Effects of persimmon peel supplementation on pork quality, palatability, fatty acid composition, and cholesterol level

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at the Kyungpook National University. All experiments were performed in accordance with the Prevention of Cruelty to Animals Act (1986).

A total of 40 crossbred pigs (Landrace × Yorkshire × Duroc; 22 gilts and 18 castrated pigs) with an average initial body weight of 48.9 ± 2.63 kg at 103 d of age were used in this experiment. The pigs were randomly allocated to 4 groups consisting of 10 pigs per group, and all 40 individuals were weighed at the beginning and at the end of the experiment. The environmental conditions to which the pigs were exposed were the same both before and after slaughter. The nutrient levels used for the control and in the treatments were based on standards described in National Research Council [7]. There were no significant differences between the genders or among pig pens in any measurements. Persimmon peel was obtained from persimmon trees growing in Sangju, GyeongBuk, South Korea, and air-dried for a month, then pulverized. The experiment was divided into 4 groups consisting of basal diet (Table 1) with premix supplementation at the level of 0 % (control); 0.3 % persimmon peel powder (Treatment 1; T1); 0.6 % persimmon peel powder (T2); and 0.9 % persimmon peel powder (T3) (Table 2). Each pig was fed ad libitum. The major components of persimmon peel powder supplement are presented in Table 3. Persimmon peel powder showed a higher level of total phenolic compound, and was rich in tannin content (24.1 mg/100 g).

All pigs were transported to a commercial abattoir under the same handling conditions at the same age (188 d of age; 135 ± 3.4 kg), and slaughtered by electric stunning during the autumnal period under the supervision of the Korean grading service for animal products. All animals were exsanguinated and placed in a dehairer at 65 °C for 5 min, with remaining hair removed with a flame and knife after exiting from the dehairer. Carcasses were weighed following evisceration. The mean of the loin eye area (measured at the level of the last rib) and back-fat thickness (measured at the 11th and last thoracic vertebrae) was used.

At 45 min postmortem, muscle samples were taken from the longissimus dorsi muscles at the 7th and 8th thoracic vertebrae to measure muscle pH (pH_{45 min}). After 24 h postmortem in a 4 °C cold room, pork loins were collected for meat quality measurements, and samples were frozen and stored at −20 °C for later analysis of sensory characteristics, fatty acid composition, and cholesterol content of cooked pork.

In order to estimate the postmortem glycolytic rate, muscle pH at 45 min (pH_{45 min}) and 24 h (pH_{24 h}) postmortem were measured directly by piercing the carcasses at the 8th and 9th thoracic vertebrae with a spear-type portable pH meter (IQ-150 pH meter and PH77-SS probe, IQ Scientific Instruments Inc., San Diego, CA).

At 24 h postmortem, meat color was measured with a Minolta chromometer (CR-400, Minolta Camera Co., Japan) following surface exposure for 30 min at 4 °C. Results were expressed as Commission Internationale de l’Eclairage [8] lightness (L^*), redness (a^*), and yellowness (b^*) values, with the average of three measurements used. Drip loss, filter-paper fluid uptake (FFU), and cooking loss were measured to assess water-holding capacity (WHC). To determine drip loss, samples weighing approximately 80 g were trimmed and accurately weighed. Each sample was then placed in a square netting (25 cm²) and then suspended in an inflated plastic bag for 48 h at 4 °C, ensuring that the sample did not make contact with the bag, after which they were re-weighed [9], with drip loss calculated as the percentage change in weight [9]. For FFU and cooking loss, pork loins were cut into 2-cm thick chops. To measure FFU, a filter paper (Whatman #2, 42.5 mm in diameter) was pre-weighed, placed on the surface of a chop for less than 2 s to absorb its fluids, and then weighed again; FFU was expressed as mg of exudates absorbed into the filter paper [10]. To measure cooking loss, the samples were put in thin-walled polyethylene bags and placed in a water bath (80 °C) until the internal temperature (measured using a thermometer with a handled probe; TES-1300, TES Electrical Electronic Co., Taiwan) reached 71 °C. Pork samples were then cooled in ice water for 15 min. The cooled samples were taken from the polyethylene bag, blotted dry, and weighed. Cooking loss was determined by weighing the chops before and after cooking [9]. Preparation of cooked meat samples for Warner–Bratzler shear force (WBS) was similar to the procedure for cooking loss, and WBS was performed with six to ten cores (1.27 cm in diameter) removed from the steak parallel to the longitudinal orientation of the muscle fibers. WBS was determined using an Instron Universal Testing Machine (Model 1011, Instron Corp., USA) equipped with a Warner–Bratzler shearing device.

Color and marbling were evaluated using fresh pork loin at 24 h postmortem based on the National Pork Producer Council (NPPC) color and marbling standards [11]. Twenty-mm thick pork loin chops were exposed to air for 30 min at 4 °C to bloom. Color scores ranged from 1 (pale) to 6 (dark), and marbling score ranged from 1 (1 % intramuscular fat [IMF] content) to 10 (10 % IMF content).

A sensory panel consisting of 12 pork-consuming individuals (20–40 years of age, 6 females and 6 males) was employed to evaluate the sensory attributes of the cooked pork. Panelists were selected based on their interest and availability. All training and testing was conducted at Kyungpook National University. Before sensory evaluation, all panelists were trained for at least 6 months (3 times per wk) and for up to 1 h in each training session. Panelist training was performed in accordance with practices described by the American Meat Science Association [12] and previously published procedures [13]. All pork samples were evaluated twice.

Samples were thawed overnight at 4 °C and then cooked in a humid heat oven (MCS312CF4, Electrolux, Sweden, set at 180 °C) to an internal temperature of 71 °C, as measured by a TES-1300 thermometer (TES Electrical Electronic Co., Taiwan). Samples were then immediately sliced into 1.3 cm × 1.3 cm × 1.3 cm pieces, which were subsequently randomly selected to minimize bias. The chosen samples were placed into 1-ounce lidded glass jars labeled with random three-digit codes and kept in a water bath (54 °C) until presented to the panelists. All samples were presented to panelists simultaneously on a compartmented plate, with an interval of approximately 5 min between evaluations of consecutive samples. For sensory evaluation, panelists were seated in private booths under incandescent light, and served distilled water (at room temperature) and salt-free crackers before the first sample and between samples to cleanse their palate.

Cooked samples were evaluated for softness (force required to compress the meat sample between the molar teeth; 1 = very hard; 9 = very soft), initial tenderness (force required to chew 3 times after initial compression; 1 = very tough; 9 = very tender), chewiness (energy required at the 9th chew to swallow at a constant rate; 1 = very chewy; 9 = very tender), rate of breakdown (number of chews required for the sample to disintegrate during the mastication process in preparation for swallowing; 1 = very slow; 9 = very fast), mouth coating (amount of oil/fat left in the mouth surface; 1 = none; 9 = very high), amount of perceptible residue (amount of connective tissue remaining upon complete disintegration of meat sample; 1 = abundant; 9 = none), juiciness (amount of moisture released after 5 chews; 1 = not juicy; 9 = extremely juicy), flavor intensity (intensity of pork flavor after 8 chews; 1 = no pork flavor; 9 = full pork flavor), and off-flavor intensity (intensity of any flavor or after-taste perceived as inappropriate to cooked pork; 1 = very strong; 9 = very weak) [12, 13].

The content of IMF was determined using the Soxhlet method with a solvent extraction system [14]; fat was extracted based on the method described by Folch et al. [15]. Briefly, 1.5 g of homogenized meat was twice blended with an extraction solvent of chloroform/methanol (2:1, v/v), filtered, and then placed in a separator funnel and mixed with saline solution (0.9 % NaCl). After separation into two phases, the aqueous methanol fraction was discarded, whereas the chloroform lipid fraction was washed with extraction solvent. Following further filtration and evaporation by means of a rotary evaporator, the lipid extracts were transferred to test-tubes for subsequent gas chromatographic analysis. Duplicates of lipids were methylated by adding 2 ml of sodium methoxide, distilled water, and heptane. Gas chromatograph analysis was performed on a Gas Chromatography–Mass Selective Detector (GC, Agilent 7890 N, USA; MSD, Agilent 5975A, USA) equipped with a HP-INNOWAX column (length 30 m, internal diameter 0.25 mm, film thickness 0.25 μm). The operating conditions consisted of a helium flow rate of 0.7 ml/min, the FID detector set to 260 °C, the split-splitless injector set to 220 °C with an injection rate of 120 ml/min, and an injection volume of 1 μl. The temperature program of the column consisted of 4 min at 140 °C followed by a subsequent increase to 220 °C at a rate of 4 °C/min. The retention time and area of each peak were computed using Agilent software. The individual fatty acid peaks were identified by comparing the retention times with those of known mixtures of standard fatty acids (FAME, Sigma-Aldrich CO., USA). Fatty acid composition was expressed as the percentage of total methylated fatty acid. The hypocholesterolemic/hypercholesterolemic potential of the pork samples was calculated according to procedures set out by Santos-Silva et al. [16] using the h/H index [h/H = (18:1n-9 + 18:2n-6 + 20:4n-6 + 18:3n-3 + 20:5n-3 + 22:5n-3 + 22:6n-3)/(14:0 + 16:0)].

Concentration of total and low-density lipoprotein (LDL) cholesterol in muscle tissues were determined with enzymatic method using a Bio Merieux kit (Bio-Meraux Lab, Paris, France), and concentrations of high-density lipoprotein (HDL) and total cholesterol in muscle tissues were measured using Alpha Diagnostics kit (Alpha Diagnostics Inc., CA, USA).

Statistical analyses were performed using SAS software [17]. Analyses of variance were conducted to evaluate potential differences (P < 0.05) between the different treatments. The results are presented as means for each group along with their standard deviations.

Carcass characteristics of the control and treatment groups are presented in Table 4. There were no significant differences in carcass weight, loin eye area, and back-fat thickness among the treatments (P > 0.05). The effects of persimmon peel supplementation on meat quality characteristics are shown in Table 5. There were no significant differences in the postmortem glycolytic rate, including muscle pH_{45 min} and pH_{24 h}, among the groups (P > 0.05). Lightness (48.1 and 47.1, P > 0.05), drip loss (2.68 and 2.47 %, P > 0.05), and cooking loss (22.8 and 22.5 %, P > 0.05) were also not significantly different between the control and T3 groups, respectively. No significant difference was also observed between the control and T3 groups in FFU, even though pork samples from group T3 had a significantly lower FFU than that of samples from group T1 (29.0 vs. 69.6 mg, P < 0.05). WBS as an indicator of tenderness was lower in the T3 group than in the control group (39.7 vs. 48.9 N, P < 0.05). Neither NPPC color nor marbling scores were affected by dietary supplementation (P > 0.05).

Eating quality characteristics of pork loin for each group classified by the supplemental levels of persimmon peel are presented in Table 6. As with the instrumental tenderness parameters, a few tenderness attributes were found to be significantly different among the treatment groups. Pork samples from group T3 had higher values for softness (6.54 vs. 5.07, P < 0.05) and initial tenderness (6.20 vs. 4.82, P < 0.05) compared to samples from the control group, respectively, whereas softness (5.51, P > 0.05) and initial tenderness (5.19, P > 0.05) values of the T1 group were similar to those of the control group. There were no significant differences among the treatment groups (P > 0.05) for other tenderness attributes, including chewiness, rate of breakdown, mouth coating, and amount of perceptible residue; moreover, there were no significant difference in juiciness and flavor intensity among the treatment groups (P > 0.05).

There were marked differences in fatty acid composition and cholesterol levels (Table 7), even though no significant differences were observed in IMF among the treatment groups (P > 0.05). Compared with pigs fed the control diet, pork from pigs fed T3 diets had a lower percentage of saturated fatty acid (SFA, 41.5 vs. 43.7 %, P < 0.05) and higher percentage of polyunsaturated FA (PUFA, 13.6 vs. 12.1 %, P < 0.05). The lower SFA in the T3 group was most probably caused by differences in the composition of palmitic acid (16:0); the percentage of palmitic acid was higher in the control group than in the T3 group (26.95 vs. 23.9 %, P < 0.01; data not shown). There were no significant differences in SFA (43.2 %, P > 0.05), monounsaturated FA (MUFA, 44.1 vs. 43.7 %, P > 0.05), n-6 (11.6 vs. 12.4 %, P > 0.05), and n-3 (0.57 vs. 0.67 %, P > 0.05) between the control and T2 groups. The h/H index of the T3 group was the highest compared to the other groups (2.09, P < 0.01), and no significant difference was observed between the control and T1 groups (1.78 vs. 1.83, P > 0.05). Pork samples from group T3 had lower concentrations of total cholesterol (94.4 vs. 99.1 mg/100 g, P < 0.05) and LDL cholesterol (47.4 vs. 51.4 mg/100 g, P < 0.05) than those of the control group, whereas no significant differences were observed in the concentration of total cholesterol between the control and T1 groups (96.4 mg/100 g, P > 0.05).