Oxidative stress and postmortem meat quality in crossbred lambs

Abstract

The objective of this study was to evaluate effects of different levels of lipopolysaccharide (LPS)-mediated oxidative stress on fresh meat quality. Crossbred lambs (n = 29) were blocked by weight and fed a standard finishing ration for the duration of the study. Lambs were individually housed and treatment groups were administered one of three intravenous injections every 72 h across a three-injection (9-day) cycle: saline control (control), 50 ng LPS/kg body weight (BW) (LPS50), or 100 ng LPS/kg BW (LPS100). Rectal temperatures were measured to indicate inflammatory response. Lambs were harvested at the Loeffel Meat Laboratory and 80 mg of pre-rigor Longissimus lumborum were collected in control and LPS100 treatments within 30 min postmortem for RNA analysis. Wholesale loins were split and randomly assigned 1 or 14 d of wet aging. Chops were fabricated after aging and placed under retail display (RD) for 0 or 7 d. Animal was the experimental unit. LPS-treated lambs had increased (P < 0.05) rectal temperatures at 1, 2, 4, and 24 h post-injection. Transcriptomics revealed significant (P_raw < 0.05) upregulation in RNA pathways related to generation of oxidative stress in LPS100 compared with control. A trend was found for tenderness (Warner–Bratzler shear force, WBSF; P = 0.10), chops from LPS50 having lower shear force compared with control at 1 d postmortem. Muscle from LPS50 treatment lambs exhibited greater troponin T degradation (P = 0.02) compared with all treatments at 1 d. Aging decreased WBSF (P < 0.0001), increased sarcoplasmic calcium concentration (P < 0.0001), pH (P < 0.0001), and proteolysis (P < 0.0001) across treatments. Following aging, chops increased discoloration as RD increased (P < 0.0001), with control chops aged 14 d being the most discolored. Chops from lambs given LPS had higher (P < 0.05) a* values compared with control at 14 d of aging. The L* values were greater (P < 0.05) in LPS100 compared with both LPS50 and control. Aging tended (P = 0.0608) to increase lipid oxidation during RD across either aging period. No significant differences (P > 0.05) in sarcomere length, proximate composition, fatty acid composition, or isoprostane content were found. These results suggest that defined upregulation of oxidative stress has no detriment on fresh meat color, but may alter biological pathways responsible for muscle stress response, apoptosis, and enzymatic processes, resulting in changes in tenderness early postmortem.

Introduction

During postmortem aging, the tenderization of muscle relies on several biological mechanisms, including endogenous enzymes, such as calpains, caspases, and proteasomes (Ouali et al., 2006; Bhat et al., 2018). These endogenous mechanisms activate early postmortem with the utilization of free calcium released from mitochondria and sarcoplasmic reticulum (Rossi and Dirksen, 2006; Santo-Domingo and Demaurex, 2010). With regards to postmortem aging, an increasing number of investigations have focused on the influence of apoptosis on meat quality. Kemp and Parr (2012) indicated apoptotic events can contribute to proteolytic degradation of many structural proteins during muscle aging.

Apoptosis is the mechanism responsible for regulated cell death, portrayed by multiple biochemical and molecular pathways (Sierra and Olivan, 2013). This intrinsic mechanism involves mitochondria, as increased apoptotic events promote mitophagy, indicated by the release of cytochrome c from the mitochondrial membrane into the cytoplasm (Ott et al., 2007; Kagan et al., 2009; Ke et al., 2017). Cytosolic cytochrome c binds to Apaf-1 genes and produce apoptosomes, large quaternary proteins used to initiate the caspase system and begin proteolytic events (Porter and Jänicke, 1999; Momeni, 2011). Initiation of apoptotic activity has been linked to oxidative stress-mediated events (Slimen et al., 2014).

The onset of oxidative stress results from the overwhelming production of reactive oxygen species (ROS) compared with homeostatic endogenous antioxidants present within the system (Powers et al., 2011). As the name implies, ROS are highly reactive substances primarily produced as byproducts during oxidative phosphorylation (Paradies et al., 2001). Additionally, ROS act as signaling molecules to upregulate a homeostatic response (Dröge, 2002). This action facilitates antioxidant enzymes (superoxide dismutase, catalase, and glutathione) capable of changing ROS to more stable products within muscle cells. However, conditions of oxidative stress overwhelm antioxidant function, allowing ROS to alter protein, lipid, or nucleic acid morphology and functionality (Bekhit et al., 2013). Concurrently, ROS can interact with nitrosative species (nitric oxide), producing hybrid radicals which can target proteins responsible for organelle function (Stamler and Meissner, 2001). Given the right conditions, oxidative stress factors have been theorized to impact muscle cells and create conditions critical to meat quality.

Intrinsic and extrinsic conditions promoting oxidative stress have been implicated to alter meat quality (Warner et al., 2005; Ponnampalam et al., 2017; Wang et al., 2018; Mitacek et al., 2019). However, there is inconsistency within the literature describing the impact of oxidative stress factors on meat quality, as this phenomenon is likely dependent on a multitude of factors. These factors include degree and duration of oxidative stress, source of oxidative stress generation (in vivo vs. in vitro), the influence of individual oxidative and nitrosative species on cellular constituents, and composition of muscle tissue (Cottrell et al., 2015; Niu et al., 2016; Ke et al., 2017; Wang et al., 2018). Concurrently, there is a gap in the literature relating specific genetic pathways and their oxidative stress-mediated regulation to the impact of meat quality.

The mechanism of meat tenderization is well-recognized. However, the impact of oxidative stress on skeletal muscle, and its influence on factors critical to meat quality has yet to be understood. Additionally, the impact of oxidative stress and its relationship toward the muscle transcriptome are not fully understood. Therefore, the objective of this study was to evaluate the effects of controlled oxidative stress in vivo on oxidative biomarkers and meat quality attributes, including tenderness, color stability, and lipid oxidation.

Materials and Methods

All animal use protocols were approved by the University of Nebraska-Lincoln’s Institutional Animal Care and Use Committee (Protocol No. 1751).

Lambs

A total of 29 cross-bred (Hampshire × Dorset × Polypay) wethers (initial body weight [BW] = 29 ± 2.68 kg) were group housed (University of Nebraska Life Sciences Annex in Lincoln, NE) for 42 days on a standard finishing ration developed at Eastern Nebraska Research and Extension Center (ENREC; Mead, NE). Lambs were then individually housed, and underwent a 22-day acclimation period preceding the immune challenge, wherein all lambs were introduced to human handling, being placed on trim stands for intravenous injections, and rectal temperatures measured prior to the immune challenge. After the acclimation period, lambs were stratified by BW, 50.8 ± 2.7 kg, and blocked into treatment groups with treatment being randomly assigned within a block. Lambs were individually fed according to BW to maintain a 0.34 kg/day weight gain for the remainder of the in vivo analysis.

Lipopolysaccharide treatments

Lambs were randomly assigned a 2 ml intravenous injection of either saline control (control, n = 10), 50 ng lipopolysaccharide/kg BW (LPS 50, n = 9), or 100 ng LPS/kg BW (LPS 100, n = 10) treatment. Concentrations of LPS O111: B4 (Sigma-Aldrich, St. Louis, MO; L2630) were determined based on previous research performed at University of Nebraska-Lincoln using sheep as a model for LPS injections (Cadaret et al., 2019a; Posont et al., 2019). Three injections were administered across a 9-day immune challenge, with a subsequent injection occurring every 72 h period. Injection times were partitioned across the 9-day immune challenge, with two sets of 14–15 lambs in each group. Two immune challenge groups were completed in succession with lambs blocked by BW, designed so all animals would reach a similar final BW prior to harvest. All methods of live animal handling were approved by UNL Institutional Animal Care and Use Committee (IACUC).

Rectal temperatures

Rectal temperatures were measured at 0, 1, 2, 4, 8, 12, 24, 48, and 72 h post-injection time (0600 hours). Lambs were secured on a trim stand prior to taking readings. Rectal temperature was measured using a Vicks Thermometer (ComfortFlex, Marlborough, MA; V966US) by inserting 2.54 cm in from the tip of the anus to the rectum of the lamb and held for 10 s. Temperatures were recorded as degrees centigrade.

Muscle sample collection

Upon completion of the final 72 h cycle, lambs were held 48 h prior to slaughter, then transported from the live animal unit within the Animal Science Complex on East Campus to the Loeffel Meat Laboratory, which is in the same Complex (Lincoln, NE). Within 30 min of exsanguination, an 80 mg pre-rigor sample was taken from the posterior end of the Longissimus lumborum from control (n = 8) and LPS100 (n = 8) treatment groups. Samples were extracted using a scalpel blade and sampling station sterilized in ethanol in between sampling (Feather Safety Razor Co. LTD., Osaka, Japan; No. 11 2976). Pre-rigor samples were removed of exterior fat and connective tissue, then placed in a 2 mL cryotube (Cryogenic Vial CryoClear 3012 Globe Scientific, Mahwah, NJ) and frozen for future analysis (−80°C). The carcasses were then chilled for 24 h.

Fabrication

After 24 h postmortem, carcasses were fabricated and the loin portion from the 9th rib to the last lumbar vertebrae was retained. Each loin section was split down the spine using a band-saw (Biro MFG. Co., Marblehead, OH; Model 3334). Within each animal, sides were randomly assigned to one of two aging periods (1 or 14 d). On d 1, longissimus muscle from one side of each loin was deboned and removed of excess subcutaneous fat. Beginning at the posterior end, one 5.08 cm chop was cut and utilized for all laboratory analysis at each aging period. One 2.54 cm chop at each aging period was utilized for analysis of pH and proximate composition. Three 2.54 cm chops were fabricated adjacent to one another for each aging period and analyzed for tenderness using Warner–Bratzler shear force (WBSF) for 0 d retail display (RD). One 2.54 cm chop was cut to measure lipid oxidation via thiobarbaturic acid reactive substances (TBARS) for 0 d of RD. One 2.54 cm chop at each aging period was trimmed of all subcutaneous fat, and utilized to measure visual discoloration, objective color, and lipid oxidation for 7 d of RD. One 2.54 cm chop at the anterior end at each aging period was cut for extra laboratory sample, if needed. At d 1 aging, chops for laboratory analysis, pH and proximate composition, laboratory analysis were vacuum packaged (MULTIVAC 500, Multivac, Inc., Kansas City, MO) in Prime Source Vacuum pouches (3 mil STD barrier, Prime Sources, St. Louis, MO). Chops for laboratory analysis, pH, and proximate composition were frozen for further analysis (−80°C). The opposite loin halves were lined with Boneguard (Boneguard Traditional Perforated, JVR Industries, Lancaster, NY), vacuum packaged, and aged (2°C) under dark storage for 14 days total. All chops were separated from the loin starting from the anterior end of the loin. The same fabrication sequence was used for both aging periods. Samples for color, lipid, and protein oxidation analysis were placed on foam trays (21.6 × 15.9 × 2.1 cm, Styro-Tech, Denver, CO) and overwrapped with an oxygen permeable film (Prime Source PSM 18 #75003815, Bunzl Processors Division, North Kansas City, MO). Trays were placed under simulated RD conditions for 7 d (3°C under white fluorescence lighting at 1,000–1,800 lux) and randomly rotated daily. All frozen chops utilized for laboratory analysis and lipid oxidation were removed from −80°C, and tempered for 30 min at room temperature, enough to finely dice, freeze in liquid nitrogen and then powdered in a metal cup blender (Model 51BL32, Waring Commercial, Torrington, CT), and held at −80°C for up to 23 d until further analysis.

RNA transcriptomics

Total RNA isolation from muscle was completed using the RNeasy Fibrous Tissue Mini Kit (QIAGEN, #74704, Hilden, Germany) and RNase-Free DNase Set (QIAGEN, #79254) as shown by Cadaret et al. (2019b). Samples were sent to the University of Nebraska-Medical Center (Next Generation Sequencing, Genome Core Facility, Omaha, NE) for poly-A+ library preparation and sequencing (150 bp paired-end) with a targeted coverage of 20 million reads/sample.

Resulting data were quality trimmed using Trim Galore! and aligned to the Oar_rambouillet_v1.0 reference genome STAR (Dobin et al., 2016). Differential expression (control vs. LPS100 treatment) was evaluated using transcript counts in DESeq2 (Love et al., 2014). Loci with P_adj < 0.05 were considered to be differentially expressed; those with P_raw < 0.05 were utilized for pathway exploration in Ingenuity Pathway Analysis (Qiagen).

Warner–Bratzler shear force

Three chops (2.54 cm) from each side were measured for tenderness via WBSF per sample. Internal temperatures were measured prior to cooking using a quick disconnect T-type thermocouple (TMQSS-062U-6, OMEGA Engineering, Inc., Stamford, CT) with a handheld thermometer (OMEGA 450-ATT, OMEGA Engineering, Inc.) in the geometric center of steaks shortly after removal from refrigeration. Weights of chops were collected prior to cooking using a precision balance scale (PL6001E, Mettler Toledo, Hogentogler and Co. Inc., Columbia, MO). All chops were cooked to an internal temperature of 35°C and turned over until they reached a target internal temperature of 70°C on an electric indoor grill (Hamilton Beach-31605A, Hamilton Beach Brands, Glen Allen, VA). After cooking, final weights were recorded. The steak was then bagged (PB-90-C, 0.85 mil., 6 × 3 × 15 in. PITT PLASTICS, Pittsburg, KS) and stored overnight at 2°C. The following day, 2 (1.27 cm diameter) cores per chop were removed with a drill press parallel to muscle fibers and sheared using a Food Texture Analyzer (TMS-Pro, Food Technology Corp., Sterling, VA.) with a triangular Warner–Bratzler blade. The mean of six cores was calculated for statistical analysis.

Troponin T and desmin degradation

Troponin T degradation was quantified according to the procedure described by Chao et al. (2018) as modified by Ribeiro et al. (2019) and desmin degradation was evaluated according to the procedure described by Carlson et al. (2017), with slight modifications. The procedures were quite similar and are thus described together. Proteins (4 mg/mL) were loaded (30 μL for troponin T degradation and 10 μL for desmin degradation) onto 4–20% Mini-PROTEAN TGX precast polyacrylamide gels (Bio-Rad Laboratories, Hercules, CA) with a 5 μL (troponin T) or 10 μL (desmin) pre-stained standard (Precision Plus Protein Kaleidoscope, #1610375, Bio Rad) using a Bio-Rad Mini-PROTEIN 2 Cells system (Bio-Rad Laboratories), and run at a constant voltage of 200 V (troponin T) or 360 V (desmin) for 45 min with an electrophoresis buffer (1X Tris/Glycine/SDS, #161–0732, Bio-Rad Laboratories). Proteins in the gels were blotted (60 min at 180 mA for troponin T or 90 min at 90 V for desmin) with ice-cold transfer buffer (25 mM Tris-base, 192 mM Glycine, 20% methanol; pH at 9.2). Membranes were blocked for 60 min in Odyssey Blocking Buffer (LI-COR, Lincoln, NE) and incubated for 60 min at room temperature in monoclonal anti-Troponin-T antibody (JLT-12 Sigma-Aldrich) or monoclonal anti-desmin antibody (DE-U-10 Sigma-Aldrich) at a dilution of 1:10,000 in Odyssey blocking buffer containing 0.2% TWEEN (troponin T) or 0.2% TWEEN-20 and 5% nonfat dry milk (desmin). Membranes were incubated overnight at 4°C, washed with tris buffered saline (containing 0.1% TWEEN-20 [TBST] for 40 min for troponin T) or tris buffered saline (containing 0.2% TWEEN-20 [TBST] for 10 min for desmin). The membranes were then incubated in IRDye 680 LT Conjugated Goat Anti-Mouse IgG1 secondary antibody (LI-COR) at a dilution of 1:10,000 in Odyssey blocking buffer containing 0.2% TWEEN-20 for 60 min. Membranes for troponin T were washed with tris buffered saline containing 0.1% TWEEN-20 for 30 min, then with tris buffered saline for 30 min. Membranes for desmin were washed three times with TBST. After washing, membranes were scanned using Odyssey Infrared Imaging system (LI-COR) at 700 nm. Degradation was evaluated by quantifying band intensities (pixel intensity) using Odyssey application software version 1.1. For troponin T, bands at 37 kDa were designated as intact and bands at 30 kDa were designated as degraded. For desmin, bands at 55 kDa were designated as intact and bands at 38 kDa were designated as degraded. Percent degradation was calculated by (intensity of degraded bands/intensity of intact bands) * 100.

Free calcium concentration

Free calcium was quantified according to the procedure described by Hart et al. (2019). About 3 g of powdered sample were centrifuged at 196,000 × g (Beckman Optima XPN-90 Ultracentrifuge, Type 50.2 Ti rotor, Beckman Coulter, Brea, CA) at 4°C for 30 min. About 700 μL of the supernatant were collected and treated with 0.1 mL of 27.5 trichloroacetic acid (TCA). Samples were centrifuged at 6,000 × g (accuSpin Micro 17R, ThermoFisher Scientific, Waltham, MA) for 10 min at 4°C. About 400 μL of supernatant were transferred to a syringe, and the volume was brought to 4 mL with deionized, distilled water. The diluted sample was filtered through a 13 mm diameter Millex-LG 0.20 μm syringe filter (Milliore, Bedford, MA). Calcium concentration was quantified at Ward Laboratories (Kearney, NE) using an inductively coupled plasma emission spectrometer (iCAP 6500 Radial; Thermo Electron, Cambridge, UK) with an appropriate calcium concentration standard.

Isoprostanes

Meat samples were analyzed using OxiSelect 8-iso-Prostaglandin F2aplha ELISA Kit (Cell BioLabs, INC., STA-337, San Diego, CA). Results were read immediately on a microplate reader using 450 nm as the target wavelength. Units of isoprostane content were designated as pg/mL.

Sarcomere length

Sarcomere length was determined using the helium–neon laser diffraction method described by Cross et al. (1981) and Dolazza and Lorenzen (2014).

Proximate composition

Proximate composition was evaluated using methods identical to Ribeiro et al. (2019). Moisture and ash percentage were quantified using a LECO Thermogravimetric Analyzer in duplicate (Model 604-100-400, LECO Corporation, St. Joseph, MI). Fat content was quantified, in triplicate, using ether extraction in accordance with the Soxhlet procedure-AOAC method 920.39 (1990). Protein content was calculated by the difference of 100 − (moisture + ash + fat).

Fatty acids

Fatty acid profiles were obtained via gas chromatography as described by Folch et al. (1957). After extraction, lipids were converted to fatty acid methyl esters according to Morrison and Smith (1964) and Metcalfe et al. (1966), with modifications as described by Hart et al. (2019). The percentage of fatty acids were determined by the peak areas in the chromatograph and values were converted to mg/100 g tissue: Fatty acid mg/100g tissue = (% of fatty acid peak area * fat content of samples) * 100.

pH analysis

Powdered sample from chops from all aging periods with 0 d RD were weighed in 5 g duplicates into 250 mL plastic beakers and placed on a stir plate. About 45 mL of distilled deionized water and a magnetic stir bar were added to ensure constant mixing during the measurement process. The pH was measured using a pH meter (Orion 410Aplus: ThermoFisher Scientific; Waltham, MA) that was calibrated using 4.0, 7.0, 10.0 standards. The mean measurement of the duplicates was utilized for all analysis.

Instrumental color in simulated RD

Objective color measurements were taken once daily every 24 h for 7 days during simulated RD at all aging time points. Chops (2.54 cm) were placed on Styrofoam trays (21.6 × 15.9 × 2.1 cm, Styro-Tech, Denver, CO), overwrapped with oxygen permeable film (Prime Source PSM 18 #75003815, Bunzl Processors Division, North Kansas City, MO), and placed under RD conditions (3°C under white fluorescence lightening at 1,000–1,800 lux). Commission international de l’éclairage (CIE) L* a* b* values were obtained using a Minolta CR-400 colorimeter (Minolta, Osaka, Japan) set with a D65 illuminant, 2°C, with an 8 mm diameter measurement area. Three measurements were made per chop and the mean was calculated for statistical analysis. The colorimeter was calibrated daily with a white ceramic tile (Calibration Plate, Serial No. 14933058, Konica Minolta, Japan) lined with oxygen permeable film. Lightness (L*) is measured with a range from 0 (black) to 100 (white), a* measures redness with the range between red (positive) and green (negative), and b* is a measure of yellowness from yellow (positive) to blue (negative). Delta E measures the magnitude of difference in L*, a*, and b* color space. Delta E was calculated for all chops across both aging periods in accordance with Hunt and King (2012). Color readings were recorded at the same time each day.

Subjective discoloration in simulated RD

Visual discoloration was assessed daily every 24 h during the 7 d of RD utilizing five trained panelists comprised of graduate students from the University of Nebraska. Panelists were trained using a standardized discoloration guide. Discoloration percent was estimated to the nearest percent from 0% to 100% with 0% meaning no discoloration present and 100% being a fully discolored chop. A reference guide showing discoloration in 10% increments (0%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, and 100%) was provided for panelists to provide consistent evaluation of chop discoloration. Chops were randomly relocated daily along the display tables to avoid any variable change due to light intensity (1,000–1,800 lux) during RD.

Lipid oxidation

Lipid oxidation was determined using TBARS values for all aging periods at 0 and 7 d RD described by Ahn et al. (1998), as modified by Henriott et al. (2020). Results were expressed in mg of malonaldehyde per kg of tissue.

Statistical analysis

Statistical analysis was conducted with SAS (version 9.4, Cary, NC). Rectal temperature was analyzed as a repeated measures design with treatment as the whole plot and time as the repeated measures. Objective and subjective color data were analyzed as a split-plot repeated measures design with treatment as the whole-plot, aging period as the split-plot and RD as the repeated measures. Tenderness, troponin T, desmin, calcium, and pH were analyzed as a spilt-plot design with treatment as the whole-plot and aging period as the split-plot. Lipid oxidation was a split-split-plot design with treatment as the whole plot, aging period as the split-plot and RD time as the split-split-plot. Transcriptomics, sarcomere length, fatty acids, and isoprostanes were analyzed as a completely randomized design. Data were analyzed using the PROC GLIMMIX procedure of SAS with animal as the experimental unit for rectal temperature, sarcomere length, fatty acids, isoprostanes, and transcriptomics. Chop was the experimental unit for objective and subjective color, tenderness, proteolysis, calcium, pH, and lipid oxidation. Correlations were evaluated using the PROC CORR procedure of SAS across all postmortem analyses. All tenderness and proteolysis means within aging periods were separated using SLICE function in SAS. All means were separated using the LS MEANS statement with an α level of less than or equal to 0.05 and tendencies were considered at an α level of 0.06–0.10.

Results

Rectal temperatures

Treatment had a significant effect (P < 0.05) on rectal temperatures of lambs (Figure 1), and a time effect (P < 0.05) was identified. Lambs administered LPS50 and LPS100 had significantly greater (P < 0.05) rectal temperatures than lambs administered the saline control. Additionally, LPS-treated lambs exhibiting the greatest increase in rectal temperature at 1, 2, and 4 h post-injection, with a numerically greater rectal temperature in LPS100-treated lambs compared with LPS50 at each timepoint listed. These results are in agreement with Yates et al. (2011), who found LPS treatments to consistently increase rectal temperatures and peak at 4 h post-injection, followed by a steady decline to basal temperature around 24 h. The increase in rectal temperatures is indicative of oxidative stress caused by treatment with LPS (Sternberg, 2007; Powers et al., 2011; Yates et al., 2011).

Figure 1.

Rectal temperatures of lambs administered intravenous injections of control, LPS50, or LPS100. Superscripts denote statistical differences (P < 0.05) within day.

Transcriptomics

About 6,493 transcripts were differentially expressed in the skeletal muscle transcriptome comparing control and LPS100 treatments. Considering all transcripts with differential expression (P_raw < 0.05, N = 309), pathway analyses identified 68 conical pathways predicted to be altered due to treatments. Pathway analyses are illustrated in Table 1. In the LPS100-treated samples, genes with evidence of dysregulation due to treatment were found in pathways that predicted an upregulation cellular biosynthesis, oxidative stress generation, cellular defense systems, nucleic acid alteration, and skeletal muscle function.

Table 1.

Conical pathway expression between Con and 100 ng LPS treatments

Function	Pathways	Fold change (−log P-value)1	Z-score2
Cell biosynthesis and turnover	IGF-1	2.9	0.707
	EGF	2.6	0.816
	ErbB2-ErbB3	2.24	−0.447
	ILK	2.05	1
	cAMP	1.88	−0.302
	PI3K	1.63	1.414
	ERK5	1.46	1.342
	Ceramide	0.848	1.89
Nucleic modification	Unfolded protein	5.93	0.378
	Telomerase	3.43	0.707
	HMGB1	2.5	1.414
	EIF2	1.94	0.333
	Neurotrophin/TRK	1.91	0.816
	JAK/Stat	1.81	0.816
Oxidative response/autophagy	NRF2 oxidative stress resp.	6.48	1.265
	IL-6	3.49	−0.905
	p38 MAPK	3.09	1
	Sumoylation	2.35	1.633
	TNFR-2	2.19	2
	CXCR4	2.01	0.707
	IL-8	1.52	0.632
	NO/ROS prod. In Macrophages	1.31	0.333
	IL-3	1.31	1.342
Muscle function oxidative stress	eNOS signaling	2.63	−2.121
	Agrin	1.84	1.342
	Calcium signaling	1.81	−1.633
	PPARα/RXRα	1.65	−1.265
	D-myo-inositol-tetrakiphosphate	1.59	0.707

¹Pathway expressions are statistically significant (P < 0.05). ²Numerical values denote degree of z-score (positive = upregulated to 100 ng LPS; negative = upregulated to Con) and gross fold change (−log P-value).

Tenderness

WBSF for lambs across treatment and aging are shown in Table 2. No aging-time-by-treatment interaction (P = 0.13) was seen. A significant aging effect (P < 0.0001) was identified across aging periods, along with a trend (P = 0.10) across treatments. No aging time-by-treatment interaction (P = 0.13) was seen.

Table 2.

The effects of LPS inclusion (control, LPS50, and LPS100) and aging (1 or 14 d) on tenderness (WBSF), proteolysis (Troponin-T, Desmin), and isoprostane content (n = 29)

Variable	Aging	Treatment1				P-values
		Control	LPS50	LPS100	SEM	Trt	Age	Trt*Age
WBSF, kg	1	8.06x	6.59x	7.27x	0.39	0.10	<0.0001	0.13
	14	2.77y	2.42y	2.36y	0.13	0.90
Troponin-T degradation, %	1	6.85by	10.32ay	6.24by	1.08	0.02	<0.0001	0.88
	14	47.73x	49.19x	41.68x	8.33	0.78
Desmin degradation, %	1	3.01y	4.17y	3.54y	1.49	0.85	<0.0001	0.08
	14	55.02x	41.73x	54.94x	4.92	0.10
Isoprostane content2	1	165.51	239.51	219.95	30.61	0.21	n/a	n/a

¹Control, saline injection; LPS50, 50 ng LPS/kg BW; LPS100, 100 ng LPS/kg BW.

²Larger 8-isoprostane content indicates a greater degree of oxidative stress within the sample, pg/mL.

^a,bMeans within the same row with different superscripts denote treatment differences (P < 0.05).

^x,yMeans within the same column of a variable with different superscripts denote significant differences (P < 0.05).

A significant aging effect (P < 0.0001) was found, as 1 day-aged chops exhibited greater WBSF compared with 14 day-aged chops (7.31 and 2.52 kg, respectively). Treatment tended to affect WBSF (P = 0.10), as chops from lambs administered LPS50 had lower shear force values compared with chops from lambs administered saline (control; 4.51 and 5.41 kg, respectively), with no differences in shear force values between LPS100 and control. Interestingly, chops from lambs administered LPS50 had noticeably lower WBSF compared with control (6.59 and 8.06 kg, respectively) within 1 d aging. There were no differences (P > 0.05) in tenderness between LPS100 and control among aging periods, however, chops from lambs administered LPS100 had numerically lower shear force values than control at 1 day of aging (7.27 and 8.06 kg, respectively). Chops aged for 14 days did not differ in WBSF across treatments (P > 0.05).

Proteolysis (troponin T and desmin degradation)

After 1 day postmortem, a treatment effect was found (P = 0.02), with LPS50 samples having significantly greater percent troponin-T degradation compared with control and LPS100 (Table 2). No differences in degradation were seen at 14 days postmortem. A significant aging effect (P < 0.0001) was found, as 1 day-aged chops exhibited less troponin-T degradation compared with 14 day-aged chops. There was no aging time-by-treatment effect (P > 0.05). These results parallel the WBSF data.

In this study, a significant aging effect was found (P < 0.0001), as 1-day aged chops exhibited less desmin degradation compared with 14-day aged chops (3.57% and 50.6%, respectively; Table 2). An aging-by treatment effect trend (P = 0.08) was found. Within 1-day aging, no differences (P > 0.05) were found across treatments. However, both control and LPS100 tended (P = 0.10) to be greater in percent degradation compared with LPS50 at 14-days aging.

Free calcium concentration

Free calcium concentration for samples across treatments and aging are shown in Table 3. An aging effect was shown for free calcium, as 14 day-aged samples had notably higher (P < 0.0001) free (sarcoplasmic) calcium compared with 1 day-aged samples. No differences (P > 0.05) were found across treatments.

Table 3.

The effects of LPS inclusion (control, LPS50, and LPS100) and aging (1 or 14 d) on sarcomere length, proximate composition, pH, and free calcium (n = 29)

		Sarcomere length1	Proximate composition				pH2		Free calcium content3
		μm1	Moisture	Protein	Fat	Ash	Days aging		Days aging
Treatment4							1	14	1	14
Control		1.7	75.1	15.13	8.18	1.59	5.71x	5.84y	46.72x	108.02y
LPS50		1.73	75.51	14.43	8.29	1.77	5.68x	5.86y	40.71x	104.51y
LPS100		1.71	75.45	13.91	8.99	1.66	5.73x	5.91y	43.63x	103.12y
SEM		0.03	0.20	0.99	0.98	0.09	0.05		4.46
P-value	Trt	0.71	0.31	0.68	0.82	0.44	0.13		0.49
	Age	n/a	n/a	n/a	n/a	n/a	<0.0001		<0.0001
	Trt*Age	n/a	n/a	n/a	n/a	n/a	0.76		0.88

¹Sarcomere length denotes the distance from one Z-disk to another Z-disk, μm.

²The potential for hydrogen; lower values are more acidic, higher values are more alkaline.

³μg Ca²⁺/μg mitochondrial protein.

⁴Control, saline injection; LPS50, 50 ng LPS/kg BW; LPS100, 100 ng LPS/kg BW.

^x,yMeans with the same variable with different superscripts denote differences across aging (P < 0.05).

Isoprostane content

There were no differences (P = 0.21) across treatments for F₂-isoprostane content (Table 2). Interestingly, LPS50 and LPS100-treated samples had numerically greater total F₂-isoprostane content compared with control chops (239.51, 219.95, 165.51 pg/mL, respectively).

Sarcomere length and proximate analysis

No differences in sarcomere length among treatments were observed (P = 0.70; Table 3).

In this study, treatment had no effect (P ≥ 0.31) on proximate composition of 1 d aged chops (Table 3). The mean values for the proximate composition were: 75.35% moisture, 14.49% protein, 8.49% fat, and 1.67% ash.

Fatty acids

There were no differences (P ≥ 0.21) found in amount of saturated fatty acids, monounsaturated fatty acids, polyunsaturated fatty acids, or trans fats, saturated:unsaturated ratio, along with no differences (P > 0.05) in any individual fatty acids identified (Table 4).

Table 4.

Amount of fatty acids from lamb chops (L. lumborum) aged for 1 day from lambs administered saline control (0), 50, or 100 ng/kg BW of LPS (n = 29)¹

Fatty acid3	Treatments2			P-value
	Control	LPS50	LPS100
C 10:0	2.80	2.96	2.80	0.5146
C 12:0	8.11	4.71	9.08	0.549
C 13:0	1.41	0.96	0.002	0.5844
C 14:0	181.12	183.25	213.03	0.6693
C 14:1	14.54	13.17	16.72	0.7083
C 15:0	38.15	33.03	40.04	0.5734
C 15:1	72.22	64.09	63.96	0.8796
C 16:0	1,754.75	1,786.67	1980.12	0.7341
C 16: 1T	30.31	21.73	31.14	0.2356
C 16:1	152.23	149.30	182.38	0.442
C 17:0	156.03	134.15	151.33	0.6655
C 17:1	174.30	151.26	173.56	0.703
C 18:0	1,099.64	1,111.18	1,117.99	0.9944
C 18:1T	308.13	293.64	352.97	0.6015
C 18:1	3,195.42	3,282.10	3,543.10	0.7905
C 18:1V	117.31	125.53	139.87	0.5919
C 18:2T	26.38	30.49	34.56	0.3524
C 18:2	493.34	547.45	584.04	0.6917
C 18: 3w3	15.77	20.04	22.23	0.218
C 18: 3w6	5.82	6.72	6.99	0.7998
C 19:0	6.63	3.98	4.54	0.6645
C 20:0	0.87	0.52	5.55E−17	0.4054
C 20:1	35.2	36.45	37.92	0.9419
C 20:2	12.62	12.88	9.48	0.6035
C 20: 4w6	170.12	175.81	166.74	0.9781
C 20:5	0.53	1.20	2.78E−18	0.5108
C 22:0	10.92	11.34	9.97	0.8824
C 22:5	13.64	14.10	14.71	0.9647
C 22:6	0	1.09	0	0.3413
C 24:0	19.96	19.59	18.83	0.9686
C 24:1	0	1.64	3.33E−18	0.3413
Total	8,126.97	8,249.87	8,940.64	0.8164
SFA	32,889.07	3,301.18	3,560.29	0.8603
MUFA	4,099.67	4,138.91	4,541.61	0.7737
PUFA	738.23	809.78	838.74	0.8157
Trans	364.82	345.86	418.67	0.5457

¹Amount (mg/100 g tissue) of fatty acid in powdered loin samples determined by gas chromatography.

²Control, saline injection; LPS50, 50 ng LPS/kg BW; LPS100, 100 ng LPS/kg BW.

³C16:1T, palmitoleic acid; C18:1T, elaidic acid; C18:1V, vaccenic acid; C18:2T, linolelaidic acid; C 18:3w3, α-linolenic acid; C 18:3w6, γ-linolenic acid; C 20:4w6, arachidonic acid; SFA, saturated fatty acids; MUFA, monounsaturated fatty acids; PUFA, polyunsaturated fatty acid.

pH

Data for pH across treatment and aging in lamb chops are exhibited in Table 3. There was no treatment effect for pH (P > 0.05), regardless of aging period. An aging effect was found for pH (P < 0.0001), as 14 day-aged chops had a greater pH (5.87) compared with 1 day-aged chops (5.71). However, the average difference in values are likely of little practical significance. There was no treatment-by-aging interaction (P = 0.7621).

Color (objective)

Color values are shown in Table 5. A days of aging-by-RD interaction was detected (P < 0.05) for objective color measures. In general, L* values increased and a* and b* values decreased as aging and RD increased, regardless of treatment.

Table 5.

The effects of LPS inclusion (control, LPS50 [50], LPS100 [100]) and aging (1 or 14 d) on objective (L*, a*, b*) and subjective percentage discoloration (%), and delta E values (n = 29)

Variable	Age	Treatment1				P-values
		Control	50	1,000	SEM	Trt	Age	Day	Trt*Age	Trt*Day	Age*Day	Trt*Age*Day
L* 2	1	44.37	45.47	45.82	0.64	0.002	0.68	<0.0001	0.92	0.99	<0.0001	0.99
	14	44.36	45.73	46.06
	$\overline{x}$	44.37b	45.6ab	45.97a
a* 2	1	13.7Ay	13.67Ay	13.31Ay	0.26	0.01	<0.0001	<0.0001	0.0008	0.96	0.05	0.18
	14	15Bx	16.01Ax	15.7Ax
b* 2	1	6.93C	8.2A	7.39BC	0.29	0.12	0.12	<0.0001	0.02	0.99	<0.0001	0.78
	14	7.88A	7.55A	8.04A
Percent discolorati-on	1	7.81A	3.34A	9.27A	3.70	0.35	0.22	<0.0001	0.02	0.99	0.48	0.74
	14	16.43A	3.32B	5.58B
ΔE3	1	6.01	4.46	6.18	1.42	0.19	0.74	n/a	0.07	n/a	n/a	n/a
	14	7.66	4.07	4.27

¹Control, saline injection; LPS50, 50 ng LPS/kg BW; LPS100, 100 ng LPS/kg BW.

²L*, black to white color space; 100 = light (white), 0 = dark (black); a*, red to green color space; + value (red), − value (green); b*, yellow to blue color space; + value = yellow, − value = blue.

³Larger delta (ΔE) values indicate a larger change in overall color over time; ΔE = [(ΔL* ²)+(Δa* ²)+ Δb* ²)]^0.5.

^a,b,cMeans within the same variable with different superscripts denote treatment differences (P < 0.05).

^x,yMeans within the same variable with different superscripts denote age differences (P < 0.05).

^A,B,CMeans within the same variable with different superscript denote Trt*age interactions (P < 0.05).

L* values were statistically different (P = 0.0017) among dietary treatments. Regardless of days of aging and RD, L* values were greater (lighter color) in LPS100 compared with control treatments (45.97 and 44.37, respectively). Chops from the LPS50 treatment was not different from the other treatments. There were no days of aging-by-treatment interaction for L* values (P > 0.05).

There was a days of aging by treatment effect (P = 0.0008) in a* (redness) values. Regardless of RD, chops from control lambs exhibited the lowest a* value across treatments at 14 days of aging, with no difference when comparing LPS50 and LPS100. There were no significant differences found for a* at 1 day of aging. An aging by-treatment effect (P = 0.02) was observed in b* values. Within 1 day of aging, meat from lambs treated with LPS50 exhibited (P < 0.05) the highest b* values compared with all other treatments. There were no significant differences at 14 days of aging. However, chops from lambs treated with LPS100 exhibited the highest b* values compared with all other treatments.

There were no significant differences (P > 0.05) in delta E values across treatments (Table 5). However, control chops exhibited numerically greater color change after 14 days aging, as shown by a larger delta E value, compared with LPS50 and LPS100 treated chops (7.66, 4.07, 4.27, respectively).

Discoloration

As expected, percent discoloration of chops increased (P < 0.0001) as days of RD increased (d 1–7 = 0.03, 0.13, 0.57, 1.67, 4.41, 9.95, 19.66, 27.64, respectively). A days of aging-by-treatment effect was shown in Table 5 (P = 0.02). Chops from control lambs clearly had the most discoloration across all treatments at 14 days of aging (P < 0.05). There were no differences among treatments at 1 day of aging.

Thiobarbaturic acid reactive substances

No differences (P > 0.05) in treatment were identified for TBARS values (Figure 2). As days of RD increased, lipid oxidation (mg malonaldehyde) also increased (P < 0.0001). A days of aging tendency was found, as 14 day-aged chops tended (P = 0.06) to have more malonaldehyde content compared with 1 day-aged chops (2.26 mg and 1.65 mg malonaldehyde, respectively). While not significant (P = 0.17), chops aged for 14 days with 7 days of RD (14–7) had the greatest TBARS values compared with all other age-by-RD combinations.

Figure 2.

Malonaldehyde content of lamb chops (L. lumborum) for 1- and 14-days aging from lambs administered control, LPS50, LPS100. Superscripts within the same day per aging period are different (P > 0.05; SEM control = 0.5491, SEM LPS50 = 0.5788, SEM LPS100 = 0.5491).

Discussion

LPS treatment and oxidative response mechanisms

This research explored the relationship between oxidative stress response via LPS exposure and the subsequent impacts of oxidative processes on meat quality. There is a wide variety of commercially available LPS preparations and the type of LPS used in an analysis dictates the degree of physiological stress induced. Therefore, effects due to LPS treatment and its impact on oxidative stress are rather complex (Suliman et al., 2004). LPS are covalently bound lipids derived from the outermost membrane of many Gram-negative bacterium (Escherichia coli). Acting as an endotoxin, LPS promotes an acute inflammatory response (Sternberg, 2007; Powers et al., 2011). As a result, innate immune response mechanisms are activated, such as nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB), which start transcription of pro-inflammatory cytokines interleukin-1β (IL-1), interleukin-6 (IL-6), and tumor necrosis factor-α (TNF-α; Halawa et al., 2013). Similar to hormones like cortisol, cytokines have broad functionality, including effects related to cellular signaling of homeostatic mechanisms during biological stress.

Given the complexity of oxidative stress in vivo, administration of LPS provided a dose-dependent method to quantitatively induce oxidative stress. The LPS model (E. coli O111:B4) promoted a quick inflammatory response in vivo, as indicated by the rapid change in rectal temperatures of lambs (Yates et al., 2011). Under conditions of high LPS concentration, pro-inflammatory mediators such as ROS are produced as a result of LPS toxicity, inducing oxidative stress and altering composition of muscle cells and apoptotic mechanisms (Suliman et al., 2004). While recent literature (Niu et al., 2016) has used LPS as a means to generate oxidative stress and influence meat quality in poultry, the impact of reintroducing LPS every 72 h in an immune challenge, as in the study presented here, has yet to be determined. We postulated that the constant reintroduction of an oxidative promoter would provide a basal level of oxidative stress prior to harvest, replicating the effects of events that could potentially promote oxidative promoters in livestock production systems (e.g., genetics, diet, environment, and handling) which may impact final meat quality.

The development of ROS can impact gene expression in muscle cells, promoting changes in cell proliferation, differentiation, function, and turnover. Using proteomic analysis, Malheiros et al. (2019) observed greater damage to proteins from oxidative stress in tender beef, compared with tough beef. The proteomic profile identified oxidation to antioxidant enzymes, heat shock proteins, and structural proteins, suggesting that biochemical changes within muscle induced by oxidative stress prior to harvest can impact meat tenderness. Oxidation of individual proteins was not assessed in this study. In the present study, transcriptomics was utilized to evaluate changes in gene transcripts among samples induced by a LPS challenge. Lambs administered LPS100 exhibited an upregulation of genes in stress response pathways compared with the control. This was evident by the predicted changes in pathways responsible for cell biosynthesis and turnover (ILK, Ceramide, IGF-1, PI3K, EGF, ERK5). These pathways are predominantly involved in cellular metabolism, proliferation, differentiation, protein transfer, and cell signaling. Interestingly, certain pathways act as hybrid models, both acting for cell biosynthesis but have secondary functions to inhibit apoptosis (PI3K). This is expected, as a biological system that has undergone biological stress would endure cellular damage, and thus need mechanisms to repair, grow, and reinforce new cellular structures, organelles, and signaling mechanisms. Conversely, in control samples, pathways predicted to be upregulated compared with LPS100 samples include biosynthesis pathways (cAMP, ErbB2-ErbB3), as homeostatic maintenance of cells requires continuous cellular turnover. It is noteworthy that genes upregulated in LPS100-treated lambs included those responsible for high levels of cellular growth and differentiation, suggesting they can account for increased cellular turnover as a result of acute biological disruption. Additionally, LPS-treated lambs predicted to upregulate pathways responsible for nucleic acid function. These proteins are responsible for the modification and development of chromosomes, chromatin function, and regulation of protein transcription, some of which are also capable of inducing cytokine signaling (JAK/Stat, HMGB1). Most notably, LPS-treatment resulted in a predicted upregulation of oxidative stress-related genes, all of which act for cellular oxidative response mechanisms (IL-6, IL-8, IL-3, NRF2, Chemokine), stress signaling (p38 MAPK, CXCR, TNFR-2, and Sumoylation), or autophagic initiation. Although overall statistical significance was just 0.05, the increased upregulation of systematic processes in response to LPS treatment in this study supports that oxidative stress was generated within our test subjects, allowing stress response systems to be evaluated.

Oxidative stress also changed mRNA expression related to muscular function and development. In particular, LPS-treated samples exhibited an upregulation of NO/ROS generation in macrophages, neuromuscular signaling between neurons and myofibrils (Agrin), and alterations in messenger molecules used to selectively block epithelial calcium channels (D-myo-inositol tetrakiphosphate biosynthesis pathways). This implies the induction of oxidative stress not only generated ROS, but also upregulated different pathways related to proteins which inhibit calcium regulation. From the perspective of muscle contraction and early postmortem tenderization, these factors hold tremendous implications on the development of muscle function by neuromuscular signaling in vivo in addition to enzymatic action in post-rigor muscle tissue. Interestingly, genes upregulated in control samples have the opposite impact on muscle tissue. Specifically, an increase in calcium signaling was found, suggesting that a homeostatic environment facilitates increased availability of calcium in myocytes. Additionally, increases in PPAR α/RXRα, responsible for glucose and lipid metabolism, were identified in control lambs. Lastly, eNOS signaling was upregulated in control samples (compared with LPS100 samples), reflecting support of endothelial homeostasis within the lining of blood vessels, in addition to maintenance of blood flow. Perhaps eNOS acts on mitochondrial biogenesis and fission within homeostatic environments (Tengan et al., 2012). This is plausible, as an increase in oxidative stress would induce mitophagy rather than mitochondrial fission. In total, these measures help to validate the presence of oxidative stress within our experiment, as well as identifying subsequent mechanisms which act in response to oxidative stress.

When discussing oxidative products, biomarkers like isoprostanes have recently been used as measures of downstream products of sustained oxidative stress (Montuschi et al., 2004). F₂-isoprostanes are the family of prostaglandin-like compounds formed by nonenzymatic, free-radical oxidation of arachidonic acid (20:4) by reactive oxygen species. Due to an integrated mechanism required to produce isoprostanes, its generation is contingent on ROS-mediated oxidation (Lawson et al., 1999; Milne et al., 2011). Generated during lipid peroxidation, F₂-isoprostanes are produced in the form of esterified fatty acids in phospholipid membranes, and then released using phospholipase action (Montuschi et al., 2004). Compared with other oxidative products, F₂-isoprostanes are very stable compounds that are detectable in all normal biological fluids and tissues. It is common to sample muscle biopsies or anoxic muscle tissue for isoprostane content, as the lack of blood present prevents F₂-isoprostanes from being transferred and metabolized in blood. Given the relationships between isoprostane generation and oxidative stress activity, detection of this molecule is regarded as the standard for evaluating sustained oxidative stress. Isoprostane has been linked to oxidative stress in animals due to its evaluation across a multitude of treatments speculated to influence oxidative stress in vivo, such as exercise (Karamouzis et al., 2004). Ponnampalam et al. (2017) proposed the change in isoprostanes as an in vivo indicator of meat quality deterioration postmortem (in vitro). Using blood samples from finishing lambs, they detected an increase of 8-isoprostane PGF₂α levels within plasma, with lambs fed a high energy feedlot diet exhibiting the greatest concentration of plasma 8-isoprostane PGF₂α at 4 and 8 weeks of feeding trials. This is due to both the increase in energy content of diets, but also the amount of α-tocopherol content increased in roughage-based diets (ryegrass and lucerne). Concurrently, positive relationships between 8-isoprostane PGF₂α levels and lipid oxidation (malonaldehyde content) were observed. Interestingly, positive relationships between 8-isoprostane PGF₂α levels, arachidonic (20:4), and linolenic (18:2n−6) acid were seen. This supports Milne et al. (2011) on how F₂-isoprostanes can potentially be generated from other PUFAs (eicosapentaenoic acid [20:5n−3], adrenic acid [22:4n−6], and docosahexaenoic acid [22:6n−3]) due to the orientation of their double bonds, but the meat quality impact of these fatty acids being able to generate F₂-isoprostanes is not fully understood. The generation of F₂-isoprostanes is an excellent marker for in vivo oxidative damage as a result of oxidative stress. We did not find a statistical difference between control and LPS-treated samples. However, it is noted that LPS50 and LPS100 treated samples had noticeably greater isoprostane content compared with the control (~44.7% and 32.9% greater content, respectively). In conjunction with changes in RNA transcripts for genetic pathways, this supports our hypothesis that the presence of increased in vivo oxidative stress occurred in LPS-treated samples.

Oxidative stress and meat tenderness

Tenderness is recognized as one of the most important factors for consumer palatability and repurchasing of meat (Koohmaraie and Geesink, 2006; Kemp et al., 2010). Subtle variations (~0.5 kg) in tenderness within muscles have been shown to greatly influence consumer likeness to different retail products (Martinez et al., 2017). While marbling acts as an indicator of tenderness (Emerson et al., 2013), a complete understanding of the biological mechanism of this relationship is not known. Such knowledge might be used to induce and increase the rate or extent of tenderization in meat. Oxidative stress has been linked to factors influencing meat quality, as oxidative stress has been suggested to impact organelles linked to normal muscle function and promote cellular turnover via apoptotic mechanisms (Stamler and Meissner, 2001; Kagan et al., 2009; Bolisetty and Jaimes, 2013). It has been postulated that oxidative stress alters proteins responsible for muscle structure, color stability, and organelle components interconnected with proteolytic mechanisms. As a result, oxidative stress could influence the degree of postmortem tenderization, and greatly impact both product quality and consumer palatability (Ott et al., 2007; Mitacek et al., 2019; Xing et al., 2019).

This study was not sufficient to claim LPS decreased WBSF. However, a tendency (P = 0.10) was shown that LPS impacts tenderness of Longissimus muscle early postmortem, as indicated by numerical differences in shear force at 1 day of aging. This tendency in early postmortem tenderization is reflected in the troponin-T degradation. Lonergan et al. (2001) indicated that degradation of troponin-T early postmortem (~2 days) could be used as a consistent indicator of proteolysis and muscle tenderness throughout muscle aging time up to 14 days aging. Oxidative stress catalyzes apoptotic events, possibly upregulating enzymatic degradation of proteins such as troponin-T (Sierra and Olivan, 2013). In the present study, a statistically significant (P = 0.02) increase in troponin-T degradation at 1 day postmortem, in comparison with the control, was found for LPS50 samples, but not for LPS100 samples. Additionally, chops from lambs treated with LPS50 had numerically greater degradation compared with control and LPS100 at both 1-day aging and 14-day aging. Our analysis suggests that early postmortem degradation of troponin-T can be impacted under certain conditions of increased oxidative stress, influencing shear force and overall tenderness of meat. While these data are promising when hypothesizing an oxidative stress—tenderness relationship, no differences were observed for desmin degradation or free calcium.

It appears oxidative stress induced by LPS50 was associated with lower shear force values compared with control and LPS100 treatments. The statistical differences in troponin-T degradation at 1 day aging would support this change in WBSF early postmortem. Using much higher doses of LPS (3–6 mg/kg) in poultry, Niu et al. (2016) found greater WBSF in 1 day-aged samples than controls. In addition, there were significant correlations of the oxidative stress biomarker isoprostane to early postmortem tenderness and proteolytic degradation in the study by Niu et al. (2016). Given this information, an argument can be made that different levels of LPS-mediated inflammation can induce different degrees of metabolic response across livestock species and therefore alter the impact on meat tenderness.

The influence of oxidative stress on meat is likely dependent on the time frame in which samples are exposed to an oxidative stress promoter. From the literature, tenderness has been examined from samples administered oxidative stress pre- and postmortem. Cook et al. (1998) and Wang et al. (2018) found lower shear force values when injecting pre-rigor Longissimus muscle with nitric oxide promoters and H₂O₂ solution, respectively. It can be postulated that the overwhelming increase in reactive species exhausted antioxidant systems, and initiated expression of pro-apoptotic mechanisms (Bax, Bcl-2) responsible for organelle instability and mitophagy (Wang et al., 2018). Concomitantly, the degradation of organelle stability promotes efflux of molecular ions bound within organelles (cytochrome c), which facilitates calcium-dependent enzymatic degradation of proteins. Cottrell et al. (2008) identified increased tenderness in Longissimus muscle when lambs were induced with nitric oxide inhibitors pre-slaughter. However, the same parameters resulted in higher shear force (less tender) values in Semitendinosus samples. Suster et al. (2005) reported increasing the time to inject nitric oxide inhibitors prior to slaughter caused lower shear forces in both Longissimus and Semitendinosus muscles. These values paralleled increased myofibrillar fragmentation, an indicator of tenderness/proteolysis. Both experiments examined specific reactive species in response to meat quality. Warner et al. (2005) postulated reactive nitrogen species (RNS), working in concert with reactive oxygen species (ROS), can generate highly radical compounds capable of disrupting proteins responsible for organelle function pertinent to postmortem tenderization.

A multitude of parameters were examined in this study which could influence oxidative stress. Factors such as metabolizable energy in animal diets, fatty acid composition, and structure of muscle fibers have all been implicated to impact meat tenderness, with evidence suggesting altering these factors by way of oxidative stress may accentuate changes in meat quality (Chauhan et al., 2016; Starkey et al., 2016; Ribeiro et al., 2019). To prevent confounding parameters on oxidative stress-influence meat quality, these factors were considered. No differences were found in muscle structure (sarcomere length), as similar results were found by Starkey et al. (2015) who reported postmortem sarcomere lengths of 1.77 μm in lamb longissimus muscle. Additionally, no statistical differences were found across fatty acid composition of muscle tissue. This is reasonable, given all lambs were fed the same ration throughout the study.

The results from this study suggest that LPS-mediated oxidative stress has the potential to increase early postmortem tenderness, which is likely due to the intricate relationship between the prolonged generation of oxidative stress and in vivo alterations among biological mechanisms responsible for muscle tenderization within the myofibril postmortem. We speculate the duration of oxidative stress in vivo and degree of oxidative stress induced by the concentration of oxidative stress promoter, as shown in this experiment, might be responsible for the impact of oxidative stress on meat tenderness. Additionally, the isolation and analysis of individual muscles, as well as any additional extrinsic conditions used in normal management strategies which could promote oxidative stress, such as diet, temperature, exercise, or genetic selection of livestock, could all influence tenderness via oxidative stress.

Oxidative stress and stability of color and lipids (pH, isoprostanes)

Given the amount of biological stressors produced during oxidative stress, it is critical to evaluate conditions which are detrimental to meat quality (Xing et al., 2019). Muscle pH was not greatly influenced by LPS treatment, suggesting our treatments did not generate sufficient stress to exhaust glycogen supplies and increase ultimate pH in meat. Additionally, neither lipid nor protein oxidation were affected by LPS treatment. Correlation coefficients (data not shown) reinforced the relationship of lipid oxidation and pH, with some indication that lipid oxidation can coincide with increased proteolysis (troponin T, calcium). These data, along with color values, indicate that LPS treatment was not detrimental toward color stability or lipid oxidation. With regards to our experiment, oxidative stress-induced samples tended to maintain color stability, exhibiting sustained redness (a*) and increased lightness (L*) in chops from LPS-treated lambs compared with control samples. This is supported by noticeably less discoloration in LPS-treated lamb chops after aging. Oxidative stress damage was not supported by lipid oxidation, as there were no noticeable changes in malonaldehyde content. Interestingly, a numerical increase in oxidative biomarker (8-isoprostanes) content was seen in the absence of lipid oxidative differences, suggesting that oxidative stress occurred to a greater extent in LPS-treated lambs, but not to a degree to promote detrimental effects on lipid oxidation. This observation conflicts with (Mitacek et al., 2019) who reported that prolonged aging promoted mitochondrial degeneration, depletion of color reducing enzymes, and decreased color stability in their study.

Conclusion

In conclusion, LPS-mediated oxidative stress triggered the onset of biochemical pathways responsible for muscle composition, proliferation, sustainability, and apoptotic mechanisms related to meat quality. Although there were few statistical differences in factors contributing to meat tenderization caused by LPS treatment, there was a trend that showed lambs subjected to LPS had lower shear force values early postmortem compared with the saline control group. This trend in shear force did not continue at 14 days of aging. These findings could be the result of greater proteolysis of troponin-T early postmortem in chops from lambs administered LPS50. There were no detriments to lipid oxidation and color stability.

Glossary

Abbreviations

BW

body weight

cAMP

cyclic adenosine monophosphate

ELISA

enzyme-linked immunosorbant assay

H2O2

hydrogen peroxide

IGF

insulin-like growth factor

LPS

lipopolysaccharide

MAPK

mitogen-activated protein kinase

MUFA

monounsaturated fatty acids

NO

nitric oxide

PUFA

polyunsaturated fatty acid

RNS

reactive nitrogen species

ROS

reactive oxygen species

SFA

saturated fatty acids

TBARS

thiobarbaturic acid reactive substances

WBSF

Warner–Bratzler shear force

Conflict of interest statement

The authors have no conflicts of interest to declare.