Pork Quality and Muscle Fiber Type


The study of the regularities of the development of muscular and adipose tissues of pigs is part of the problem of creating high-quality agricultural meat raw materials for further human consumption, including through the definition of mechanisms responsible for the formation of necessary structural features. It is known that pork has phenomenal market importance for humankind, which is caused by the relative ease of cultivation and observation and the high nutritional value of animal muscle and adipose tissues. According to statistical calculations by Ritchie and Roser, pork is the most popular sort of meat in the world, and global production has increased fivefold since 1961. Thornton points out that since the 2000s, humanity has consumed more than a billion pigs annually, and that number is overgrowing. Strictly speaking, these figures lead to the conclusion that there is an urgent need both to improve quality control procedures and to investigate the patterns associated with the development of pigs’ muscle and adipose tissues.

It is known that mammalian muscle tissue is evolutionarily represented by two types of fibers: the primary fibers form a matrix framework on which the secondary muscle fibers develop. The process of muscle fiber formation is activated at the fusion of myoblasts, which, in natural conditions, is associated with the beginning of muscle cell differentiation. In this process, many different genes that trigger the mechanisms of fiber evolution are involved in the coordination (Glaser and Suzuki 70). It seems evident that the process of building muscle tissue, mainly composed of primary and secondary fibers, can be altered by certain factors, including the perinatal development of the pig embryo.

Numerous studies on the relationship between animal embryonic development, mass at birth, and muscle and adipose tissue formation have been conducted in the fields of veterinary medicine, agricultural sciences, and medicine. Understanding which factors directly or indirectly influence the intensity of muscle fiber formation, its quantity, size, and availability of the most important biochemical molecules, such as actin, myosin, tropomyosin, troponin, and other enzymes, makes it possible both to predict the development of the population and to manipulate the conditions for growing animals with the given properties. Finally, it is well known that the quality of supplied market pork is directly affected by the intensity of the tissue location, size, and volume of muscle fibers, as well as by its differentiation (Kim et al. 5530). This becomes especially important in the conditions of market sales of agricultural raw materials, as customers are primarily interested in buying pork with a structured and dense fiber material rich in nutrients. Nevertheless, there is no unequivocal opinion in the academic community on the relationship between pig birth weight and the intensity of development of primary and secondary muscle fibers. While some researchers are convinced that there are no statistically significant differences between groups of pigs with different weight parameters at birth and the number of muscle fibers of both types contained in m. semitendinosus, others refute these findings with their research (Handel and Stickland 316; Gondret 101). On the other hand, not only the definition of the cause-and-effect mechanism of functional muscle tissue development but also possible differentiation by fiber type has scientific value. In particular, one cannot deny the potential effect of the low body weight of piglets on the development or underdevelopment of muscle fibers of the first and second types at different life stages. Future case studies should determine the nature of the relationship between variables and explain the mechanism of fiber development. This dictates the need for experimental work to determine muscle fiber types and the quality of raw pork products, which is the main objective of this research paper.


Sample Preparation

Histological analysis of muscle tissue samples begins with the isolation of biological material from the slaughtered animal. Fragment of muscle fibers taken from pig m. semitendinosus was pre-labeled and then subjected to rapid cooling to prevent morphological distortions of the fibers’ cellular structure. Instant freezing of the sample is not acceptable, as this process can stimulate the intensification of intracellular vapor barrier processes, so pre-cooling of the cut muscle tissue fragment in the cold room for 20-30 minutes was used. After cooling, the sample was placed in an instant freezing medium depending on the method used: acetone, dry ice, isopentane, or liquid nitrogen. The plastic foam chamber was filled with coolant, and then a stainless-steel metal cup was placed inside, filled with an alternative freezer so that the liquid levels matched. A muscle sample was placed in a metal bowl for a period of ten to sixty minutes, depending on the type of refrigerant. After a while, the frozen sample was pulled out of the refrigeration chamber and moved into a plastic container with dry ice at -80 ℃ for long-term storage. For further investigation, a 1 cm × 1 cm × 1.5 cm cube was cut off with a sharp knife and placed in a cryostatic chamber at -25 ℃ for 25-30 minutes. From the sample prepared for cryo-section, the material 8-10 μm thick was cut off with a sharp blade and placed on a slide. To extend the shelf life and prevent scratches and cracks, a piece of muscle tissue was covered with a second slide.

NADH-TR staining methods

Chemical reagents have been prepared in advance with the NADH-TR to investigate the qualitative characteristics of a muscle tissue sample and determine the type of differentiated muscle fibers. Solutions of 0.2 M of sodium dihydro phosphate and 0.2 M of disodium dihydro phosphate were mixed to produce a sodium phosphate buffer (7.4 pH). An additional acetate buffer was obtained by mixing 0.2 M of sodium acetate and 0.2 M of acetic acid. The incubation environment consisted of a mixture of nitro blue tetrazolium chloride (240 mg) with prepared phosphate buffer (120 ml), NADH (0.125 g), and distilled water (120 ml). After incubation in the medium for 20 minutes, a sample of muscle tissue was placed in the acetate buffer for 50, minutes, washed with distilled water, and subsequently studied under a microscope.

Myosin ATPase staining methods

For the preparation of alkaline incubation environment 0.1 M sodium barbital (144 ml), 0.18 M calcium chloride (72 ml), and distilled water (504 ml) were mixed. By adding sodium hydroxide to the solution, the pH was regulated at 9.4. Subsequently, the sample was placed in an ATP incubation solution synthesized by mixing ATP (364 mg), 0.1 M sodium barbital (48 ml), 0.18 M calcium chloride (24 ml), and distilled water (72 ml): pH was controlled at 9.4 by adding sodium hydroxide or hydrochloric acid. The third environment for incubation was an acid solution, which is an acetate buffer from the previous section with pH 4.3 (or 4.6). Calcium chloride (240 ml), 0.1 M sodium barbital (240 ml), cobalt chloride (4.8 g per 240 ml water), and ammonium sulfide (0.5%) were placed in a dyeing glass at room temperature. The painted sample was washed with distilled water and placed on a slide for further investigation.

Works Cited

Glaser, Jennifer, and Masatoshi Suzuki. “Skeletal Muscle Fiber Types in Neuromuscular Diseases.” Intech Open, 2018, Web.

Gondret, Florence, et al. “Low Birth Weight is Associated with Enlarged Muscle Fiber Area and Impaired Meat Tenderness of the Longissimus Muscle in Pigs.” Journal of Animal Science, vol. 84, no. 1, 2006, pp. 93-103.

Handel, S. E., and N. C. Stickland. “Muscle Cellularity and Birth Weight.” Animal Science, vol. 44, no. 2, 1987, pp. 311-317.

Kim, Gap-Don, et al. “Relationship Between Pork Quality and Characteristics of Muscle Fibers Classified by the Distribution of Myosin Heavy Chain Isoforms.” Journal of Animal Science, vol. 91, no. 11, 2013, pp. 5525-5534.

Ritchie, Hannah, and Max Roser. “Meat and Dairy Production.” Our World in Data, 2017, Web.

Thornton, Alex. “This is How Many Animals We Eat Each Year.” World Economic Forum, 2019, Web.