A total of 15 barrows of similar genetics (maternal-line nonselect offspring comprised of Yorkshire and Landrace breeds) were procured from the university research farm and slaughtered under federal inspection at The Ohio State University Meat Science Teaching Laboratory with approved humane slaughter procedures using single-file electrical stunning. The pigs were slaughtered in 2 slaughter events (i.e., blocks) at an average live weight (± SD) of 122.47 kg (± 3.79 kg). The first slaughter event consisted of 8 barrows, whereas the second slaughter event consisted of 7 barrows. Backfat thickness was measured at the last rib (midline) location. Carcasses weighed an average weight of 93.23 kg (± 3.02 kg) and measured with an average last rib backfat thickness of 3.88 cm (± 0.55 cm).

The pH and temperature of the LT, PM, ST, TB, and GM muscles from the left side of each carcass were measured at 1-h, 3-h, 6-h, 9-h, 12-h, and 24-h postmortem during the conventional chilling process. An approximately 15 cm × 15 cm area of skin (approximately 3–5 cm thick) on the exterior surface of the carcass was partially incised and reflected like a curtain using a clean, sharp knife to expose the ST, GM, LT, and TB muscles. This approach allowed for repeated pH and temperature measurements to be collected throughout the sampling period without substantially affecting the temperature decline of each muscle, as the skin flap could cover the area between measurements. The LT was measured at the 10th rib location of the carcass, while the PM was measured on the ventral muscle below the transverse process. The ST was measured on the posterior face of the ham. The TB was measured at the triangular area bound by the ventral edge of the scapula and the posterior edge of the humerus. The GM was measured between the ham and the loin, at the lateral face of the ilium bone. pH was obtained by inserting a glass-tipped probe attached to a portable pH meter into the muscle at a new location within the exposed 15 cm × 15 cm area for each measurement (Meat Probes Inc., Topeka, Kansas, USA). The pH meter was calibrated with pH = 4.01 and pH = 7.00 buffer solutions that were stored at refrigerated temperatures (≤4°C) prior to use. Temperature was measured using a digital meat thermometer probe (Taylor USA, Oak Brook, Illinois, USA). At each time point when pH and temperature measurements were conducted (1-h, 3-h, 6-h, 9-h, 12-h, and 24-h postmortem), approximately 100 g samples from the LT, PM, ST, TB, and GM muscles were collected using a scalpel blade, placed in a sterile 15 mL plastic centrifuge tube, and then immediately placed in liquid nitrogen for flash freezing before further storage in a −80°C freezer.

The LT, PM, ST, TB, and GM muscles were removed from the carcasses at 24-h postmortem. The thoracis portion (anterior to last rib) of the LTL muscle was utilized, while the entire PM, ST, TB, and GM muscles were used. After trimming subcutaneous fat and connective tissue, the muscle samples from the right side of the carcasses were vacuum packaged and allotted to 10 d of postmortem aging at 4°C and then frozen at −20°C until use for proximate analysis and sensory evaluation. The muscle samples from the left side of the carcasses were divided into seven 2.54-cm thick sections. One of the muscle sections was allotted to fresh meat quality analysis at 1-d postmortem. The remaining 6 muscle sections were vacuum packaged, and randomly allocated to 1 of the 6 treatments, which consisted of a 3 × 2 factorial design with 3 postmortem aging time points (1-d, 3-d, and 10-d) and 2 endpoint cooking temperatures (63°C or 71°C). After each postmortem aging period elapsed, the muscles were frozen at −20°C until the Warner-Bratzler shear force (WBSF) analysis took place. The remaining samples from the left side of the carcasses were cut into 3 small cuts (approximately 100 g each) and were then placed in 3 sterile 50 mL plastic centrifuge tubes, which were assigned to storage for either 1-d, 3-d, or 10-d postmortem at 4°C, respectively. After each of the respective storage days, the samples were immediately placed in liquid nitrogen for flash freezing before further storage in a −80°C freezer for further proteomic analysis.

Samples (used for the quality analysis at 1-d postmortem) were allowed to bloom for a period of 30 min prior to instrumental color evaluation. Instrumental Commission Internationale de l’Éclairage (L*, a*, b*) color values were evaluated using a handheld colorimeter with D₆₅ light source, an 11 mm aperture, and a 10° observer (Minolta CM-700d; Minolta Corp., Osaka, Japan) on the cut surface of each sample following carcass fabrication at 24-h postmortem. Drip loss was assessed using the EZ cup method described previously by Rasmussen and Andersson (1996). A 25 mm diameter core from each sample was weighed at 24-h and 72-h postmortem after being placed within specialized EZ drip loss containers (Sarstedt product no. 86.290; Sarstedt Ag & Co., Nümbrecht, Germany) to quantify drip loss over a 48 h period at the storage temperature of 4°C.

Approximately 100 g samples (from right-side carcasses) had their subcutaneous fat removed and were individually homogenized using a food processor (Hamilton Beach model 70760, type FP19; Hamilton Beach Brands, Glen Allen, Virginia, USA). Moisture was measured using the air oven drying procedure described by AOAC method 990.19, while intramuscular fat (IMF) was measured using the Soxhlet extraction technique described by AOAC method 960.39 (AOAC International, 2016). Duplicate homogenized pork samples (7 g each) were placed on aluminum weighing pans with 2 layers of #1 filter paper, weighed, and subsequently dried in a drying oven (Thermo Scientific Isotemp 100 L Forced Air Oven; Thermo Fisher Scientific, Waltham, Massachusetts, USA) at 100°C for 24 h. After drying, the samples were cooled in a vacuum-sealed desiccator for 15 min before being reweighed to determine the moisture content. Aluminum weighing pans containing samples were washed multiple times with warm chloroform methanol (mixed at a ratio of 87:13) in the Soxhlet apparatus for 9 h and dried in a drying oven at 100°C for 24 h before being reweighed to determine the IMF. Results from duplicate samples with a coefficient of variance less than 15% were averaged and reported.

Pork samples were removed from the freezer and placed in a single layer at an ambient temperature of 2°C to thaw for 24 h prior to WBSF analysis. Samples were assigned to endpoint cooking temperatures of either 63°C or 71°C. Samples were cooked for 35 min using the sous-vide technique in a precision water bath (Thermo Scientific Precision GP20; Thermo Fisher Scientific) that was set to either 63.5°C or 71.5°C, respectively. The water bath was set 0.5°C above the target endpoint temperatures to ensure that the samples reliably reached 63°C and 71°C. Internal cooking temperature was monitored throughout cooking by inserting a digital temperature logger with 2 type K thermocouple wires connected to a handheld digital scanning thermometer (ThermoQ; ThermoWorks, American Fork, Utah, USA) into the geometric center of a nonstudy pork LT sample cut to similar dimensions to study samples. Once the internal temperature reached the targeted endpoint cooking temperature, samples were immediately moved to a refrigerated room (ambient temperature of 2°C) and placed on a drying rack in a single layer. After 12 h of refrigerated storage, cooked samples were removed from vacuum packages and reweighed. The percentage of weight difference between initial raw weight and cooked weight was calculated as cooking loss. Six 1.25 cm diameter cores cut parallel to the muscle fibers were obtained from each sample using a handheld coring device. Cores were sheared perpendicular to the direction of muscle fibers with a Warner-Bratzler shear attachment equipped to a TA-XT Plus 100 Connect texture analyzer (Texture Technologies Corp., Hamilton, Massachusetts, USA) at a crosshead speed of 2 mm/s. The peak force (kg) required to shear through each core was recorded, and the average peak force of 6 cores was calculated for each sample.

Frozen pork samples were homogenized and powdered in liquid nitrogen. Protein extraction was conducted following the procedure of Richardson et al. (2018) with a few modifications. Powdered frozen samples (approximately 0.15 g) were homogenized with 1 mL of whole muscle buffer (2% sodium dodecyl sulfate wt/vol and 10mM sodium phosphate, pH 7.0) in a bead mill homogenizer (Bead Mill 4 Homogenizer; Thermo Fisher Scientific) at the #2 speed setting for 200 s. The homogenate was then centrifuged at 3220 × g for 15 min. Protein concentration of the supernatant was determined using a bicinchoninic acid protein assay kit (Pierce Protein Research Products, Rockford, Illinois, USA). Samples were adjusted to a final protein concentration of 4.0 mg/mL by mixing 0.03 mL of beta mercaptoethanol, 0.25 mL of Laemmli sodium dodecyl sulfate sample buffer tracking dye, and whole muscle buffer. Samples were then vortexed and heated on a heating block for 15 min at 50°C, followed by frozen storage at −80°C until further analysis occurred.

Sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis was used for running load checks to ensure proper protein concentrations were achieved. Protein samples (40 μg per well) were loaded into 4–12% Bis-Tris (bis[2-hydroxyethyl]amino-tris[hydroxymethyl]methane) gels (Invitrogen Bolt 4–12% Bis-Tris PlusGels, 10-well; Thermo Fisher Scientific) for desmin and troponin-T analysis. Calpain-1 analysis was conducted using 8% Bis-Tris gels (Invitrogen Bolt 8% Bis-Tris PlusGels, 10-well; Thermo Fisher Scientific). Gels were electrophoresed using 2-morpholinoethanesulphonic acid SDS running buffer and were then transferred to a polyvinylidene fluoride blotting membrane using the iBlot 2 western blotting workflow system. Membranes were blocked using 5% nonfat dry milk in phosphate buffered saline (PBS)-tween solution for 1 h at room temperature and washed 3 times for 10 min each with PBS-tween solution before transferring to the iBind flex western device (Thermo Fisher Scientific). The membranes were incubated (1 target protein at a time) with the following primary antibodies, which were diluted with iBind flex solution (Thermo Fisher Scientific): 1:2000 mouse monoclonal anti-desmin immunoglobulin G ([IgG]; D1022; Sigma-Aldrich, St. Louis, Missouri, USA), 1:2000 mouse monoclonal anti-troponin-T IgG (T6277, Sigma-Aldrich), 1:1000 mouse monoclonal anti-calpain-1 IgG (MA3–940; Thermo Fisher Scientific), to determine desmin (and its degradation products), troponin-T (and its degradation products), and calpain-1 autolysis. The secondary antibody used for each of the membranes was goat-anti-mouse IgG, horseradish–peroxidase-linked antibody (A28177; Thermo Fisher Scientific), which was diluted with iBind flex solution (Thermo Fisher Scientific) to the same dilution as primary antibodies, respectively. Blotting membranes were then washed with PBS-tween solution 3 times before protein detection. Protein bands were detected using a western blotting chemiluminescent detection kit (Pierce ECL western blotting substrate, Thermo Fisher Scientific). Immunoreactive bands were visualized using an iBright1500 imager (Thermo Fisher Scientific), and the area-defined density of each band was determined with iBright Analysis Software (Thermo Fisher Scientific). A consistent internal reference sample was loaded on each gel to standardize the intensity of protein bands. The intensity of targeted protein bands was quantified as a fold-change difference relative to the reference sample on the same gel. The reference sample used for desmin and troponin-T was a nonstudy LT muscle sample aged for 14-d postmortem, while the reference sample used for calpain-1 was a 1-h postmortem LT muscle sample. For desmin and troponin-T, the intensities of 55 kDa intact desmin, 42 kDa and 38 kDa degraded desmin, 37 kDa intact troponin-T, and 30 kDa and 28 kDa degraded troponin-T bands were quantified as a comparative ratio of the sample band to the corresponding reference protein band on each gel (Carlson et al., 2017) (Figure 1). Calpain-1 autolysis was determined as the percentage of the 80 kDa, 78 kDa, or 76 kDa bands within each sample (Ma and Kim, 2020) (Figure 2). All the samples were analyzed in duplicate across different gels, with a coefficient of variance less than 20% to ensure repeatable results and the average results from the 2 gels were reported.

The 5 muscle cuts from the right side of each pork carcass were subjected to 10 d of postmortem aging and cooked to endpoint cooking temperatures of either 63°C or 71°C to evaluate the influence of muscle and endpoint cooking temperature on sensory traits. A total of 11 panelists were trained to evaluate pork samples for tenderness, juiciness, chewiness, and pork flavor. Tenderness and chewiness training consisted of cooking a pork PM muscle to an internal endpoint cooking temperature of 63°C and a pork LT muscle to an endpoint cooking temperature of 71°C. Juiciness training consisted of cooking several LT muscles to different endpoint cooking temperatures including 63°C, 71°C, and 85°C. Flavor training consisted of cooking a pork ST muscle to an endpoint cooking temperature of 63°C and a LT muscle to an endpoint cooking temperature of 71°C. Additionally, Knorr brand pork bouillon cubes (Unilever, Englewood Cliffs, New Jersey, USA) were diluted to 4 different concentrations (6 mg/mL, 12 mg/mL, 24 mg/mL, and 48 mg/mL) for panelists to evaluate differences for pork flavor. Over the course of 2 training sessions, the panelists collectively established anchors and thresholds for tenderness, chewiness, juiciness, and pork flavor.

A total subsample of 38 muscle cuts cooked to 2 different endpoint cooking temperatures were used for the sensory testing (N = 76 samples; 5 muscle cuts from 6 different carcasses and 4 muscle cuts from 2 different carcasses). Samples were removed from −20°C storage and placed in a single layer on a plastic tray in a refrigerated room at 2°C for 24 h for proper thawing before each session. The samples were cooked to an endpoint cooking temperature of either 63°C or 71°C in a water bath using sous-vide cookers (Anova precision cooker; Anova Culinary, San Francisco, California, USA), then cut into 1.27 cm cubes (1.27 cm × 1.27 cm × 1.27 cm). The 76 samples were evaluated over 13 sensory sessions. Each sensory session consisted of no more than 6 pork samples, representing differences in muscles and endpoint cooking temperatures. Samples were assigned to sensory sessions using a balanced, incomplete block design, allowing every muscle and endpoint cooking temperature combination to be compared an equal number of times. Prepared samples were served to 6 predetermined trained panelists from the pool of 11 trained panelists in a randomized order during each sensory session. Panelists were provided with a pair of red shaded glasses to mask color differences between samples, along with a writing utensil, napkin, water, apple juice, and unsalted crackers. Tenderness, juiciness, chewiness, and pork flavor for each sample were assessed on 15 cm continuous-line scales, with anchors at 0, 7.5, and 15, where 0 was labeled as extremely tough, extremely dry, not chewy, or no pork flavor, and 15 was labeled as extremely tender, extremely juicy, extremely chewy, or very flavorful. Results from all panelists were averaged with coefficient of variation for tenderness, juiciness, chewiness, and pork flavor of 22.22%, 30.27%, 34.60%, and 14.18%, respectively. Least-squares means for each sample were computed based on the average responses of the 6 panelists per sample.

The assumptions of analysis of variance were assessed by evaluating the residuals for normality and homogeneity of variance using the UNIVARIATE procedure in SAS, with consideration given to the Shapiro-Wilk test for normality using SAS v9.4 (SAS Inst. Inc., Cary, NC, USA). The rate and the extent of pH and temperature decline were analyzed as repeated measures with a compound symmetry covariance structure using the MIXED procedure of SAS, with muscle, time, and their interaction as the fixed effects, and slaughter day as a random effect. When selecting the covariance structure for repeated measures, the Akaike Information Criterion and Bayesian Information Criterion were considered, and the covariance structure presenting the smallest values were selected. Instrumental color, drip loss, and calpain-1 autolysis were analyzed as a randomized complete block design using the GLIMMIX procedure of SAS, with muscle as the fixed effect, and pork carcass nested within slaughter event as the random effect. WBSF and cooking loss were analyzed as a split-split-plot design, with muscle as the whole plot factor, postmortem aging time as the subplot factor, endpoint cooking temperature as the sub-subplot factor, and slaughter day and pork carcass as random effects. The density of desmin, troponin-T, and their degradation products were analyzed as a split-plot design, with muscle as the whole plot factor, postmortem aging time as the subplot factor, and pork carcass and gel as random effects. Sensory traits were analyzed as an incomplete block design, with fixed effects of muscle, endpoint cooking temperature, and their interaction, and random effects of panel session, panelist, pork carcass, and their interactions. Nonsignificant random effects were excluded from the final model. Least-squares differences were determined using the LSMEANS statement with a Tukey-Kramer adjustment. Differences for all analyses were considered significant at P < .05. Stepwise regression analysis was performed with the REG procedure with selection criteria of SLENTRY = 0.15 and SLSTAY = 0.15 to develop prediction equations for the parameters of interest while including slaughter event as dichotomous variables in the model.

The influence of muscle on pH and temperature at each time point was significant (P ≤ .02) apart from temperature at 24 h (P = .34; Table 1). An interaction between muscle and postmortem time (P < .01) was observed for pH and temperature decline, indicating muscles demonstrated a different rate of pH and temperature decline over the 24-h postmortem period (Figure 3). The rate of pH and temperature decline were further quantified by calculating the slope, defined as the change in pH or temperature per hour.

The pH of PM muscle declined the fastest (occurring before the first measurements in this study occurred at 1-h postmortem) among the muscles evaluated in this study (this will be discussed at length in the subsequent section). The LT muscle demonstrated a faster (P < .05) rate of pH decline from 1-h to 3-h postmortem compared with the TB and PM muscles, while no differences (P > .05) in the rate of pH decline at other time periods (3–6h, 6–9 h, 9–12 h, and 12–24 h) were observed among muscles. Overall (1–24 h postmortem), the LT, ST, and GM muscles had similar rates of pH decline, which were greater (P < .05) than the TB muscle, while all muscles had faster (P < .05) rates of pH decline compared with the PM muscle. Muscle-specific differences (P < .01) in temperature decline were observed for all measured time points, except between the 9-h to 12-h postmortem period (P = .49). Overall, when comparing the rate of temperature decline from 1 h to 24 h, the LT and GM muscles had a faster (P < .05) rate of temperature decline compared with the PM and ST muscles.

The rate of pH decline is driven by—and a result of—adenosine triphosphate (ATP) hydrolysis, while ultimate pH is largely determined by the muscle glycogen reserves available at the time of slaughter (Matarneh et al., 2021). Muscle fiber composition plays a critical role in the rate and extent of pH decline. Glycolytic muscle fibers, characterized by greater adenosine triphosphatase (ATPase) activity, elevated levels of glycolytic enzymes, and a more extensive sarcoplasmic reticulum, predominantly undergo glycolytic metabolism. Consequently, glycolytic muscle fibers may exhibit more rapid rates of ATP hydrolysis and pH decline during the early postmortem period compared with oxidative fibers, which primarily rely on oxidative metabolism. Results from this study indicated that the LT muscle underwent a faster rate (P < .05) of pH decline than the TB muscle, especially during the first 3 h of the postmortem period. Rates of pH decline were intermediate for the ST and GM muscles compared with the LT and TB muscles. The PM muscle experienced the slowest and lowest extent of pH decline (P < .05) among the muscles studied in this study (this will be discussed at length in the subsequent section).

The LT, GM, and ST muscles contain predominantly glycolytic type IIB muscle fibers with a low percentage of oxidative type I and type IIA muscle fibers, whereas the TB muscle contains a greater percentage of oxidative type I and type IIA muscle fibers (Melody et al., 2004; Christensen et al., 2004; Karlsson et al., 1999; LeMaster et al., 2024). The greater abundance of glycolytic type IIB muscle fibers in the LT, GM, and ST muscles allowed for a more rapid depletion of glycogen reserves, and a faster progression through the phases of rigor, which accounted for their faster rate of pH decline compared with the more oxidative TB muscle. Moreover, the LT muscle demonstrated the lowest ultimate pH among the 5 muscles analyzed in this study. Similarly, Bohrer et al. (2024) reported that the LT muscle demonstrated the lowest ultimate pH among 9 pork muscles, including the TB, ST, serratus ventralis, biceps femoris, semimembranosus, adductor, rectus femoris, and vastus lateralis muscles.

In contrast to the other muscles evaluated in this study, including the glycolytic LT and GM muscles, the oxidative PM muscle demonstrated the lowest (P < .05) pH at 1-h postmortem (pH = 5.73) compared with all other muscles evaluated. However, the pH of the PM muscle at 12-h and 24-h postmortem (pH = 5.71 and 5.67, respectively) was not different (P > .05) compared with the ST, TB, and GM muscles and was actually greater (P < .05) than the LT muscle. This demonstrates that the majority of the pH decline in the PM muscle occurred within the first hour postmortem. A similar observation was documented by Melody et al. (2004), Lefaucheur (2010), and Zhang et al. (2013), all of whom reported that the PM muscle had a lower pH at 1-h postmortem and a greater pH at 24-h postmortem compared with the LTL muscle. The low pH at 1-h postmortem in the PM muscle could be due to its limited glycogen reserves at slaughter, which are nearly depleted within the first hour postmortem, thereby restricting postmortem glycolysis and pH decline, ultimately leading to its relatively high pH at 24-h postmortem. Moreover, upon ATP depletion, the impaired calcium ATPase pump in the sarcoplasmic reticulum may have resulted in a cytosolic calcium overload, which could have been subsequently sequestered by mitochondria. Calcium overload in the mitochondria has been shown to induce the translocation of H⁺ from the matrix to the intermembrane space, facilitating ATP hydrolysis (Matarneh et al., 2023). The lower membrane integrity and greater permeability of the mitochondrial membrane in the PM muscle compared with other muscles could allow for a greater rate of ATP hydrolysis (Zhang et al., 2020).

Instrumental color of each muscle was evaluated on day 1 postmortem. Muscle influenced (P < .01) L* (lightness), a* (redness), and b* (yellowness). Pork LT, ST, and GM muscles demonstrated greater (P < .05) L* than PM and TB muscles. The PM muscle exhibited the greatest (P < .05) a*, while a* of the ST and TB muscles were at intermediate levels compared with other muscles, and the LT and GM muscles had the lowest (P < .05) a*. The PM and ST muscles had greater (P < .05) b* compared with other muscles, while the LT muscle had greater (P < .05) b* compared with the TB muscle.

Muscle-specific variation in pork color has been well documented by several previous studies. For instance, Brewer et al. (2001), Arkfeld et al. (2016), and Bohrer et al. (2024) suggested that pork loin (LTL muscle) exhibited greater lightness, lower redness, and greater yellowness compared with ham muscles, including the ST, GM, biceps femoris, semimembranosus, adductor, rectus femoris, and vastus lateralis muscles as well as shoulder muscles, such as the TB and serratus ventralis muscles. Moreover, Barkley et al. (2023) reported that the LTL muscle demonstrated lower oxygen consumption rates, greater metmyoglobin reducing activity, and greater color stability than the TB or PM muscles during retail display. Color differences among pork muscles could be largely attributed to the variation in muscle fiber types, which influence myoglobin content, mitochondrial functionality, pH, and water-holding capacity. Oxidative muscle fibers contain greater concentrations of myoglobin and mitochondria for aerobic metabolism and generate more energy than glycolytic muscle fibers. Particularly, myoglobin content has been shown to strongly relate to the red pigmentation of pork (Kim et al., 2010). Therefore, muscles with a greater proportion of glycolytic type IIB muscle fibers and a lower proportion of oxidative type I muscle fibers tend to exhibit greater lightness and lower redness, often resulting in a “pale” appearance. This characteristic aligns with the findings of this study, where the LT, ST, and GM muscles—predominantly composed of glycolytic type IIB muscle fibers—demonstrated greater lightness and lower redness compared with the PM and TB muscles, which contain a greater proportion of oxidative type I fibers.

Stepwise regression analysis was conducted to predict the instrumental color of each individual muscle using pH and temperature at different time points during the early postmortem process (data presented in supplementary tables). For the LT muscle, pH at 1-h postmortem explained the greatest variation (R² = 0.61) in L* (measured at 1-d postmortem), while temperature at 6-h postmortem explained the greatest variation (R² = 0.43) in L* (measured at 1-d postmortem). For the PM and GM muscles, pH at 24-h postmortem explained the greatest variation in a* (measured at 1-d postmortem; R² = 0.33 and R² = 0.24, respectively). For the TB muscle, pH at 24-h postmortem explained the greatest variation in L* (measured at 1-d postmortem; R² = 0.23). These findings suggested that the relationship among color (measured at 1-d postmortem), rate and extent of pH decline, and rate of temperature decline appears to differ among muscles.

Drip loss was influenced (P < .01) by muscle. The LT muscle had the greatest (P < .05) drip loss, whereas the PM, ST, and TB muscles exhibited the lowest (P < .05) drip loss. Drip loss for the GM muscle differed (P < .05) when compared with all other muscles and was at intermediate levels compared with the LT muscle and the PM, ST, and TB muscles. Variation in water-holding capacity is known to be attributed to the differences in the rate and extent of pH decline, degradation of intermediate filament proteins (Melody et al., 2004; Johnson et al., 2023), and protein oxidation (Lund et al., 2007). The current study evaluated the differences in the rate and extent of pH decline and proteolysis among the muscles, which are likely responsible for the observed variation in drip loss. In the current study, the lower ultimate pH observed in the LT and GM muscles (approximately 0.2 U and 0.1 U lower, respectively) compared with the PM, ST, and TB muscles likely contributed to their reduced water-holding capacity, as evidenced by greater levels of drip loss. Similarly, Warner et al. (1993) compared pork quality characteristics of 10 muscles and documented greater levels of exudate in the LTL, GM, and ST muscles compared with the PM and TB muscles. Melody et al. (2004) and LeMaster et al. (2024) reported that the LTL muscle exhibited greater levels of drip loss and purge loss compared with the PM muscle.

Stepwise regression analysis suggested that pH at 1-h postmortem accounted for the greatest variation in drip loss (R² = 0.50) for the ST muscle (data presented in supplementary tables). For the LT muscle, pH at 6-h postmortem explained the greatest variation in drip loss (R² = 0.30), while temperature at 1-h postmortem explained the greatest variation in drip loss (R² = 0.32). In contrast, pH and temperature at other measured time points explained less than 30% of variation in drip loss for other muscles (R² < 0.30). It is worth noting that pH at specific temperatures during the early postmortem period may be a more meaningful predictor of drip loss when compared with pH at specific time points during the early postmortem period. For instance, Dorleku et al. (2025) suggested that pH at 36°C explained 27.5% of the variability in purge loss of the pork LT muscle.

The ST muscle had the greatest IMF (P < .05; approximately 10.29%) and the lowest moisture content (P < .05; approximately 69.43%) compared with the other muscles. Consistent with our findings, Kim et al. (2008) reported the ST muscle demonstrated the greatest IMF, followed by the TB, GM, and LT muscles.

Interactions were observed for WBSF between postmortem aging day and endpoint cooking temperature (P = .02) as well as between muscle and endpoint cooking temperature (P < .01; Table 2). While WBSF decreased when extending postmortem aging from 1 d to 10 d and with a reduction in the endpoint cooking temperature from 71°C to 63°C, the numerical change for WBSF by lowering endpoint cooking temperature was less for samples aged 10 d (Δ = 0.35 kg) compared with samples aged for 1 or 3 d (Δ = 0.54 kg and Δ = 0.55 kg, respectively; Figure 4A). This indicated that reducing endpoint cooking temperature from 71°C to 63°C played a more critical role for pork aged for 1-d or 3-d postmortem compared with those subjected to 10 d of aging. Moreover, different muscles demonstrated varying levels of reduction in WBSF in response to the reduction of endpoint cooking temperature from 71°C to 63°C (Figure 5A). As the endpoint cooking temperature was reduced from 71°C to 63°C, the GM muscle experienced the greatest levels of numerical change in WBSF values (Δ = 0.71 kg), while the LT and TB muscles exhibited the lowest levels of numerical change in WBSF values (Δ = 0.28 kg and Δ = 0.27 kg, respectively), and the PM and ST muscles were intermediate (Δ = 0.59 kg and Δ = 0.55 kg, respectively). This suggested that lowering the endpoint cooking temperature had a greater impact on the instrumental tenderness of the GM compared with other pork muscles.

Cooking loss was influenced by muscle (P < .01), and the interaction between postmortem aging days and endpoint cooking temperature (P = .02). The greatest (P < .05) level of cooking loss was observed in the ST and GM muscles, while the LT muscle exhibited the lowest (P < .05) level of cooking loss compared with all other muscles. Similar to WBSF, the magnitude of change in cooking loss caused by endpoint cooking temperature varied across different postmortem aging days. Specifically, decreasing the endpoint cooking temperature from 71°C to 63°C contributed to a greater level of numerical change for cooking loss in muscles subjected to 3 d and 10 d of postmortem aging (Δ = 5.50% and Δ = 5.73%, respectively) compared with those aged for 1-d postmortem (Δ = 3.94%; Figure 4B).

In agreement with our results, numerous research studies have concluded that pork cooked to greater endpoint cooking temperatures exhibited greater WBSF and cooking loss (Rincker et al., 2008; Bryan et al., 2019; Gaffield et al., 2020). Muscle-specific differences in pork tenderness observed in this study could be primarily attributed to variations in muscle fiber composition (Kim et al., 2013), collagen content and solubility (Rhee et al., 2004; Żochowska et al., 2005), IMF (Nishimura, 2010), sarcomere length (Wheeler et al., 2000; Warner et al., 2021), and the extent of proteolysis (Muroya et al., 2010). These structural and biochemical characteristics influence thermal stability (LeMaster et al., 2024), muscle fiber shrinkage (Vaskoska, 2020), and connective tissue behavior during cooking (Vaskoska et al., 2020), ultimately leading to differences in WBSF among muscles.

An effect (P < .01) of muscle was observed for unautolyzed calpain-1 (80 kDa), partially autolyzed calpain-1 (78 kDa), and fully autolyzed calpain-1 (76 kDa; Table 3). The PM and ST muscles exhibited a greater proportion (P < .05) of unautolyzed calpain-1 (80 kDa) compared with the LT and GM muscles, with the TB muscle demonstrating an intermediate level. The ST and TB muscles had greater proportion (P < .05) of partially autolyzed calpain-1 (78 kDa) compared with the LT and GM muscles, while the PM was at an intermediate level. Moreover, the LT and GM muscles demonstrated greater proportion (P < .05) of fully autolyzed calpain-1 (76 kDa) compared with the PM, ST, and TB muscles. This suggested that LT and GM muscles exhibited greater extent of calpain-1 autolysis at (and prior to) 24-h postmortem compared with the PM, ST, and TB muscles.

An interaction (P < .05) between muscle and postmortem aging time was observed for desmin (and its degradation products) and troponin-T (and its degradation products; Table 4). This indicated that the extent of proteolysis varied among muscles, as evidenced by differential changes in the abundance of intact desmin (55 kDa), desmin degradation products (42 kDa and 38 kDa), intact troponin-T (37 kDa), and troponin-T degradation products (30 kDa and 28 kDa) during postmortem aging. The ST and TB muscles exhibited a greater abundance (P < .05) of intact desmin (55 kDa) compared with the LT, PM, and GM muscles at both 1-d and 10-d postmortem. Moreover, the ST and TB muscles demonstrated limited changes (P > .05) in intact desmin (55 kDa) and desmin degradation products (42 kDa and 38 kDa) during postmortem aging. In contrast, the LT, GM, and PM muscles exhibited a decrease (P < .05) in the abundance of intact desmin (55 kDa) and an increase (P < .05) in the abundance of desmin degradation product (38 kDa) between 1-d and 10-d postmortem aging.

The LT muscle demonstrated the greatest abundance (P < .05) of intact troponin-T (37 kDa) at 1-d postmortem, and the greatest abundance (P < .05) of troponin-T degradation products (30 kDa and 28 kDa) at 10-d postmortem compared with other muscles. Additionally, an increase (P < .05) in the abundance of troponin-T degradation products (30 kDa and 28 kDa) were observed from 1-d to 10-d postmortem in the LT and GM muscles, whereas the PM, ST, and TB muscles exhibited minimal changes (P > .05) in intact troponin-T (37 kDa) and troponin-T degradation products (30 kDa and 28 kDa) during postmortem aging.

Overall, these results suggested that the LT and GM muscles exhibited the greatest level of postmortem proteolysis during aging, followed by the PM muscle, while the ST and TB muscles demonstrated the lowest level of proteolysis. This indicated that the LT and GM muscles, which demonstrated greater WBSF but also greater proteolytic potential, may benefit the most from extended periods of postmortem aging—potentially up to 10 d to 14 d. In contrast, shorter aging periods may be sufficient for the ST, TB, and PM muscles.

In agreement with our findings, Wheeler et al. (2000) documented pork LTL muscles had greater levels of desmin degradation compared with the ST and TB muscles. Christensen et al. (2004) reported a faster rate of troponin-T and desmin degradation for pork LTL and semimembranosus muscles compared with ST, vastus intermedius, and soleus muscles, attributing these differences to the variations in muscle fiber type, calpain-1 activity, and the rate of pH decline among muscles. Melody et al. (2004) reported a faster rate of pH decline, an increased rate of calpain-1 autolysis, and earlier degradation of desmin (within the first 24-h postmortem) for the PM compared with the LTL muscle. However, the earlier activation of calpain-1 in the PM might result in an earlier loss of calpain-1 activity, leading to reduced protein degradation and tenderization during extended periods of postmortem aging (up to 120-h postmortem) compared with the LT muscle. Together, these previous studies imply that the rate of pH decline within the first 6-h postmortem could influence the calpain-1 activity and ultimately influence drip loss and the rate of postmortem tenderization (Melody et al., 2004).

Stepwise regression analysis was conducted to predict the calpain-1 autolysis for individual muscles using pH and temperature at different time points (data presented in supplementary tables). For the LT muscle, pH at 1-h postmortem accounted for the greatest variation (R² = 0.34) in autolyzed calpain-1 (76 kDa). For the PM and GM muscles, pH at 3-h postmortem explained the greatest variation (R² = 0.20 and R² = 0.23, respectively) in autolyzed calpain-1 (76 kDa). For the ST muscle, pH at 6-h postmortem explained the greatest variation (R² = 0.27) in autolyzed calpain-1 (76 kDa). For the TB muscle, pH at 9-h postmortem explained the greatest variation (R² = 0.35) in autolyzed calpain-1 (76 kDa). Additionally, for the LT and ST muscles, temperature at 24-h postmortem accounted for the greatest variation (R² = 0.31 and R² = 0.17) in autolyzed calpain-1 (76 kDa). The contributions of pH and temperature to calpain-1 autolysis during the early postmortem process (and the differences observed among muscles) may be partially attributed to the rate and extent of pH decline and the rate of temperature decline. Carlin et al. (2006) reported that calpain-1 exhibited greater activity at pH 6.5 than pH 7.5. Pomponio et al. (2010) suggested that a faster rate of pH decline in pork resulted in increased autolysis of calpain-1 and consequently an earlier loss of its activity. Therefore, while the rate of early pH decline could influence the activity of calpain-1, it may not necessarily impact the extent of proteolysis that occurs. Mohrhauser et al. (2014) documented that calpain-1 activity and troponin-T degradation were not influenced by fast or slow pH decline treatments in beef. However, increased activation of calpain-1 and a reduction in troponin-T were observed at warmer temperatures. Ramos et al. (2020) observed the delayed calpain-1 autolysis in Brahman longissimus lumborum (LL) muscle that exhibited resistance of pH decline and greater ATP content. The authors of that study suggested that calpain-1 autolysis was influenced by mitochondria related to energy production and calcium sequestration. These results indicated that the variation pH, temperature, and energy availability may collectively lead to muscle-specific proteolytic activity of calpain-1.

An interaction (P < .01) between muscle and endpoint cooking temperature was observed for sensory tenderness and sensory juiciness (Figure 5B and Figure 5C). Generally, decreasing endpoint cooking temperature improved sensory tenderness and sensory juiciness scores; however, the extent of reduction varied among the muscles. At an endpoint cooking temperature of 63°C, the PM muscle received greater (P < .05) sensory tenderness scores compared with the LT and TB muscles, with the ST and GM muscles at intermediate levels for sensory tenderness that did not differ from other muscles. At an endpoint cooking temperature of 71°C, the PM muscle was scored as the most tender (P < .05) among the muscles, while the other muscles demonstrated similar (P > .05) scores for sensory tenderness when compared with one another. Furthermore, the sensory tenderness of the PM muscle was not influenced (P > .05) by endpoint cooking temperature (either 63°C or 71°C), whereas improvements (P < .05) in sensory tenderness were observed for other muscles at endpoint cooking temperatures of 63°C compared with 71°C. Overall, these findings suggested that endpoint cooking temperature was less impactful for the PM muscle compared with other muscles in terms of sensory tenderness. When cooked to an endpoint temperature of 63°C, the ST muscle exhibited greater (P < .05) sensory juiciness compared with the LT and PM muscles, with the TB and GM muscles at intermediate levels for sensory juiciness. At an endpoint temperature of 71°C, the ST muscle received a greater (P < .05) sensory juiciness scores compared with the LT, GM, and PM muscles, while the TB muscle was an intermediate level for sensory juiciness. Although all muscles experienced an improvement (P < .05) in sensory juiciness as the endpoint cooking temperature was reduced from 71°C to 63°C, the LT muscle exhibited the greatest numerical increase in sensory juiciness (Δ = 4.70), whereas the ST muscle showed the smallest numerical increase in sensory juiciness (Δ = 2.99). Furthermore, all muscles exhibited a sensory juiciness score greater than 7.5 when cooked to an endpoint temperature of 63°C, and all muscles, except for the ST, demonstrated a sensory juiciness score lower than 7.5 when cooked to an endpoint temperature of 71°C.

Sensory chewiness and sensory pork flavor were not influenced (P > .05) by the interaction between muscle and endpoint cooking temperature; however, both were influenced (P < .01) by muscle and endpoint cooking temperature (Figure 5D and Figure 5E). Reducing the endpoint cooking temperature from 71°C to 63°C lowered (P < .01) sensory chewiness levels and increased (P < .01) sensory pork flavor. The PM muscle was perceived as less chewy (P < .05) compared with most other muscles at both endpoint cooking temperatures. Exceptions for when the PM muscle was not different than other muscles were for the ST and GM muscles at the endpoint cooking temperature of 63°C and the ST muscle at the endpoint cooking temperature of 71°C (P > .05). For pork flavor, no difference (P > .05) between muscles was observed at the endpoint cooking temperature of 63°C, while the PM and ST muscles had greater levels (P < .05) of pork flavor compared with the LT and GM muscles at the endpoint cooking temperature of 71°C.

Consistent with our results, previous research documented that reducing endpoint cooking temperatures (i.e., degree of doneness) from 71°C to 63°C has been shown to improve sensory attributes by increasing sensory tenderness and sensory juiciness of pork LTL muscles (Rincker et al., 2008; Bryan et al., 2019; Honegger et al., 2019; Honegger et al., 2022). Moeller et al. (2010) demonstrated that incremental increases in cooking temperature (62.8°C, 68.3°C, 73.9°C, and 79.4°C) reduced sensory juiciness and sensory tenderness scores when evaluated by a trained sensory panel. Klehm et al. (2018) suggested that cooking pork LTL muscles to a lower endpoint cooking temperature (63°C vs. 71°C) was a more effective way to improve eating experience than the proposed US Department of Agriculture quality grades (based on subjective color and marbling) or packaging type. Moreover, Honegger et al. (2019) reported that endpoint cooking temperature of the pork LTL muscle has a greater impact on consumer eating experience than visual color, marbling, or ultimate pH.

The influence of endpoint cooking temperature on sensory attributes of pork likely depends on muscle. In the current study, while sensory tenderness was improved at lower endpoint cooking temperature across most muscles, the observed magnitude of change for each muscle could be attributed to content and solubility of connective tissue. In support, Bejerholm and Aaslyng (2004) documented that sensory tenderness of pork LTL muscles decreased when endpoint cooking temperature was increased from 65°C to 80°C. However, sensory tenderness of the pork biceps femoris muscles decreased when the endpoint cooking temperature increased from 65°C to 75°C and then was improved when the endpoint cooking temperature increased from 75°C to 80°C. It was suggested that the differences between the 2 muscles could be attributed to their variation in connective tissue content and solubility (Bejerholm and Aaslyng, 2004). Similar to our results, Wheeler et al. (2000) reported that the ST and TB muscles exhibited greater sensory tenderness, sensory juiciness, and sensory pork flavor compared with the LL muscle at an endpoint cooking temperature of 70°C. Furthermore, Channon et al. (2016) observed that pork LTL muscles obtained lower sensory tenderness, sensory juiciness, and sensory flavor scores compared with shoulder muscles such as the TB, regardless of postmortem aging period, endpoint cooking temperatures, and cooking methods.