Exploring the Longissimus Muscle: Unraveling its Correlation with Meat Quality in Bos indicus and Crossbred Bulls

Summary

We investigated skeletal muscle tissue in Bos indicus and crossbred bulls to explain differences in meat quality traits. Warner-Bratzler shear force (WBSF) was found to range from 4.7 kg to 4.2 kg. Myosin heavy chain isoforms revealed differences between animals, and myofibril fragmentation index provided further insights into tenderness (WBSF) variations.

Abstract

This study investigated muscle tissue in Bos indicus and crossbred bulls to explain differences in meat quality traits. Carcass traits, meat quality parameters, and biochemical and molecular investigations of myofibrillar proteins are described. Methods for evaluating pH, intramuscular fat (IMF), meat color (L*, a*, b*), water losses, tenderness, and molecular biology assays have been outlined. Specific procedures detailing calibration, sample preparation, and data analysis for each method are described. These include techniques such as infrared spectroscopy for IMF content, objective tenderness assessment, and electrophoretic separation of MyHC isoforms.

Color parameters were highlighted as potential tools for predicting beef tenderness, a crucial quality trait influencing consumer decisions. The study employed the Warner-Bratzler shear force (WBSF) method, revealing values of 4.68 and 4.23 kg for Nellore and Angus-Nellore (P < 0.01), respectively. Total cooking losses and biochemical analyses, including myofibril fragmentation index (MFI), provided insights into tenderness variations. Muscle fiber types, particularly myosin heavy chain (MyHC) isoforms, were investigated, with a notable absence of MyHC-IIb isoform in the studied Zebu animals. The relationship between MyHC-I and meat tenderness revealed divergent findings in the literature, highlighting the complexity of this association. Overall, the study provides comprehensive insights into the factors influencing meat quality in Bos indicus and crossbred (Bos taurus × Bos indicus) bulls, offering valuable information for the beef industry.

Introduction

Brazil has the largest commercial cattle herd globally, numbering approximately 220 million animals and ranking as the second-largest meat producer, yielding over 9 million metric tons of carcass equivalent annually¹. The beef cattle production sector significantly contributes to the national agricultural system, with total annual sales surpassing R$ 55 billion. Since 2004, Brazil has been a key player in the global meat trade, exporting to over 180 countries, which represents ~50% of the world meat trade².

Meat tenderness stands out as the paramount quality attribute influencing consumer satisfaction and meat consumption³. By employing biochemical and objective methods to measure meat tenderness, researchers aim to provide valuable insights into factors such as animal genetics, processing techniques, and storage conditions, ultimately enhancing the quality and consistency of meat products for consumers. Such information is useful because meat tenderness has gained increased importance in consumer decision-making during purchases. Moreover, meat tenderness assessment provides valuable information for quality control in meat production and processing industries. By consistently monitoring tenderness, producers can ensure that meat products meet desired standards and specifications. In this context, Brazilian beef cattle producers are progressively embracing intensive feedlot systems with crossbred animals to enhance capital turnover. This system accounts for about 10% tons of carcass produced annually in Brazil^4,5.

The escalating demand for improved meat quality by consumers has prompted beef cattle producers to crossbreed with European breeds, primarily Aberdeen Angus⁶. This strategy aims to produce F1 Angus-Nellore hybrids, known for superior performance, desirable carcass traits, and enhanced meat quality compared to pure zebu animals^7,8. In the tropical regions of Brazil, it is common practice to utilize non-castrated animals (bulls) of advanced maturity in finishing farms, potentially compromising meat quality attributes such as color, marbling, and tenderness. Notably, a survey reveals that 95% of animals finished in Brazilian feedlots are males, with 73% being Nellore, followed by 22% crossbred animals and 5% other genotypes^9,10.

Understanding the biochemical mechanisms underlying meat tenderness is crucial for improving meat quality. One key aspect is postmortem proteolysis, which affects the structural integrity of muscle fibers¹¹. The myofibril fragmentation index (MFI) is a widely used biochemical assay that quantifies the extent of myofibril degradation, providing insights into the tenderness of meat¹². The MFI method involves measuring the fragmentation of myofibrillar proteins, which directly correlates with meat tenderness. This assay complements traditional meat quality assessments and offers a deeper understanding of the biochemical processes that contribute to variations in meat tenderness.

In this context, the present study investigated the skeletal muscle of Bos indicus versus crossbred (Bos taurus × Bos indicus) bulls finished in feedlot, aiming to explain differences in meat quality traits.

Protocol

All procedures with animals complied with the ethical research standards established by the Animal Use Ethics Committee (CEUA) of the "Universidade Estadual Paulista Júlio de Mesquita Filho" - UNESP Botucatu Campus, under protocol 0171/2018.

1. Experimental animals

1. Finish 30 Nellore bulls (Bos indicus) and 30 F1 Angus-Nellore bulls (Bos taurus × Bos indicus), aged 20-24 months, in a feedlot. House both groups of animals in collective pens measuring 5 m x 6 m with a concrete floor and equipped with shell-type water troughs, accommodating up to five animals per pen. Ensure that all animals belong to the same management group (born and raised on the same farm) and are submitted to the same feedlot period.
   NOTE: In this study, the Nellore bulls had an average initial body weight of 370.7 kg while the F1 Angus-Nellore bulls had an average initial body weight of 380.8 ± 17 kg.
2. Feedlot diet
   1. Ensure that the finishing diet is comprised of 11.3% roughage (Tifton hay and sugarcane bagasse) and 88.7% concentrates (ground dry corn grain, soybean meal, corn wet distillers' grains, dry corn gluten feed, and mineral core). Feed the animals for 120 days and provide the diets ad libitum twice a day (at 10:00 a.m. and 04:00 p.m.).
3. Slaughter
   1. Register the final body weight (BWf) at the end of the experimental period. Process all animals at a nearby slaughterhouse, adhering to standard inspection procedures. Prior to slaughter, ensure that the animals undergo a minimum 16 h fast, abstaining from both feed and water.
      NOTE: F1 Angus-Nellore bulls exhibited a final body weight of 615.09 ± 57.53 kg, while Nellore bulls had a weight of 545.47 ± 11.45 kg.
4. Evaluation of carcass traits
   1. Weigh the beef carcasses initially and then subject them to a cooling period at 2-4 °C for 48 h. Measurements include hot carcass weight (HCW), rib eye area (REA), and backfat thickness (BFT) at the 12^th/13^th rib interface, as recommended¹³. Determine the REA using the grid method with a small grid (18 cm x 13 cm) and measure the BFT in millimeters using a caliper.
      1. Measure the REA in each carcass by using a reticulated grid (the same as used in the USDA Yield Grade classification system), divided into 1 cm² squares with one dot in the middle. Add all squares within the ribeye tracing perimeter and those along the contour of the tracing passing through the middle dot.
      2. Measure the BFT at a specific position on the assessment site anywhere between the 12^th/13^th ribs. To determine this position, measure the length of the rib eye; then, starting at the medial border "A", determine a point three quarters of the way along the rib eye and halfway across "B". Take a caliper through this point and at right angles to the specified rib to the interface between the subcutaneous fat and intermuscular fat. Measure the subcutaneous fat by placing the caliper at a right angle to the line of the subcutaneous fat, from the interface point (Supplemental Figure S1).
5. Sampling
   1. Sample Longissimus thoracis (LT) from the left half-carcass (portion of ± 12.0 cm of meat), between the 11^th and 13^th ribs in the cranial direction. In the laboratory, section the meat samples into steaks of 2.54 cm.
6. Aging
   1. Assess the meat quality traits after a 14 day wet-aging period at a temperature of 0-2 °C in a biological oxygen demand (BOD) incubator. Use steaks of 2.54 cm thickness for the analysis of meat color, pH, intramuscular fat, purge loss, water-holding capacity, objective tenderness, and cooking losses. Pack the steaks separately in plastic bags for high vacuum and low oxygen permeability and after the aging time is reached, keep them frozen at -20 °C until the time of analysis. Thaw the beef samples at 4 °C for 24 h and expose them to oxygen for 30 min at 4 °C (blooming time).

2. Meat pH

1. Measure the pH of the meat using a digital pH gauge equipped with a penetration probe. Calibrate it with pH 4.0 and 7.0 buffers at a room temperature of 25 °C. Measure the meat pH at three locations of the LT muscle sample. Manually record the data readings and subsequently export the datasheet; calculate the average of the three readings for meat pH.

3. Intramuscular fat

NOTE: Intramuscular fat (IMF) content was determined using near infrared (NIR) spectroscopy¹⁴ and by gravimetric method¹⁵.

1. Remove the subcutaneous fat from the LT muscle using a scalpel. Then, grind and homogenize the steak for 5 min using a mixer, incorporating approximately 180 g of the sample. Place the sample in a cup, position it inside the sample chamber, and perform subscanning of various zones of the test sample by rotating the sample cup; merge the zones for the final result.
2. Take three readings for each sample. After homogenization, place the samples in the plate for subsequent reading. Set the apparatus to NIR transmission, with a moving grating monochromator scanning the region from 850 nm to 1050 nm.
3. Export the data sheet and then calculate the average three reading for the IMF. Express the results as a percentage, using the formula: [(IMF average ÷ sample weight) × 100].
4. Combine homogenize the LT muscle samples (3.0 g) with a chloroform/methanol methanol/chloroform (2:1) solution for 2 min and subject them to centrifugation (700 × g; 10 min; 20 °C) to segregate the hydrophilic (upper), solid (middle), and hydrophobic (lower) phases.
5. Filter the hydrophobic phase obtained post centrifugation using a filter paper on a funnel with slight suction. Transfer the filtrate (bottom phase; lipids in chloroform) to a flask labeled as lipid phase and transfer at least 5 mm of filtrate to a preweighed beaker flask after letting it stand for a few minutes. Then, record the volume of the chloroform layer (at least 150 mL) and aspirate the alcoholic layer.
   1. Homogenize 100 g aliquots of the sample of fresh or frozen tissue for 2 min with a mixture of 100 mL of chloroform and 200 mL of methanol. Add 100 mL of chloroform to the mixture, blend for 30 s, add 100 mL of distilled water, and blend for another 30 s.
   2. Filter the homogenate through filter paper on a funnel with slight suction. Apply pressure with the bottom of a beaker when the residue becomes dry to ensure maximum recovery of solvent.
   3. Transfer the filtrate to a 500 mL graduated cylinder and let it stand for a few minutes to allow separation and clarification. Record the volume of the chloroform layer (at least 150 mL) and aspirate the alcoholic layer.
   4. Be sure to completely remove the top layer; the chloroform layer contains the purified lipid. For quantitative lipid extraction, recover the lipid trapped in the tissue residue by blending the residue and filter paper with 100 mL of chloroform.
   5. Filter the mixture through the funnel and rinse the blender jar and residue with a total of 50 mL of chloroform. Mix this filtrate with the original filtrate before removing the alcoholic layer.
      NOTE: Filtration is typically rapid; apply pressure with the bottom of a beaker on the dry residue to ensure maximum recovery of solvent.
6. Dry the samples in an oven, cool them in a desiccator for at least 24 h, place them in an oven at 110 °C until complete solvent evaporation, further cool them in a desiccator overnight, and finally reweigh them.
7. Determine the IMF content by calculating the difference between the initial and final weights of the beaker.

4. Meat color

1. Calibrate the device using a black and a white standard plate. Place the white calibration plate near the middle of the plate. When doing a calibration, use the area near the middle of the plate. Calibration is complete after the lamp flashes three times.
2. Take measurements after 30 min at 4 °C (blooming time). Obtain color readings from three different locations on the LT muscle sample, carefully avoiding connective tissue and fat.
3. At room temperature (20 °C), compute an average from these measurements, as recommended¹⁶.

5. Water losses

1. Assess purge loss (PL) for all samples. Determine the PL of beef loin sections by measuring the variance between the initial weight before freezing and the final weight after freezing/thawing.
   NOTE: Do not evaluate the PL of never-frozen control beef loins.
2. Gauge the water-holding capacity (WHC) by the weight difference of a meat sample (approximately 1.0 g) before and after being subjected to a pressure of 10 kg for 5 min¹⁷.

6. Objective meat tenderness

NOTE: The measurement of Warner-Bratzler shear force (WBSF) was conducted as described^18,19.

1. Position samples on a grid attached to a glass refractory and cook them in an industrial electric oven until reaching a final temperature of 71 °C. After cooking, cool, weigh, and refrigerate the samples at 4 °C for 24 h.
2. Determine cooking losses (CL) using the formula .
   1. Determine drip loss by weighing the refractory before and after cooking the sample. To this end, place the samples on a grid over a glass refractory to allow drainage of meat juices and fat during cooking.
   2. Determine evaporation loss by weighing only the sample before and after cooking.
   3. Record raw and cooked weights and calculate the percentage of DL as the weight of drip after cooking divided by the weight of the thawed meat sample.
   4. Calculate the evaporation loss percentage (EVP) using the formula [100 - (weight after cooking) ÷ raw weight × 100].
3. For the determination of WBSF, section eight cores with a diameter of 1.27 cm using a texture analyzer, equipped with a 3.07 mm Warner-Bratzler Shear Force blade and a V-shaped (60° angle) cutting edge.
   1. Report the results as the average of six values per sample, in kilograms (kg), after excluding the low and high extremes¹⁹.

7. Biochemical assay

NOTE: Postmortem proteolysis was assessed by estimating the myofibril fragmentation index (MFI), following the original procedure outlined by Culler et al.²⁰ and adapted for Bos indicus cattle by Borges et al.²¹.

1. Homogenize fragments of approximately 3 g of LT samples (fat-stripped muscle tissue and connective tissue) in a buffer solution containing 100 mM potassium chloride, 20 mM potassium phosphate at pH 7, 1 mM ethylenediaminetetraacetic acid (EDTA), 1 mM magnesium chloride, and 1 mM sodium azide at 2 °C, followed by centrifugation (1,000 × g for 15 min at 4 °C).
   1. Resuspend the sediment in 10 volumes (v/w) of isolating medium using a stir rod, then sediment it again at 1,000 × g for 15 min and decant the supernatant.
   2. Resuspend the sediment in 2.5 volumes (v/w) of isolating medium and separate the connective tissue and debris by passing it through a polyethylene strainer (18 mesh). Use an additional 2.5 volumes (v/w) to allow the myofibrils to pass through the strainer.
   3. Determine the protein concentration of the suspension of myofibrils using the biuret method of Gornall et al.²². Dilute an aliquot of the myofibril suspension with isolating medium to a protein concentration of 0.5 ± 0.05 mg/mL.
   4. Immediately measure the absorbance of this suspension at 540 nm. Determine MFI using spectrophotometry at 540 nm. Multiply the absorbance by 200 to obtain the MFI for each sample (and report it as an index without a measurement unit).

8. Molecular biology assay

NOTE: For the analysis of myosin heavy chain (MyHC), the most abundant protein in bovine skeletal muscle, LT samples from both groups were processed following the protocol described in the literature^23,24.

1. Achieve electrophoretic separation using a gradient SDS-PAGE gel (7-10%) and a 4% stacking gel. Apply 25 µL of each sample to the gel and run it at 70 V, 28 mA, and 4 °C for 1 h, followed by a run at 180 V, 12 mA, and 4 °C for 29 h.
2. Employ two different buffers in the runs: the upper gel buffer comprising glycine, Tris(hydroxymethyl)aminomethane base, sodium dodecyl sulfate (SDS), and distilled water, while the lower gel buffer is identical to the upper buffer, with the addition of mercaptoethanol.
3. Stain the gels with Coomassie Blue and capture images using appropriate software.
4. Identify MyHC isoforms (MyHC-I, MyHC-IIa, MyHC-IIx/d) based on their molecular weights (223.900, 224.243, and 223.875 kDa, respectively). Conduct semi-quantitative analysis by densitometry of the bands corresponding to each isoform, utilizing appropriate software.
5. Employ Rat soleus and Extensor digitorum longus (EDL) muscle as positive controls to classify MyHC isoforms, reserving one well in each gel for loading 40 µL of the processed sample.
6. For all data, perform the analysis of variance (ANOVA) by the F test, using the following model:
   Y_ij = µ + t_i + Ɛ_ij
   where Yij is the observed value of the experimental unit referring to treatment i in repetition j; µ is the general effect of the mean; t is the treatment effect (genetic group), and ε is the experimental error.
7. Compare the means by using the Student's t-test and adopt P-value < 0.05 as the critical probability.

Results

Table 1 displays the carcass traits of the two genetic groups investigated in this study. Notably, differences were identified (P < 0.01) in HCW, REA, and BFT, with crossbred animals exhibiting greater values, suggesting a heterosis effect.

Variable¹	Nellore	F1 Angus x Nellore	SEM	P-value

Discussion

During carcass evaluation, it is crucial to accurately measure growth and quality traits following a 48 h cooling period to obtain consistent and comparable data. The two biological models exhibited divergent carcass traits, particularly HCW, REA, and BFT, which are consistent with findings reported in other studies. The average HCW of Nellore bulls aligns with Brazilian market preferences, which prioritize greater meat production per animal unit with less fat content²⁵. Conversely, crossbred catt...

Materials

Name	Company	Catalog Number	Comments
Acetone	Merk, Darmstadt, Germany	CAS 67-64-1 | 100014	solutions used for the electrophoretic separations
Anti-MYH-1 Antibody	Merk, Darmstadt, Germany	MABT846	Rat soleus
Anti-Myosin antibody	Abcam, Massachusetts, United States	ab37484	Myosin heavy chain
Anti-Myosin-2 (MYH2) Antibody	Merk, Darmstadt, Germany	MABT840	Extensor digitorum longus (EDL)
Biological oxygen demand (BOD) incubator	TECNAL, São Paulo, Brazil	TE-371/240L	Meat aging
Chloroform; absolute analytical reagent	Sigma-Aldrich, Missouri, United States	67-66-3	Intramuscular fat
CIELab system	Konica Minolta Sensing, Tokyo, Japan	CR-400 colorimeter	Meat color
Coomassie Blue	Sigma-Aldrich, Missouri, United States	C.I. 42655)	Myosin heavy chain
Electric oven	Venâncio Aires, Rio Grande do Sul, Brazil		Meat tenderness
Ethanol	Merk, Darmstadt, Germany	64-17-5	solutions used for the electrophoretic separations
Ethylenediaminetetraacetic acid	Sigma-Aldrich, Missouri, United States	60-00-4	Post-mortem proteolysis
Glass flasks	Sigma-Aldrich, Missouri, United States		solutions used for the electrophoretic separations
Glycine	Sigma-Aldrich, Missouri, United States	G6761	Myosin heavy chain
Infrared spectroscopy - FoodScan	Foss NIRSystems, Madson, United States	FoodScan™ 2	Intramuscular fat
Magnesium chloride	Sigma-Aldrich, Missouri, United States	7786-30-3	Post-mortem proteolysis
Mercaptoetanol	Sigma-Aldrich, Missouri, United States	M6250	Myosin heavy chain
Methanol, absolute analytical reagent	Sigma-Aldrich, Missouri, United States	67-56-1	Intramuscular fat
pH meter	LineLab, São Paulo, Brazil	AKLA 71980	Meat pH
PlusOne 2-D Quant Kit	GE Healthcare Product	Code 80-6483-56	Post-mortem proteolysis
Polypropylene	Sigma-Aldrich, Missouri, United States		solutions used for the electrophoretic separations
Potassium chloride	Sigma-Aldrich, Missouri, United States	7447-40-7	Post-mortem proteolysis
Potassium phosphate	Sigma-Aldrich, Missouri, United States	P0662	Post-mortem proteolysis
R software	Vienna, Austria	version 3.6.2	Data analysis
Sodium azide	Sigma-Aldrich, Missouri, United States	26628-22-8	Post-mortem proteolysis
Sodium dodecyl sulfate (SDS)	Sigma-Aldrich, Missouri, United States	822050	Myosin heavy chain
Spectrophotometer	Perkin Elmer, Shelton, United States	Perkin Elmer Lambda 25 UV/Vis	Post-mortem proteolysis
Statistical Analysis System	SAS, Cary, North Carolina, United States	version 9.1,	Data analysis
Texture Analyzer	AMETEK Brookfield, Massachusetts, United States	CTX	Meat tenderness
Tris(hydroxymethyl)aminomethane	Sigma-Aldrich, Missouri, United States	77-86-1	Myosin heavy chain
Ultrafreezer	Indrel Scientific, Londrina, Paraná, Brazil.	INDREL IULT 335 D - LCD	Sample storage
Ultrapure water	Elga PURELAB Ultra Ionic system		solutions used for the electrophoretic separations
Ultra-Turrax high shear mixer	Marconi – MA102/E, Piracicaba, São Paulo, Brazil		Post-mortem proteolysis