sci. 2002, 15, 1458–1468. [Google Scholar] [CrossRef]
  14. Chen, R.J.; Chung, T.Y.; Li, F.Y.; Lin, N.H.; Tzen, J.T. Effect of sugar positions in ginsenosides and their inhibitory potency on Na+/K+-ATPase activity. Acta Pharmacol. Sin. 2009, 30, 61–69. [Google Scholar] [CrossRef] [PubMed]
  15. Tavendale, M.H.; Meagher, L.P.; Pacheco, D.; Walker, N.; Attwood, G.T.; Sivakumaran, S. Methane production from in vitro rumen incubations with Lotus pedunculatus and Medicago sativa, and effects of extractable condensed tannin fractions on methanogenesis. Anim. Feed. Sci. Technol. 2005, 123, 403–419. [Google Scholar] [CrossRef]
  16. Olagaray, K.E.; Bradford, B.J. Plant flavonoids to improve productivity of ruminants–A review. Anim. Feed. Sci. Technol. 2019, 251, 21–36. [Google Scholar] [CrossRef]
  17. Daehler, C.C. Performance comparisons of co-occurring native and alien invasive plants: Implications for conservation and restoration. Annu. Rev. Ecol. Evol. Syst. 2003, 34, 183–211. [Google Scholar] [CrossRef]
  18. Dantas-Queiroz, L.M.; Dubeux, J.C.B., Jr.; Sollenberger, L.E.; Wallau, M.O.; Loures, D.R.S.; da Silva Santos, E.R.; Oduor, K.; Araujo Bernardini, M.; Novo, S.F.; Pereira Neto, J.; et al. Spiny Pigweed Encroachment Affects Animal and Herbage Performance. In Proceedings of the ASA, CSSA, SSSA International Annual Meeting, Baltimore, MD, USA, 6–9 November 2022; Available online: https://scisoc.confex.com/scisoc/2022am/meetingapp.cgi/Paper/143947 (accessed on 17 September 2024).
  19. Sarker, U.; Oba, S. Nutraceuticals, antioxidant pigments, and phytochemicals in the leaves of Amaranthus spinosus and Amaranthus viridis weedy species. Sci. Rep. 2019, 9, 20413. [Google Scholar] [CrossRef]
  20. UF IFAS Extension University of Florida. How Weed Infestations in Pastures affect Cattle Performance?—Panhandle Agriculture; UF IFAS Extension University of Florida: Gainesville, FL, USA, 2021; Available online: https://nwdistrict.ifas.ufl.edu/phag/2021/08/06/how-weed-infestations-in-pastures-affect-cattle-performance/ (accessed on 17 September 2024).
  21. Tilley, J.M.A.; Terry, R.A. A two-stage technique for the in vitro digestion of forage crops. Grass Forage Sci. 1963, 18, 104–111. [Google Scholar] [CrossRef]
  22. Ruiz-Moreno, M.; Binversie, E.; Fessenden, S.W.; Stern, M.D. Mitigation of in vitro hydrogen sulfide production using bismuth subsalicylate with and without monensin in beef feedlot diets. J. Anim. Sci. 2015, 93, 5346–5354. [Google Scholar] [CrossRef]
  23. Broderick, G.A.; Kang, J.H. Automated simultaneous determination of ammonia and total amino acids in ruminal fluid and in vitro media. J. Dairy Sci. 1980, 63, 64–75. [Google Scholar] [CrossRef]
  24. Larrauri, M.; Asensio, C.M.; Martin, M.P.; Quiroga, P.R.; Grosso, N.R.; Nepote, V. Soymilk stability increase using polyphenols microcapsules. J. Food Sci. Technol. 2023, 61, 688–696. [Google Scholar] [CrossRef]
  25. Larrauri, M.; Zunino, M.P.; Zygadlo, J.A.; Grosso, N.R.; Nepote, V. Chemical characterization and antioxidant properties of fractions separated from extract of peanut skin derived from different industrial processes. Ind. Crops Prod. 2016, 94, 964–971. [Google Scholar] [CrossRef]
  26. Luximon-Ramma, A.; Bahorun, T.; Soobrattee, M.A.; Aruoma, O.I. Antioxidant activities of phenolic, proanthocyanidin, and flavonoid components in extracts of Cassia fistula. J. Agric. Food Chem. 2002, 50, 5042–5047. [Google Scholar] [CrossRef] [PubMed]
  27. Quiroga, P.R.; Riveros, C.G.; Zygadlo, J.A.; Grosso, N.R.; Nepote, V. Antioxidant activity of essential oil of oregano species from Argentina in relation to their chemical composition. Int. J. Food Sci. Technol. 2011, 46, 2648–2655. [Google Scholar] [CrossRef]
  28. Huff, S.; Rhodes, G.N., Jr. Pasture Weed Fact Sheet Spiny Amaranth. Real Life Solutions; UT Extension, Department of Agriculture, University of Tennessee: Knoxville, TN, USA, 2023; Available online: https://utia.tennessee.edu/publications/wp-content/uploads/sites/269/2023/10/W353.pdf (accessed on 17 September 2024).
  29. De Andrade Neto, A.Q.; de Almeida Souza, J.C.; de Mendonça, C.L.; Correa, F.R. Intoxicação natural por Amaranthus spinosus (Amaranthaceae) em bovinos no Agreste do Estado de Pernambuco. Rev. Bras. Zootec. 2016, 45, 1–7. [Google Scholar]
  30. Sun, X.; Cheng, L.; Jonker, A.; Munidasa, S.; Pacheco, D. A Review: Plant Carbohydrate Types—The Potential Impact on Ruminant Methane Emissions. Front. Vet. Sci. 2022, 9, 880115. [Google Scholar] [CrossRef]
  31. Hazarika, J.M.; Talukdar, B.; Choudhury, P.; Chakraborty, S.; Kumar, S.; Sarkar, S. Effect of Amaranthus spinosus (Meghnads) and Turmeric (Curcuma longa) on Chronic Arsenicosis in Cattle. J. Appl. Pharm. Sci. 2012, 2, 163–166. Available online: https://japsonline.com/abstract.php?article_id=480&sts=2 (accessed on 1 September 2024).
  32. Amabye, T.G. Evaluation of Physiochemical, Phytochemical, Antioxidant and Antimicrobial Screening Parameters of Amaranthus spinosus Leaves. Nat. Prod. Chem. Res. 2015, 4, 199. [Google Scholar] [CrossRef]
  33. Rodríguez-Prado, M.; Calsamiglia, S.; Ferret, A. Effects of fiber content and particle size of forage on the flow of microbial amino acids from continuous culture fermenters. J. Dairy Sci. 2004, 87, 1413–1424. [Google Scholar] [CrossRef]
  34. Wang, L.; Zhang, G.; Li, Y.; Zhang, Y. Effects of high forage/concentrate diet on volatile fatty acid production and the microorganisms involved in VFA production in cow rumen. Animals 2020, 10, 223. [Google Scholar] [CrossRef]
  35. Teng, Z.; Liu, S.; Zhang, L.; Zhang, L.; Liu, S.; Fu, T.; Zhang, N.; Gao, T. Tea polyphenols inhibit methanogenesis and improve rumen epithelial transport in dairy cows. Animals 2024, 14, 2569. [Google Scholar] [CrossRef]
  36. Ban, C.; Tian, X.; Lu, Q.; Lounglawan, P.; Wen, G. Enhancing rumen fermentation and bacteria community in sika deer (Cervus nippon) through varying levels of dragon fruit peel polyphenolic extract: An in vitro study. Animals 2024, 14, 1139. [Google Scholar] [CrossRef] [PubMed]
  37. Hassan, U.F.; Hassan, H.F.; Baba, H.; Lawan, D.A.; Hassan, H.F.; Ajiya, D.A.; Sambo, A.M.; Shehu, K.L.; Bello, U.; Muhammad, K.Y.; et al. Feed quality potentials of the aerial parts of Amaranthus spinosus. Chem. Res. J. 2020, 5, 29–36. [Google Scholar]
  38. Sutton, J.D. The fermentation of soluble carbohydrates in rumen contents of cows fed diets containing a large proportion of hay. Br. J. Nutr. 1968, 22, 689–712. [Google Scholar] [CrossRef] [PubMed]
  39. Lee, M.R.F.; Merry, R.J.; Davies, D.R.; Moorby, J.M.; Humphreys, M.O.; Theodorou, M.K.; MacRae, J.C.; Scollan, N.D. Effect of increasing availability of water-soluble carbohydrates on in vitro rumen fermentation. Anim. Feed. Sci. Technol. 2003, 104, 59–70. [Google Scholar] [CrossRef]
  40. Sinz, S.; Kunz, C.; Liesegang, A.; Braun, U.; Marquardt, S.; Soliva, C.R.; Kreuzer, M. In vitro bioactivity of various pure flavonoids in ruminal fermentation, with special reference to methane formation. Czech J. Anim. Sci. 2018, 63, 293–304. [Google Scholar] [CrossRef]
  41. Patra, A.K.; Min, B.R.; Saxena, J. Dietary tannins on microbial ecology of the gastrointestinal tract in ruminants. In Dietary Phytochemicals and Microbes; Patra, A., Ed.; Springer: Dordrecht, The Netherlands, 2012. [Google Scholar] [CrossRef]
  42. Makarewicz, M.; Drożdż, I.; Tarko, T.; Duda-Chodak, A. The interactions between polyphenols and microorganisms, especially gut microbiota. Antioxidants 2021, 10, 188. [Google Scholar] [CrossRef]
  43. Vasta, V.; Daghio, M.; Cappucci, A.; Buccioni, A.; Serra, A.; Viti, C.; Mele, M. Invited review: Plant polyphenols and rumen microbiota responsible for fatty acid biohydrogenation, fiber digestion, and methane emission: Experimental evidence and methodological approaches. J. Dairy Sci. 2019, 102, 3781–3804. [Google Scholar] [CrossRef]
  44. Ventura, M.; Canchaya, C.; Tauch, A.; Chandra, G.; Fitzgerald, G.F.; Chater, K.F.; van Sinderen, D. Genomics of Actinobacteria: Tracing the evolutionary history of an ancient phylum. Microbiol. Mol. Biol. Rev. 2007, 71, 495–548. [Google Scholar] [CrossRef]
  45. Goel, G.; Makkar, H.P.S. Methane mitigation from ruminants using tannins and saponins. Trop. Anim. Health Prod. 2012, 44, 729–739. [Google Scholar] [CrossRef]
  46. Yu, S.; Zhao, Y.; Li, L.; Zhao, H.; Liu, M.; Jiang, L. Flavonoids from citrus peel display potential synergistic effects on inhibiting rumen methanogenesis and ammoniagenesis: A microbiome perspective. Environ. Sci. Pollut. Res. 2024, 31, 21208–21223. [Google Scholar] [CrossRef]
  47. Jayanegara, A.; Leiber, F.; Kreuzer, M. Meta-analysis of the relationship between dietary tannin level and methane formation in ruminants from in vivo and in vitro experiments. J. Anim. Physiol. Anim. Nutr. 2011, 96, 365–375. [Google Scholar] [CrossRef] [PubMed]
  48. Yineger, H.; Kelbessa, E.; Bekele, T.; Molla, E.L. Plants used in traditional management of human ailments at Bale Mountains National Park, South Eastern Ethiopia. J. Med. Plants Res. 2008, 2, 132–153. [Google Scholar]
  49. Patra, A.K.; Saxena, J. Dietary phytochemicals as rumen modifiers: A review of the effects on microbial populations. Antonie Van Leeuwenhoek 2009, 96, 363–375. [Google Scholar] [CrossRef]
  50. Wang, K.; Xiong, B.; Zhao, X. Could propionate formation be used to reduce enteric methane emission in ruminants? Sci. Total Environ. 2023, 855, 158867. [Google Scholar] [CrossRef]
  51. Beauchemin, K.A.; McGinn, S.M.; Martinez, T.F.; McAllister, T.A. Use of condensed tannin extract from quebracho trees to reduce methane emissions from cattle. J. Anim. Sci. 2007, 85, 1990–1996. [Google Scholar] [CrossRef]
  52. Guglielmelli, A.; Calabro, S.; Primi, R.; Carone, F.; Cutrignelli, M.I.; Tudisco, R.; Danieli, P.P. In vitro fermentation patterns and methane production of sainfoin (Onobrychis viciifolia Scop.) hay with different condensed tannin contents. Grass Forage Sci. 2011, 66, 488–500. [Google Scholar] [CrossRef]
  53. Rufino-Moya, P.J.; Blanco, M.; Lobón, S.; Bertolín, J.R.; Armengol, R.; Joy, M. The inclusion of concentrate with quebracho is advisable in two forage-based diets of ewes according to the in vitro fermentation parameters. Animals 2019, 9, 451. [Google Scholar] [CrossRef]
  54. Bretas, I.L.; Dubeux, J.C.B., Jr.; Cruz, P.J.R.; Queiroz, L.M.D.; Ruiz-Moreno, M.; Knight, C.; Flynn, S.; Ingram, S.; Pereira Neto, J.D. Monitoring the Effect of Weed Encroachment on Cattle Behavior in Grazing Systems Using GPS Tracking Collars. Animals 2023, 13, 3353. [Google Scholar] [CrossRef]
  55. Hess, H.D.; Beuret, R.A.; Lötscher, M.; Hindrichsen, I.K.; Machmüller, A.; Carulla, J.E.; Lascano, C.E.; Kreuzer, M. Ruminal fermentation, methanogenesis, and nitrogen utilization of sheep receiving tropical grass hay-concentrate diets offered with Sapindus saponaria fruits and Cratylia argentea foliage. Anim. Sci. 2004, 79, 177–189. [Google Scholar] [CrossRef]
Table 1. Proportion of each morphological portion regarding the whole pigweed plant and chemical composition of the incubated forage and the selected morphological portions to be incubated.
Table 1. Proportion of each morphological portion regarding the whole pigweed plant and chemical composition of the incubated forage and the selected morphological portions to be incubated.
ItemBasal DietLeavesRoot Seed Stem WHO
Proportion 1, %-19.8 ± 6.913.49 ± 1.92.2 ± 1.659.6 ± 6.2-
Item, g/kg of DM
OM930.2 ± 2.7847.8 ± 3.1801.6 ± 2.4925 ± 3.4865.83 ± 2.8806.99 ± 5.6
CP91.5 ± 7.5294.1 ± 15.487.4 ± 7.2295.8 ± 12.489.51 ± 6.1202.82 ± 0.4
NDF727.0 ± 4.9412.3 ± 2.0624.7 ± 4.3309.4 ± 2.1675.9 ± 3.7481.07 ± 13.6
ADF362.6 ± 1.9167.1 ± 17.7499.4 ± 2.7111.9 ± 10.1431.1 ± 14.1288.07 ± 13.9
OM = Organic matter; CP = Crude Protein; NDF = Neutral Detergent Fiber; ADF = Acid Detergent Fiber; Basal Diet = Bermudagrass; WHO = Whole Pigweed Plant. 1 Proportion of each morphological portion from the pigweed plant.
Table 2. Volatile fatty acid concentration and molar proportion under three levels of inclusion of morphological portions of pigweed (Amaranthus spinosus).
Table 2. Volatile fatty acid concentration and molar proportion under three levels of inclusion of morphological portions of pigweed (Amaranthus spinosus).
Inc (%)Incubated Morphological Portion p-Value
VariableLeafRootSeedStemWHOSEMPIP × I
Acetate, mM045.6545.5945.6245.4245.360.870.050.130.02
2.546.5339.3646.5547.5047.61
541.3145.5044.7346.4341.57
1045.6342.7145.0045.9237.77
Propionate, mM024.1124.0824.1024.0023.970.480.250.030.06
2.526.3322.1926.1626.0226.66
523.6625.6525.1624.7323.52
1024.7123.2624.5124.6421.41
Butyrate, mM08.938.928.928.898.870.12<0.0010.0010.001
2.510.188.0610.099.649.37
59.308.849.428.978.27
109.328.639.088.767.84
BCVFA, mM04.574.574.574.554.540.09<0.0010.03<0.001
2.55.484.005.274.894.70
54.944.484.814.723.92
104.824.654.794.783.81
A:P01.881.881.881.871.870.020.27<0.0010.92
2.51.731.741.741.791.75
51.671.721.691.781.68
101.671.661.651.681.60
Total VFA, mM084.2884.1684.2283.8783.751.460.020.060.01
2.589.6874.5289.2189.1389.20
580.4385.3785.1185.8878.06
1085.4680.2484.2584.9471.76
Acetate, mol/100 mol053.7953.7153.7553.5353.450.220.02<0.0010.10
2.550.5751.4750.8551.9552.06
548.8050.7649.8251.3350.57
1048.0747.8847.9748.6447.36
Propionate, mol/100 mol028.4128.3828.4028.2828.240.210.29<0.0010.98
2.528.5928.9328.6228.4829.11
527.9128.2828.2327.3928.62
1025.9826.0626.3026.0926.80
Butyrate, mol/100 mol010.5210.5110.5110.4710.460.08<0.001<0.0010.01
2.511.0910.5811.0310.5410.24
510.999.8610.539.9410.08
109.839.709.729.299.85
BCVFA, mol/100 mol05.395.385.395.365.350.090.0010.0010.16
2.55.975.305.775.345.14
55.865.055.325.224.77
105.105.255.085.084.80
VFA = volatile fatty acids; Total VFA = total volatile fatty acids; A:P = acetate–propionate ratio; WHO = whole pigweed plant; BCVFA = branched chain VFA; SEM = standard error of the mean; P = portion; I = inclusion. No P × I was detected for the evaluated parameters.
Table 3. Orthogonal contrasts and comparisons for ruminal fermentation and methane production under three levels of inclusion of morphological portions of pigweed (Amaranthus spinosus).
Table 3. Orthogonal contrasts and comparisons for ruminal fermentation and methane production under three levels of inclusion of morphological portions of pigweed (Amaranthus spinosus).
VariableCON vs. Portion (p-Value)Contrast (p-Value)
VFA, mMLeafRootSeedsStemWHOLQC
      Acetate, mM0.810.100.260.030.060.030.800.36
      propionate0.540.130.210.350.200.180.040.08
      Butyrate<0.01<0.01<0.010.66<0.010.020.010.10
      BCVFA<0.010.010.010.27<0.00010.410.210.10
      A:P0.570.910.730.040.20<0.0001<0.010.17
      Total VFA0.330.050.130.090.030.040.390.11
VFA, mol/100 mol
      Acetate0.010.490.280.010.82<0.001<0.010.11
      propionate0.490.760.870.120.08<0.001<0.0010.47
      Butyrate<0.00010.070.02<0.010.07<0.001<0.010.09
      BCVFA<0.010.530.250.56<0.01<0.010.290.07
CH4
      mg/mL0.570.230.310.230.050.230.860.75
      mM/g DMI0.03<0.010.49<0.01<0.01<0.01<0.010.39
      mM/g OMF0.01<0.010.22<0.01<0.01<0.01<0.010.44
Fermentation
      IVOMD0.510.280.310.670.070.230.430.73
      OMF0.01<0.010.001<0.00010.02<0.0010.270.24
      pH0.310.70.860.020.28<0.001<0.0010.07
      Gas0.530.950.120.020.180.790.770.06
      NH3-N0.010.040.330.010.150.070.690.07
VFA = volatile fatty acids; BCVFA = branched chain volatile fatty acids; A:P = acetate–propionate ratio; Total VFA = total volatile fatty acids; CH4 = methane; DMI = dry mater incubated; OMF = organic matter fermented; IVOMD = in vitro organic matter digestibility; NH3-N = ammonia–nitrogen; CON = bermudagrass without the addition of any morphological portion of pigweed; WHO = whole pigweed plant; L = linear; Q = quadratic; C = cubic.
Table 4. Ruminal fermentation and methane production under three levels of inclusion of morphological portions of pigweed (Amaranthus spinosus).
Table 4. Ruminal fermentation and methane production under three levels of inclusion of morphological portions of pigweed (Amaranthus spinosus).
Inc (%)Incubated Morphological Portion p-Value
VariableLeafRootSeedStemWHOSEMPIP × I
CH4, mg/mL 00.0250.0270.0270.0270.0270.0040.180.660.10
2.50.0440.0380.0200.0400.006
50.0340.0320.0200.0300.033
100.0230.0360.0380.0370.028
CH4, mM/g DMI00.0440.0440.0650.0510.0640.004<0.01<0.01<0.01
2.50.0290.0540.0260.0580.022
50.0360.0520.0240.0420.019
100.0230.0520.0390.0540.015
CH4, mM/g OMF00.0060.0060.0080.0080.0090.001<0.001<0.001<0.001
2.50.0040.0080.0030.0080.003
50.0050.0070.0030.0050.002
100.0030.0070.0050.0070.002
IVOMD,%046.2250.8552.1756.2845.942.530.290.530.90
2.550.8657.9150.9655.2744.90
554.4857.6258.2153.4848.87
1053.1954.2858.6549.6954.12
pH06.326.286.186.086.240.020.24<0.0010.15
2.56.376.366.416.366.34
56.376.366.416.396.42
106.376.386.336.356.42
Gas09.439.389.239.099.330.220.140.290.72
2.59.809.149.5210.039.56
59.139.408.3210.038.53
109.469.379.0610.018.85
NH3-N09.429.379.339.289.230.21<0.001<0.0010.22
2.57.397.828.818.768.52
56.947.978.198.789.17
108.167.378.539.248.29
CH4 = methane; DMI = dry matted incubated; OMF = organic matter fermented; IVOMD = in vitro organic matter digestibility; NH3-N = ammonia–nitrogen; SEM = standard error of the mean; P = portion; I = inclusion; P × I = portion and inclusion interaction.
Table 5. Plant secondary metabolite contents of the morphological portions from Pigweed.
Table 5. Plant secondary metabolite contents of the morphological portions from Pigweed.
Item.LeavesRoot Seed Stem WHO
DPPH EC50 (µg/mL)169.8 ± 1.1 d4164 ± 10.1 a133.1 ± 2.4 e1639 ± 3.8 b1262.50 ± 5.4 c
Yield (mg/g)118.2 ± 18.3135.6 ± 9.7155.2 ± 19.3117.78 ± 7.1131.19 ± 7.9
Phenol mg GAE/g DM3.2 ± 0.1 a0.5 ± 0.02 c3.5 ± 0.1 a0.8 ± 0.03 c1.1 ± 0.04 b
Flavonoid mg QE/g DM19.7 ± 3.5 a1.3 ± 0.04 c3.7 ± 0.4 b1.7 ± 0.2 c1.9 ± 0.3 c
Data presented corresponded to average ± standard deviation. DPPH = diphenyl picryl hydrazyl radical; GAE = gallic acid equivalent; QE = quercetin equivalent; EC50 = is a measure of scavenging activity of sample concentration required to inhibit 50% radical; WHO = whole pigweed plant. Means with different letters were significantly different according to Tukey’s test (p < 0.05).
Table 6. Ruminal fermentation and methane production under two levels of inclusion of extracts from pigweed (Amaranthus spinosus).
Table 6. Ruminal fermentation and methane production under two levels of inclusion of extracts from pigweed (Amaranthus spinosus).
Incubated Extract p-ValueTend
WHOLeavesSEMPI
Inclusion (%)
VariableCON 2.552.55
VFA, mM
      Acetate60.263.656.2761.0562.545.120.630.85L
      Propionate15.716.215.715.8919.320.960.040.78L
      Butyrate6.46.45.546.457.090.660.080.76L
      Total VFA 85.388.8879.6985.9691.516.90.510.73L
      A:P 3.83.933.563.853.280.190.190.02L
VFA, mol/100 mol
      Acetate70.5771.6570.6171.0268.344.110.630.85L
      Propionate18.4318.2419.718.4921.110.540.040.78L
      Butyrate7.57.256.957.57.750.710.080.76L
CH4
      mM6.364.874.635.736.120.480.010.03L
      mmol/g OMF0.380.270.240.330.390.040.020.05L
Fermentation
            pH6.616.576.536.466.450.050.030.2L
            IVOMD, % 49.3453.2353.0651.4152.691.080.420.68L
            NH3-N, mM7.188.038.77.448.20.620.01<0.01
            Gas, mL/g OMF67.4156.4947.0758.69628.520.010.01Q
VFA = volatile fatty acids; Total VFA = total volatile fatty acids; A:P = acetate–propionate ratio; CH4 = methane; OMF = organic matter fermented; IVOMD = in vitro organic matter digestibility; NH3-N = ammonia–nitrogen; CON = bermudagrass without the addition of any morphological portion of Pigweed; WHO = whole pigweed plant; SEM = standard error of the mean; P = portion; I = inclusion. No P × I was detected for the evaluated parameters.
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Cuervo, W.; Larrauri, M.; Gomez-Lopez, C.; DiLorenzo, N. Invasive Pigweed (Amaranthus spinosus) as a Potential Source of Plant Secondary Metabolites to Mitigate Enteric Methane Emissions in Beef Cattle. Grasses 2025, 4, 14. https://doi.org/10.3390/grasses4020014

AMA Style

Cuervo W, Larrauri M, Gomez-Lopez C, DiLorenzo N. Invasive Pigweed (Amaranthus spinosus) as a Potential Source of Plant Secondary Metabolites to Mitigate Enteric Methane Emissions in Beef Cattle. Grasses. 2025; 4(2):14. https://doi.org/10.3390/grasses4020014

Chicago/Turabian Style

Cuervo, Wilmer, Mariana Larrauri, Camila Gomez-Lopez, and Nicolas DiLorenzo. 2025. "Invasive Pigweed (Amaranthus spinosus) as a Potential Source of Plant Secondary Metabolites to Mitigate Enteric Methane Emissions in Beef Cattle" Grasses 4, no. 2: 14. https://doi.org/10.3390/grasses4020014

APA Style

Cuervo, W., Larrauri, M., Gomez-Lopez, C., & DiLorenzo, N. (2025). Invasive Pigweed (Amaranthus spinosus) as a Potential Source of Plant Secondary Metabolites to Mitigate Enteric Methane Emissions in Beef Cattle. Grasses, 4(2), 14. https://doi.org/10.3390/grasses4020014

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