250 0 C:\BOOK\CHAP2.W51 June 20, 1995

CHAPTER 2 - CONNECTIVE TISSUES OF THE CARCASS

INTRODUCTION

The animal body is supportd by bone, is held together by fibrous connective tissue, and is protected against starvation and cold by adipose tissue. These three types of tissue, although they differ radically in their appearance and properties, are all classified as types of connective tissue. All three types contain cells located in a matrix that contains fibers.

In bones, both the matrix and the fibers make an important contribution to mechanical strength. The hardness of a bone originates from its calcified matrix, but its stength comes from embedded collagen fibers. The cells of bone, osteocytes, are trapped in small caves called lacunae. The gristle of the carcass is formed from tendons (by which muscles pull on bones), from ligaments (which hold bones together at the joints of the skeleton), from aponeuroses (which cover some muscles) and from fasciae (which form strong sheets between muscles). The dominant protein in gristle is collagen. Since connective tissues permeate nearly all parts of the body at the microscopic level, collagen is the most abundant protein in the animal body. The collagen fibers in meat are converted from strong fibers to jelly (gelatin) by the action of moist heat during cooking. The collagen in bones may be removed by mild hydrolysis to produce gelatin for use in other food products or for other uses such as photographic emulsions. In fat or adipose tissue there is little trace of the matrix and fibers, but the cells are bloated with stored triglyceride, the chemical component of lard.

This chapter starts with a general description of the bones that occur in the carcasses of meat animals and poultry - information that will be used later in the book to describe the anatomical locations of muscles and cuts of meat. After surveying the macroscopic structure of the skeleton, the level of magnification is increased to consider the microstructure and development of the dominant types of connective tissue in the carcass - gristle, bone and fat.

THE STRUCTURE OF THE SKELETON

Many parts of the skeleton may be seen or handled through the skin of the live animal (Figure 2-1, 2-2 and 2-3). These points of conformation are important in showing and judging animals. When comparing the conformation of a live animal with the structure of its carcass, it is important to remember that the hindlimb is normally rotated through an angle of nearly ninety degrees when the carcass is suspended in the abattoir. Thus, the muscles on the posterior face of the hindlimb bulge outwards, while the muscles on the anterior face of the hindlimb and in the belly are stretched.

Skull

The brain is situated within the cranium - a box-like posterior part of the skull. The brain is connected to the spinal cord through a large hole, the foramen magnum. The foramen magnum is flanked by two large knobs or occipital condyles that form a joint with the first cervical vertebra of the neck. The skulls of meat animals are damaged in the frontal bone region if animals have been stunned by concussion. Sinuses or spaces are present between the inner and outer cranial walls. There are considerable differences in the size and shape of the skull in different breeds of farm mammals, particularly in pigs where a long narrow skull is a feature that is often associated with a relatively large amount of fat in the carcass (Gregory and Whelehan, 1983).

The major muscles used in chewing are attached to the coronoid process which is a large expansion of the lower jaw or mandible. The coronoid process is located medially to the zygomatic arch, between the eye and the ear (Figure 2-4). The coronoid process allows muscle leverage to be exerted onto the mandible. The joint between the skull and the lower jaw is formed by a mandibular condyle. In cattle and sheep, the mandibular condyle is relatively flat and allows considerable movement in a horizontal plane. Lateral movement is important in animals whose teeth work with a grinding action.

The jig-saw pattern of suture joints on the skull surface indicates that the whole skull is formed by the fusion of a number of individual bones. A saw cut made transversely through the facial region of the skull reveals delicate rolls of the turbinate bones in the nasal cavity. The turbinate bones support a large area of nasal epithelium to warm and moisten the air travelling to the lungs, and to provide a large area for the sense of smell.

Skeleton of the neck

The vertebral column or backbone is the main axis of the skeleton and it protects the spinal cord. The spinal cord is located in a neural canal formed by a long series of neural arches, each contributed by a different vertebra (Figure 2-5). The neural arch of each vertebra is supported on the body or centrum of the vertebra. In some types of vertebrae, the neural arch extends dorsally as a prominent spine that may be called a dorsal spine, a neural spine or a spinous process. Where movement between vertebrae is possible, the centra are separated by cartilaginous intervertebral discs. In mammals, the anterior and posterior faces of the centra are almost flat. The names and numbers of the different types of vertebrae in meat animals are shown in Table 2-1.

Table 2-1. Numbers of vertebrae in meat animals (from Sisson and Grossman, 1953; Shaw, 1929; Palsson, 1940).

--------------------------------------------------------------- NAME REGION BEEF PORK LAMB ---------------------------------------------------------------

Cervical Neck 7 7 7 Thoracic Ribcage 13 13 - 17 13 - 14 Lumbar Loin 6 5 - 7 6 - 7 Sacral Sirloin 5 4 4

Caudal Tail 18 - 20 20 - 23 16 - 18 ---------------------------------------------------------------

Meat animals, like most other living mammals, usually have seven cervical vertebrae in the neck region. However, sheep sometimes have only six cervical vertebrae (Palsson, 1940), and as few as five cervical vertebrae have been reported in pigs (Berge, 1948). In cattle and sheep, but to a lesser extent in pigs, the neck is very mobile, and the cervical vertebrae have a series of interlocking articular and tranverse processes that limit excessive bending of the neck to protect the spinal cord (Figure 2-6). The first cervical vertebra, the atlas, articulates with the skull and is greatly modified in shape to form a joint that enables the animal to nod its head up and down. Rotation or twisting of the head occurs from the joint between the atlas and the next cervical vertebra, the axis. The ligamentum nuchae is a very strong elastic ligament in the dorsal midline of the neck, and it relieves the animal of the weight of its head. Were it not for the ligamentum nuchae, the head of the standing animal would droop between its forelimbs.

The ligamentum nuchae is pale yellow with a thick cord-like or funicular part and a flat sheet-like or lamellar part (Figure 2-6). Once the head is removed at slaughter, the elasticity of the ligamentum nuchae causes the neck of the carcass to curve dorsally. Beef and pork carcasses usually are split into right and left sides soon after slaughter and the series of vertebral centra that are now exposed is called the chine bone.

Ribcage

The number of vertebrae in pork carcasses is rather variable, particularly in certain breeds (Shaw, 1929, 1930). The heritability of the number of vertebrae is about 0.74 (Berge, 1948). Each extra vertebra adds about 15 mm to the length of the carcasses at slaughter weight (King and Roberts, 1960). The number of thoracic vertebrae, each bearing left and right ribs, ranges from 13 to 17. Breeds with a large size when mature and with heavy bone development tend to have more thoracic vertebrae than lighter breeds. Sometimes the ribs on extra thoracic vertebrae are only partially formed, but usually they are complete. The minimum number of lumbar vertebrae is generally found in cacasses with the maximum number of thoracic vertebrae. However, the variability of vertebral numbers frequently leads to an increase in the total number of vertebrae, so that the phenomenon is not due simply to the substitution of one type of vertebra for another. Lamb carcasses usually have either 13 or 14 thoracic vertebrae and a corresponding number of pairs of ribs (Palsson, 1940).

Experimental studies on embryonic amphibians (Detwiler, 1934) suggest that the number of vertebrae in an animal is determined by the number of somites (see Chapter 6) that develop along the length of the spinal cord. By definition, in mammals, the vertebrae that bear ribs are identified as thoracic vertebrae. In the embryo there are ossification centers on each side of the developing vertebrae. In vertebrae that do not normally develop ribs, these lateral ossification centers contribute their bone tissue to the centra of adjacent vertebrae. In the thoracic vertebrae, however, this laterally derived bone tissue remains separate from the centra and forms the ribs. Thus, the numbers of pairs of ribs and the numbers of thoracic vertebrae are determined by the developmental mechanism that controls the fate of the tissue which is derived from the lateral ossification centers.

The cage formed by thoracic vertebrae, ribs and sternum is an essential component of the respiratory system. Thoracic vertebrae are distinguished by their tall dorsal spines, many of which point towards the hindquarter and are known as the feather bones. The ribs are joined to the vertebral column dorsally so that the head of each rib articulates with the bodies of two adjacent vertebrae. Each rib has a tubercle that articulates with the transverse process of the more posterior of its two vertebrae (Figure 2-7). Ventrally, the anterior ribs articulate with the sternum and are termed sternal ribs (Table 2-2). The more posterior ribs are called asternal ribs and they only connect to the sternum indirectly via costal cartilages. The most posterior ribs have only small costal cartilages that do not reach all the way to the sternum. Some of the costal cartilages are very hard and may appear more like bones than typical cartilage. The sternum is formed by a number of closely joined bones, the sternebrae. When split through the midline, the interior structure of the sternebrae resembles that found in the centra of the vertebrae. But in an isolated cut of meat, the distinction between sternebrae and vertebral centra may be made by the presence or absence of a neural canal. A neural canal is seen in the vertebrae of carcasses that have been symmetrically separated into right and left sides.

The structure of the ribcage is rather variable in lamb carcasses. Carcasses have been found with as few as 12 ribs on one side, and left and right sides of the ribcage may differ in their number of ribs (Palsson, 0941). In lambs, rib length is determined mainly by age while the plane of nutrition determines rib thickness (Palsson and Verges, 1965).

Table 2-2. Structure of the ribcage in meat animals.

-------------------------------------------------------------- BEEF PORK LAMB -------------------------------------------------------------- Total pairs of ribs 13 13 -17 13 - 14

Pairs of sternal ribs 8 7 8

Pairsof asternal ribs 5 7 - 8 5 - 6

Number of sternebrae 7 6 6 - 7 --------------------------------------------------------------

The skeleton of the loin, sirloin and rump

In a live animal, the lumbar vertebrae act like a suspension bridge to support the weight of the abdomen. The lumbar vertebrae have flat, wing-like transverse processes that broaden the abdominal cavity dorsally (Figure 2-8) to provide a strong attachment for the muscles of the abdominal wall that carry the weight of the viscera. The propulsive thrust generated by the hindlimb during locomotion is transmitted to the sacral vertebrae by the pelvis. To strengthen the sacral vertebrae, they are fused together to form the sacrum (Figure 2-9). Fusion is incomplete in young animals and provides an important clue to animal age in the dressed carcass.

The pelvis is formed by three bones on each side (Figure 2-10). The most anterior bone on each side is the ilium. The shaft of the ilium expands anteriorly to form a flat wing attached to the sacrum. This joint is called the slip joint. When seen in a sirloin steak, the ilium may appear either as a small round bone or a large flat bone. The anterior edges of the ilia form the hooks of the live animal (Figure 2-1). The most posterior bone of the pelvis on each side is the ischium. The pelvis and the sacrum form a ring of bone completed ventrally by the pubes (Figure 2-11). The left pubis is separated from the right pubis by fibrocartilage which, at parturition, may soften to allow movement between the bones of the pelvis. The pubes are separated when carcasses are split into left and right sides in the abattoir. The pubic bone exposed on a carcass is called the aitch bone. The aitch bone is curved in steer and bull carcasses, is moderately curved in heifers, but is straight in cow carcasses (Figure 2-12). Only two caudal or coccygeal tail vertebrae are left on a commercial beef carcass.

Forelimb skeleton

The most proximal bone of the forelimb is the blade bone or scapula. It is not fused to the vertebral column (like the pelvis in the hindlimb), and this allows muscles that hold the scapula to the ribcage to function as shock absorbers during locomotion. The scapula has a distal socket joint for the next bone in the forelimb, the humerus. This socket joint is called the glenoid cavity (Figure 2-13). The glenoid cavity is wide and shallow, unlike the ball and socket joint in the hindlimb which is narrow and deep. Along the dorsal edge of the scapula, the bone merges with flexible hyaline cartilage. On the lateral face of the scapula is a prominent ridge of bone called the spine of the scapula. In beef carcasses, the scapular spine is extended distally as a prominent acromion process.

Proceeding distally down the forelimb, the bone that articulates with the scapula is the humerus (Figure 2-14). Proximally, the humerus has a relatively flat knob or head to fit into the glenoid cavity of the scapula. Two well defined condyles on the distal end of the humerus contribute to the hinge joint at the elbow (Figure 2-15). The elbow is formed by the olecranon process, an extension of the ulna. The bones of the elbow may be sold by the butcher as shank knuckle bones, a rather misleading name if compared to the anatomy of the human knuckle bones. The radius is joined to the ulna and is the shorter and more anterior bone of the pair.

Beef and lamb carcasses have a set of six compact carpal bones remaining on the carcass after slaughter. Before slaughter, the forefeet of cattle and sheep have a large cannon bone located distally to the carpal bones. Beef cannon bones are removed with the feet at slaughter since there is virtually no meat on them. Cannon bones sometimes are left on lamb carcasses in the abattoir to prevent the meat contracting proximally up the limb. Each forelimb cannon bone in ruminants is derived by the enlargement and fusion of the third and fourth metacarpal bones.

In the human hand, the metacarpal bones lie in the flat part of the hand, between the wrist and the knuckles. If the human hand is placed flat on a desk and is slowly lifted from the wrist, the thumb (digit 1) is the first digit to leave the desk, followed by the index finger and the little finger (digits 2 and 5, respectively). The third and fourth fingers remain on the desk. This demonstrates how the feet of meat animals may have evolved from an original basic plan with five digits. In pigs, digits 3 and 4 on each foot bear most of the body weight and are larger than the lightly loaded digits 2 and 5 (Figure 2-16). The first digit is absent. The evolutionary trend towards lifting of the foot and reduction of digits is even more extensive in cattle and sheep. Cattle and sheep have cursorial limbs, long limbs adapted for running. Digits 2 and 5 are reduced to dew claws behind the fetlock. Weight-bearing digits 3 and 4 are enlarged, and their metacarpals are fused to form a long cannon bone (Figure 2-16). The small bones in the toes of both fore and hind feet are called phalanges.

The feet remain on pork carcasses when they are shipped from the abattoir in shipper's style (unsplit carcasses with head plus perirenal or leaf fat) or in packer's style (split sides with leaf fat and head but not jowls removed). However, in a Wiltshire side for bacon production, the feet are removed, together with the head, pelvis, vertebral column and psoas muscles.

Hindlimb skeleton

The proximal bone of the hindlimb is the femur or round bone (Figure 2-17). Its articular head is deeply rounded and it bears a round ligament that holds it into the acetabulum. Another distinctive feature of the femur is the broad groove between the two trochlear ridges located distally. The patella or knee cap slides in this groove. The tension generated by muscles above the knee is transmitted over the knee or stifle joint by the patella to avoid having an important tendon in a vulnerable position over the anterior edge of a joint.

In beef and lamb carcasses there is a single major bone, the tibia or shank bone, located distally to the femur. In the corresponding position in a pork carcass there are two parallel bones, a large tibia and a more slender fibula (Figure 2-18). The presence of parallel bones suggests that, at some point in an animal's evolutionary past, rotation of the limb about its axis was possible. For example, rotation of the human wrist involves a partial crossing of the widely spaced ulna and radius but limb rotation is reduced as animals develop cursorial limbs. In cattle and sheep, one of the parallel bones, the fibula, has lost its shaft. Only a remnant of the head of the fibula may be found. In pigs, the fibula retains its shaft and the bone is mobile at birth. After a few years, however, the fibula becomes fused to the tibia.

Distal to the tibia are the tarsal bones of the hock. The structure of the tarsals, metatarsals and phalanges of the hindlimb is similar to that of the carpals, metacarpals and phalanges in the forelimb. Pork carcasses normally are suspended by a gambrel or hooked bar placed under the tendons of the hind feet. Beef carcasses normally are suspended by a hook under the fibular tarsal bone. This bone projects posteriorly and has a rough knob, the tuber calcis, for the insertion of the Achilles tendon at the hock.

Poultry skeleton

The skeletons of poultry are radically different from those of the farm mammals. Not only is the avian skeleton adapted for flight, but birds and mammals are only distantly related zoologically. The skull has very large eye orbits and a small cranial cavity. The long double curved neck contains 14 cervical vertebrae, and the ring-like atlas articulates to the skull with only a single occipital condyle. The axis has a large odontoid process that projects anteriorly. There are 7 thoracic vertebrae, but numbers 2 to 5 are fused. Thoracic vertebrae 6 can move freely, but the last thoracic vertebra is fused to the synsacrum. The synsacrum is a fused length of the vertebral column that contains thoracic vertebra 7, 14 lumbo-sacral vertebrae, and the first coccygeal or caudal vertebra, but skeletal fusion in the vertebral column does not occur for many weeks after hatching (Hogg, 1982). There are six caudal vertebrae that, apart from the first, are free and mobile. However, only numbers 2 to 5 are normal vertebrae, since the last one is formed into a three sided pyramidal bone called the pygostyle.

There are seven ribs: the first two are free while the last five are attached to the sternum. There are no costal cartilages. Ribs 2 to 6 each have an uncinate process which overlaps the next posterior rib. The sternum is extremely large. It has a conspicuous ventral ridge in the midline, the CARINA, which increases the area available for attachment of the flight muscles. The dorsal surface of the expanded sternum is concave and forms the floor of a continuous thoracic and abdominal cavity.

The bones of the fore limb are greatly modified to form the wing (Figure 2-19). Distal to the humerus are the widely spaced radius and ulna. The carpals, metacarpals and digits are reduced to form a stiff skeletal unit for the anchorage of the primary flight feathers. The three digits of the wing are equivalent to digits 2, 3, and 4 in other animals (Montagna, 1945). The wing articulates with the body at the glenoid cavity which is strengthed by the convergence of three bones, the scapula, the coracoid and the clavicle. In birds the coracoid is a separate bone, whereas in mammals it has been reduced to a small integral part of the scapula. The clavicles of right and left sides are fused ventrally to form the furcula or wishbone. Although many mammals have a pair of clavicles, they are absent in cattle, sheep and pigs. The clavicle functions as a strut to support the shoulder joint in animals which have complete mobility of the shoulder joint. Since cattle, sheep and pigs have cursorial limbs with a restricted fore and aft movement, they do not need clavicles. In poultry, the distal end of the coracoid is braced against the sternum (Figure 2-20). In flight, the body of a bird hangs from its wings at the shoulder joint, hence the more elaborate support for the glenoid cavity.

In poultry, the legs show many cursorial adaptations. Distal to the femur, the fibula is reduced to leave the tibia as the major bone (Figure 2-21). In the embryo this occurs as a result of the differential growth and translocation of the distal part of the fibula to become part of the tibia (Archer et al., 1983). The proximal tarsal bones are fused to the distal end of the tibia to increase its length, and the whole skeletal unit may be called the tibiotarsus. The distal tarsal bones are incorporated into the proximal end of a single bone, the tarsometatarsus, which also includes the fused metatarsals 2, 3 and 4. Of the four digits which form the bird's claw, digit 1 is directed posteriorly while digits 2, 3 and 4 are anterior. This adpatation enables the bird to perch. The ilium is fused to the synsacrum. Instead of being fused in the midline, the pubic bones are separate, and they project backwards as thin rods (Figure 2-22). The open structure of the pelvis in the ventral region facilitates the passage of eggs from the body cavity. The ilium, ischium and pubis all contribute to the acetabulum, but the ilium forms more than half of the socket and the floor is membranous (Harrison, 1975).

THE MICROSTRUCTURE AND DEVELOPMENT OF CONNECTIVE FIBERS

Collagen fibers

Collagen is an elongated protein that forms extremely strong microscopic fibrils. Collagen fibrils may be bound together to form microscopic fibers which, in large numbers, may appear as gristle in raw meat. Collagen is the most abundant protein in the animal body, and the collagen which occurs in meat may be an important source of meat toughness. Beef carcasses have to be graded according to age (Chapter 4) mainly because of age-related changes in collagen that cause meat from older cattle to be tough. Large amounts of collagen are found in animal skin. In pig skin for example, collagen fibers are tightly woven from two directions so as to form a tight meshwork (Meyer et al., 1982). Collagen is a raw material for major industries in leather, glue and cosmetics.

Under a light microscope, collagen fibers in the connective tissue framework of meat range in diameter from 1 to 12 microns. They do not often branch and, when branches are found, they usually diverge at an acute angle. Collagen fibers from fresh meat are white, but usually they are stained in histological sections. Eosin gives them a pink color. Unstained collagen fibers may be seen by polarized light since they are birefringent (Pimental, 1981). By rotating the plane of polarized light, collagen fibers appear bright against an otherwise dark background. The birefringence of collagen fibers in meat is lost at the point during heating when gelatinization occurs (Figure 2-23; Swatland, 1989). Collagen fibers have a wavy or crimped appearance which disappears when they are placed under tension.

Collagen fibers fluoresce with a blue-white light when excited with UV (Swatland, 1987a) so that the amount of connective tissue on a cut meat surface may be measured very rapidly (Swatland, 1987b). Peak excitation is around 370 nm so that the prominent 365 nm peak emission of a mercury source may be used (Swatland, 1987c). Some indication of collagen fiber diameter may be obtained by spectrofluorometry since the fluorescence is quenched fairly rapidly. Thus, large fibers retain a core with a pre-quenching emission spectrum for longer than small fibers (Figure 2-24; Swatland, 1987d). Adipose tissue only fluoresces weakly, to about the same extent as areas of muscle with a low connective tissue content (Swatland, 1987e). Thus, as considered in more detail in Chapter 3, it is possible to make fiber-optic probe measurements of the connective tissue in meat. By a fortunate coincidence, collagen fluorescence increases with age (Odetti et al., 1992, 1994) so that probe measurements may have a promising future in beef grading.

Electron microscopy reveals that collagen fibers are composed of parallel bundles of small fibrils with diameters ranging from 20 to 100 nm. Collagen fibrils typically have diameters which are multiples of 8 nm and may reflect the manner in which they grow radially (Craig and Parry, 1981). Gotoh et al. (1983) have proposed that collagen fibers are composed of a slightly helical coil of 11 nm microfibrils surrounded by a banded sheath. Collagen microfibrils may appear to have a tubular structure with an electron-lucent lumen. Inoue and Leblond (1986) suggested that the surface bands around the tubules may secure the string of segments from which microfibrils appear to be formed.

Collagen fibrils are formed from long molecules of tropocollagen which are staggered in arrangement but tightly bound laterally by covalent bonds (Figure 2-25). When negatively stained with heavy metals that spread into the spaces between the ends of molecules, collagen fibrils appear to be transversely striated. The periodicity of these striations is 67 nm but often shrinks to 64 nm as samples are processed for examination. There is considerably more detail to the periodicity of the transverse striations than is indicated in Figure 2-25 and negative staining techniques may be used to relate the periodicities of staining to the molecular structure of tropocollagen (Tzaphlidou, 1986). The initial stages of collagen fibril assembly may be intracellular with fibril morphology being regulated by a special site on the fibroblast membrane (Trelstad and Hayashi, 1979).

Tropocollagen molecules

Tropocollagen is a high molecular weight protein (300,000) formed from three polypeptide strands twisted into a triple helix. Each strand is a left-handed helix twisted on itself, but this is not shown in Figure 2-26 where only the larger right-handed triple helix which involves all three polypeptide strands is shown. The triple helix is responsible for the stability of the molecule and for the property of self assembly of molecules into microfibrils. The flexible parts of each strand projecting beyond the triple helix (telopeptides) are responsible for the bonding between adjacent molecules (Kuhn and Glanville, 1980). In other words, the cross links that bind tropocollagen molecules together laterally are made between the helical shaft of one molecule and the non-helical extension of an adjacent molecule.

In the polypeptide strands, glycine occurs at every third position, and proline and hydroxyproline account for 23% of the total residues. The regular distribution of glycine is required for the packing of tropocollagen molecules and has been claimed as evidence that all animals are derived by evolution from a single ancestral stock, since the chance development of this unique regularity in unrelated animals is thought unlikely (Finerty, 1981). Since hydroxyproline is quite rare in other proteins of the body, an assay for this imino acid provides a measure of the collagen or connective tissue content in a meat sample (O'Neill et al., 1979). Tropocollagen also contains a fairly high proportion of glutamic acid and alanine as well as some hydroxylysine. Methods for the detection and estimation of the collagen content in meat products are reviewed by Etherington and Sims (1981).

Types of collagen

Each tropocollagen molecule is composed of three alpha chains but 19 unique alpha chains have been identified, giving rise to 11 different types of collagen. These have been divided by Miller (1985) into three general classes:

(1) molecules with a long (about 300 nm) uninterupted helical domain,

(2) molecules with a long (300 nm or greater) interupted helical domain, and

(3) short molecules with either a continuous or an interupted helical domain.

Various types of collagen of interest in understanding the structure of meat animals are detailed in Table 2-3. Other types are found in locations such as placental villi (Type VI), placental membranes (Type VII), endothelial cells (Type VIII), and hyaline cartilage (Types IX, X and K). Different types of collagen can be identified histochemically as well as biochemically (Bock, 1977, 1978).

Table 2-3. Derivation of different types of collagen from different combinations of alpha chains (from Gay and Miller, 1978, Bailey and Sims, 1977, Burgeson et al., 1976, Bentz et al., 1978, and Bailey et al., 1979).

------------------------------------------------------------------

Types of Types of alpha chains

Collagen ------------------------------------------------------

    1I1II1III1IV2    A    B -----------------------
-------------------------------------------

  I        2								1 

II 3

III 3

IV 3

V 1 2

------------------------------------------------------------------

Type I collagen forms striated fibers between 80 and 160 nm in diameter in blood vessel walls, tendon, bone, skin and meat. It may be synthesized by fibroblasts, smooth muscle cells and osteoblasts.

Type II collagen fibers are less than 80 nm in diameter and occur in hyaline cartilage and in intervertebral discs. It is synthesized by chondrocytes.

Type III collagen forms reticular fibers in tissues with some degree of elasticity, such as spleen, aorta and muscle. It is synthesized by fibroblasts and smooth muscle cells, contributes substantially to the endomysial connective tissues around individual muscle fibers, provides a small fraction of the collagen found in skin (Ramshaw, 1986) and occurs in the large collagen fibers dominated by Type I collagen. It may have some function regulating collagen fiber growth (Keene et al., 1987).

Type IV collagen occurs in the basement membranes around many types of cells and may be produced by the cells themselves, rather than by fibroblasts (Laurie et al., 1980). Although basement membranes were once regarded as amorphous, many of them now are thought to be composed of a network of irregular cords (Inoue and Leblond, 1988). The cords contain an axial filament of Type IV collagen, ribbons of heparin sulfate proteoglycan, and fluffy material (laminin, entactin and fibronectin). Type IV collagen occurs in the endomysium around individual muscle fibers (Light and Champion, 1984). Instead of being arranged in a staggered array (as shown in Figure 2-25), the molecules are linked at their ends to form a loose diagonal lattice (Martin et al., 1985).

Type V collagen is found prenatally in basement membranes and cultures of embryonic cells. It is synthesized by myoblasts, smooth muscle cells and, possibly, by fibroblasts (Kuhn and Glanville, 1980). Type V collagen is composed of -A and -B chains (Burgeson et al., 1976) in a 1:2 ratio (Bentz et al., 1978). Type V collagen has also been found in the basement membranes of muscle fibers, except at the point where muscle fibers are innervated (Sanes and Cheney, 1982).

Type VI collagen is a tetramer of Type VI. It forms a filamentous network and has been identified in muscle and skin (Keene et al., 1988). The molecule consists of a short triple helix about 105 nm in length with a large globular domain at each end.

Tendons often extend into the belly of a muscle or along its surface before they merge with its connective tissue framework, and types I and III collagen both may be extracted from meat. Even within tendons, there may be some Type III collagen forming the endotendineum or fine sheath around bundles of collagen fibrils (Duance et al., 1977). In fibers composed of collagen Types I and II, fibrils have a straight arrangement whereas, in fibers of Type III collagen, the fibrils have a helicoidal arrangement (Reale et al., 1981).

Small diameter type III collagen fibers are called reticular fibers since, when stained with silver for light microscopy, they often appear as a network or reticulum of fine fibers. The larger diameter collagen fibers formed from Type I collagen are not blackened by silver. The identification of reticular fibers by silver staining has a long and complex history which is reviewed by Puchtler and Waldrop (l978).

Collagen fibers shrink when they are placed in hot water, and they are ultimately converted to gelatin. Around 65oC, the triple helix is disrupted and the alpha chains fall into a random arrangement. The importance of this change is that it tenderizes meat with a high connective tissue content. Tropocollagen molecules from older animals are more resistant to heat disruption than those from younger animals. In early studies, it was suggested that reticular fibers, unlike collagen fibers, did not yield gelatin when treated with moist heat. The original suggestion that reticular fibers survive unchanged after cooking is wrong, but a modification of the idea is plausible. Since a piece of meat may contain different types of collagen, and since these types may differ in the thermal stability of their cross links, it is possible that, at an intermediate level of cooking around 65oC, endomysial collagen and perimysial collagen may differ in the extent to which they are affected by the cooking treatment (Bailey and Sims, l977). Heat-induced solubilization of Type I collagen is more important in improving meat tenderness by cooking than is the effect of heat on Type III collagen (Burson and Hunt, 1986a).

Collagen biosynthesis

The synthesis of the different polypeptide strands that are combined to make different types of collagen is genetically regulated by the production of messenger RNA. The synthesis of polypeptide strands occurs on membrane-bounded polysomes, but the hydroxylation of lysine and proline occurs after the strands are assembled. Ascorbic acid is required for the hydroxylation of lysine and proline. Polypeptide strands enter the cisternae of the endoplasmic reticulum, the terminal extensions of the strands are aligned, and then the strands spiral around each other. Procollagen or immature collagen has long terminal extensions protruding from each end of the newly formed triple helix. Procollagen moves to the golgi apparatus and is packaged into vesicles that are moved to the cell surface, probably by microtubules. Except for some Type III procollagen molecules, the long terminal extensions are then enzymatically reduced in length.

Outside the cell, collagen molecules become aligned in parallel formations, and then they link up laterally to form fibrils. It is likely that tropocollagen monomers are partially assembled together in groups before they are added to an existing collagen fibril (Trelstad, 1982). Firstly, vacuoles containing procollagen fuse to form a fibril-containing compartment. Then the cytoplasmic extensions withdraw from between several fibril-forming compartments to create a bundle-forming compartment (Birk and Trelstad, 1986; Figure 2-27). Sometimes collagen fibrils occur intracellularly, but it is not clear whether this is collagen taken up by phagocytosis or a surplus of newly synthesized collagen (Michna, 1988).

The characteristic parallel staggered arrangement of tropocollagen molecules in a collagen fibril is caused by the 67 nm repeating pattern of oppositely charged amino acids along the length of the tropocollagen molecule (Miller, 1982). The degree of overlapping of adjacent molecules and the gaps left between the ends of molecules cause the striated appearance of collagen fibers seen by electron microscopy (Figure 2-25). The fibroblasts of young animals are metabolically more active than those of older animals, particularly for aerobic metabolism (Floridi et al., 1981).

Accumulation of collagen in meat

Although the relative proportions of Types I and III collagen in a muscle may be related to meat tenderness (Burson and Hunt, 1986b), the overall amount of collagen and its degree of crosslinking also are important. The absolute amount of collagen in an animal may increases as animals become older, and this may have an effect on meat toughness, but rapid growth of muscle fibers also may dilute the relative amounts of collagen in meat. Considering the supposed importance of collagen in meat toughness, the absence of overwhelming evidence from taste panel studies is rather curious. It seems reasonable that stewing beef is tougher than prime steak because it has more collagen, but is collagen responsible for differences in tenderness between the same cut of steak from different commercial carcasses? Some of this uncertainty may originate from the fact that collagen concentration may be the primary determinant of eating quality, while collagen solubility may determine shear strength measured mechanically (Young and Braggins, 1993; Young et al., 1993).

Recent research using a UV fiber-optic probe for connective tissues in meat is starting to shed new light on this old problem. The dominant change in beef animals from 12 to 17 months is an increase in the incidence of fluorescence peaks as layers of perimysium grow to became detectable. From 17 to 24 months, the incidence of peaks decreases as layers of perimysium are pushed apart by muscle fiber growth. Fluorescence peaks became wider from 17 to 24 months as layers of perimysium grow in thickness. The effects of connective tissue on the tenderness of cooked meat evaluated by taste panels tend to be more important as animals grow older (24 months) and in cuts such as round steak that have an intermediate level of tenderness.

Within a carcass, there are considerable differences in collagen content between different muscles and this is reflected in their retail price (Chapter 4). Collagen content also may differ between sexes. For example, the hydroxyproline content is higher in pork from females than castrated males (Sellier and Boccard, 1971). However, the amount of collagen in meat, when expressed as a proportion of wet sample weight, also is affected by fat content. In steaks from a veal carcass, for example, the collagen content might exceed 0.5%, but could be much less in the same region from a steer carcass in which fat had accumulated to "dilute" the collagen content.

Collagen in meat may be studied by measuring collagen fibril diameters in electron micrographs (Rowe, l978). In equine tendons, fibril diameters in the fetus are unimodal but become bimodal in the adult (Parry et al., l978b). Large diameter fibrils may have more intrafibrillar covalent cross links, while small diameter fibrils may have more interfibrillar non-covalent cross links. Thus, fibril diameter may be related to fibril strength and elasticity (Parry et al., l978a). Meat with large diameter collagen fibers tends to be tougher than meat with thinner collagen fibers (Light et al., 1985).

Little is known about the mechanisms by which collagen fibers become arranged in a muscle, or about the interactions which occur between fibroblasts and the fibers that they produce, although it is possible that glycosaminoglycans play some part in this interaction (Parry et al., l978a).

Collagen is very important in muscle development. Myoblasts, the cells that form muscle fibers, develop a parallel alignment when cultured on a substrate of Type I collagen, but they do not become elongated or aligned on Type V basement membrane collagen (John and Lawson, 1980). Myoblasts may themselves form Types I, III and V collagen (Sasse et al., 1981), while myotubes also may form collagen (Mayne and Strahs, 1974) but, perhaps, only when associated with fibroblasts (Lipton, 1977). The identification of collagen in developing muscle is complicated by the fact that the tail unit of the acetylcholinesterase molecule has a collagen-like sequence that contains hydroxyproline and hydroxylysine (Lwebuga-Mukasa et al., 1976).

Crosslinking of collagen molecules

Within an individual collagen molecule, the three polypeptide strands are linked together by stable intramolecular bonds that originate in the non-helical ends of the molecule. The great strength of collagen fibers, however, originates mainly from the stable intermolecular covalent bonds between adjacent tropocollagen molecules. Stable disulfide bonds between cystine molecules in the triple helix also occur. During the growth and development of meat animals, covalent cross links increase in number, and collagen fibers become progressively stronger. Meat from older animals, therefore, tends to be tougher than meat from the same region of carcasses from younger animals. This relationship is complicated in young animals by the rapid synthesis of large amounts of new collagen. New collagen has fewer cross links so that, if there is a high proportion of new collagen, the mean degree of cross linking may be low, even though all existing molecules are developing new cross links. As the formation of new collagen slows down, the mean degree of cross linking increases. Another complication is that many of the intermolecular cross links in young animals are reducible - the collagen is strong but is fairly soluble. In older animals, reducible cross links are probably converted to non-reducible cross links - the collagen is strong but is far less soluble and more resistant to moist heat. The chemistry of these changes is still a subject for debate.

Nakano et al. (1985) have suggested that pyridinoline, a non-reducible cross-link, may be involved in the increased heat stability of epimysial connective tissues from older animals. Although changes in collagen solubility might be an important factor affecting the tenderness of beef from older animals, the effect in younger animals at a typical commercial slaughter weight may be relatively slight (Hall and Hunt, 1982). With increasing USDA maturity levels, however, the pyridinoline content and thermal stability of intramuscular collagen both increase (Smith and Judge, 1991).

Differences in the degree of cross linking may occur between different muscles of the same carcass, and between the same muscle in different species. For example, collagen from the longissimus dorsi is less cross-linked than collagen from the semimembranosus, and collagen from the longissimus dorsi of a pork carcass is less cross-linked than collagen from the bovine longissimus dorsi. Nutritional factors such as high-carbohydrate diet, fructose instead of glucose in the diet, low protein, and pre-slaughter feed restriction may reduce the proportion of stable cross links. Nonenzymic glycosylation (between lysine and reducing sugars) may be involved in the interaction between diet and collagen strength (Furth, 1988). In general, the turnover rate of collagen is accelerated in catle fed a high energy diet (Wu et al., 1981). The rate of collagen turnover in skeletal muscle may be about 10% per day (Laurent, 1982) and the turnover time for collagen may be inversely proportional to collagen fibril diameter (Svoboda et al., 1983).

Elastin and elastic fibers

Individual collagen fibers only lengthen by about 5% when stretched and little elasticity is possible where collagen is formed into cable-like tendons. However, much of the collagen that is present in meat forms a meshwork so that stretching of the whole meshwork is possible because its configuration changes. Fibers with truly elastic properties, however, are necessary in structures such as the ligamentum nuchae and the abdominal wall. All arteries, from the aorta down to the finest microscopic arterioles, rely on elastin fibers to accommodate the surge of blood from contraction of the heart. Elastin fibers may be stretched to several times their original length but rapidly resume their original length once released. Elastin is found in all vertebrates except primitive jawless fish, and in evolution it appeared first in cartilagenous fish (Sage, 1983).

Elastic fibers are made of the protein elastin. Elastin resists severe chemical conditions, such as the extremes of alkalinity, acidity and heat that destroy collagen. Fortunately, there are relatively few elastic fibers in muscle, otherwise cooking would do little to reduce meat toughness. The elastin fibers in muscles that are used frequently for locomotion are larger and more numerous than those of less frequently used muscles (Hiner et al., 1955). Elastin fibers in the epimysium and perimysium of beef muscles range from 1 to 10 microns in diameter (Rowe, 1986). Elastin is synthesized by arterial smooth muscle cells, but the origin of elastin in non-vascular locations is not properly understood. In the lung, for example, large amounts of elastin are synthesized by various types of lung cells (Mandl et al., 1986) but the cellular source of the elastin fibers in meat is unclear at present. Some elastic fibers in muscle are involved in the attachment of sensory organs called neuromuscular spindles (Cooper and Gladden, l974). During the digestion of meat in the human gut, elastic fibers are broken down by ELASTASE, an enzyme from the pancreas that would not be there if our evolutionary ancestors had not been at least partly carnivorous.

Elastic fibers are pale yellow, as seen in the ligamentum nuchae, but may be stained for light microscopy by a variety of techniques (Munz and Meves, 1974; Brissie et al., 1975; Puchtler et al., 1973, 1976; Gotta-Pereira et al., 1976). When elastic fibers are stretched, they may become visible in polarized light without staining, but this requires careful attention to the refractive index of the mounting medium. In the bovine ligamentum nuchae, the pattern of birefringence indicates that there are two micellar structures, one arranged circularly on the outside and the other arranged axially in the centers of the fibers (Romhanyi, 1983). Elastic fibers in meat have a small diameter (aproximately 0.2 to 5 microns) although they are much larger in the ligamentum nuchae. Elastic fibers in the connective tissue framework of meat are usually branched.

Electron microscopy reveals that elastic fibers are composed of bundles of small fibrils approximately 11 nm in diameter embedded in an amorphous material. In the bovine ligamentum nuchae, fibrils may be constructed from smaller units or filaments approximately 2.5 nm in diameter (Serafini-Fracassini et al., 1978). Elastin filaments are bound by non-covalent interactions to form a three-dimensional network (Rucker and Lefevre, 1980). Elastic fibers are assembled in grooves on the fibroblast surface where initially rope-like aggregations of fibrils become infiltrated with amorphous elastin (Ross and Bornstein, 1971). Unlike the situation in elastic ligaments, where elastin forms fibers, the elastin of the arterial system occurs in sheets that condense extracellulerarly in the absence of fibrils.

Although elastin resembles tropocollagen in having a large amount of glycine, it is distinguished by the presence of two unusual amino acids, desmosine and isodesmosine (Partridge, 1966). Like collagen, elastin contains hydroxyproline, although it may not have the same function of stabilizing the molecule. Tropoelastin, the soluble precursor molecule of elastin (molecular weight 70,000 to 75,000), is secreted by fibroblasts after it has been synthesized by ribosomes of the rough endoplasmic reticulum and processed by the Golgi apparatus. In the presence of copper, lysyl oxidase links together four lysine molecules to form a desmosine molecule. Isodesmosine is the isomer of desmosine. The aorta may be fatally weakened by a lack of mature elastin in animals deprived of dietary copper. Elastin in the arterial system is produced by smooth muscle cells instead of fibroblasts.

The functional properties of elastin in different tissues such as lung and aorta may be related to differences in the ratio of tropoelastin A to B (Barrineau et al., 1981). The elastin of elastic cartilage might be a different genetic type to that found in the vascular system but, overall, the diversity of different genetic types of elastin is far less than for collagen.

THE CELLS OF FIBROUS CONNECTIVE TISSUE

The dominant cell type in the fibrous connective tissue of meat is the fibroblast, but other cells also exist. Macrophages or histioicytes are sometimes quite numerous and, when inactive, may resemble fibroblasts in appearance. However, the motility of macrophages is soon revealed by tissue inflammation or the injection of colloidal dyes. Macrophages migrate through the tissue and act as scavengers by engulfing invasive microorganisms or foreign particles by phagocytosis.

Cells from the vascular system may wander through connective tissues and even compact structures such as tendons have their own lymphatic and vascular supply (Edwards, 1946), something that is not easily seen in an exsanguinated carcass. The vascular cells include a variety of lymphocytes and the plasma cells responsible for antibody production. Eosinophils are cells with bilobed nuclei and numerous cytoplasmic granules readily stained by eosin. The skeletal muscles of cattle, and sometimes sheep (Harcourt and Bradley, 1973), may become inundated with eosinophils (eosinophilic myositis). The affected areas appear as irregular pale lesions and often are detected by meat inspectors looking for muscle parasites. Eosinophils may be attracted to areas of antibody activity and eosinophilic myositis may be an allergic response (Oghiso et al., 1977).

Mast cells (Selye, 1965) also occur within the skeletal muscles of meat animals (Swatland, 1978), mainly in the perimysium and epimysium. The numbers of mast cells may be increased in pathological situations (Helliwell et al., 1990) and, in denervated muscle, mast cells may move from the central tendon into the belly of the muscle (Sánchez-Mejorada and Alonso de Florida, 1992). The cytoplasm of mast cells contains large numbers of metachromatic granules. Metachromasia is a color change of dyes such as methylene blue so that metachromatic granules are purple while the surrounding tissue is blue. Mast cells contain heparin and histamine. Heparin prevents the coagulation of blood and histamine increases the permeability of small blood vessels. Heparin also activates the enzyme lipoprotein lipase involved in the accumulation of triglyceride by adipose cells. Mast cells also may release a substance that activates mitosis in nearby cells (Franzen and Norrby, 1980). Thus, the development of intramuscular fat in meat may have some relationship to the distribution of mast cells. Mast cells sometimes come into close contact with skeletal muscle fibers (Heine and Forster, 1974), but most mast cells are located along fine branches of the lymphatic system in the perimysium and endomysium (Stingl and Stembera, 1974). Mast cells also have been implicated in the regulation of collagenase activity (Simpson and Taylor, 1974).

CARTILAGE

Cartilage cells or chondrocytes occupy lacunae in a stiff flexible matrix formed from collagen fibers in a proteoglycan ground substance. Hyaline cartilage has a white translucent appearance and occurs on the smooth surfaces of joints. In the larynx, trachea and bronchi, hyaline cartilage forms the rings that hold these air ducts open during respiration. Flexible units of the skeleton, such as the dorsal part of the scapula and the linkages between the sternum and the posterior ribs, also are formed from hyaline cartilage. Most of the bones of the carcass are initiated prenatally as cartilagenous models that subsequently become ossified. Complete ossification is a slow process, and the bones of young meat animals are more flexible than those of adults. The state of ossification is a useful clue to animal age in carcass grading.

Chondrocytes are derived from mesenchymal cells and are initially capable of both mitosis and matrix formation. Clusters of related cells are pushed apart by their new matrix in a process called interstitial growth (Figure 2-28). Cartilaginous models of prenatal bones are covered by a membrane known as the perichondrium. Inner perichondrial cells differentiate into chondrocytes so that, in addition to interstitial growth, new cells and matrix may be added superficially in a process known as appositional growth (Figure 2-29). Cartilage may acquire numerous elastic or collagen fibers to become elastic cartilage or fibrocartilage, respectively. The dominant type of collagen in hyaline cartilage is Type II and accounts for 50 to 70% of dry weight. Cartilage also contains some unusual minor types of collagen. Type M collagen, for example, is much shorter than other collagen molecules, has a helix that is stabilized by disulfide bonds, and occurs in the matrix immediately around chondrocytes (Duance et al., 1984).

There are a number of histological features of cartilage that indicate an animal's physiological age.

(1) With age, the matrix becomes increasingly rigid.

(2) Appositional growth is the dominant growth process in older animals.

(3) Young chondrocytes are flattened or elliptical with their long axis parallel to the surface of the cartilage.

(4) Old chondrocytes are large (up to 40 micrometres diameter) and rounded in shape.

(5) In older animals, chondrocytes form nests or isogenous groups within single lacunae.

(6) With age, the basophilia of the matrix decreases.

(6) Relative to young chondrocytes, older chondrocytes have less endoplasmic reticulum, less extensive Golgi apparatus, and more stored glycogen.

THE MICROSTRUCTURE AND DEVELOPMENT OF BONE

Oxygen, nutrients and waste products may travel to and from the chondrocytes in cartilage by diffusion through the surrounding matrix but, when the matrix becomes ossified by the deposition of submicroscopic hydroxyapatite crystals, diffusion is greatly reduced. In bone, osteocytes can only survive if they develop long cytoplasmic extensions radiating from the lacunae to regions where exchange by diffusion can take place. These cytoplasmic extensions run through fine tubes or canaliculi in the ossified matrix, but are limited in length. Consequently, large numbers of blood vessels permeate the matrix of bone. Most of these blood vessels run longitudinally through the bone in large haversian canals surrounded by concentric rings of osteocytes and bone lamellae (Figure 2-30). Bones are covered by a connective tissue membrane called the periosteum.

The prenatal formation of bone is initiated by either of two basic mechanisms, (1) intramembranous ossification or (2) endochondral ossification. Intramembramous ossification is typical of the bones that form the vault of the skull, and it occurs when sheets of connective tissue produce osteoblasts which then initiate centers of ossification. Endochondral ossification is more common, and is the process by which cartilagenous models become ossified to form the bones of a commercial meat carcass.

The internal structure of carcass bones becomes visible when they are split longitudinally on a band saw (Figure 2-31). The shaft of a bone is called the diaphysis. The knob at each end of a bone is called the epiphysis. Between the diaphysis and each epiphysis is a cartilagenous growth plate called the epiphyseal plate. In a young animal, the chondrocytes of the epiphyseal plate are constantly dividing to form new matrix. However, on each face of the plate, cartilage is continuously resorbed and is replaced by bone (Howlett, 1980) so that the thickness of the epiphyseal plate tends to remain constant in growing animals. This process allows a bone to grow longitudinally without disrupting the articular surface on the epiphysis. The rate of the longitudinal growth of bones is the product of two factors; (1) the rate of production of new cells, and (2) the size that cells reach before they degenerate at the point of ossification (Thorngren and Hansson, 1981). The strength and thickness of epiphyseal plates is modified by sex hormones (Oka et al., 1979). At puberty, chondrocyte growth slows down and fails to keep pace with ossification on the surface of the epiphyseal plate. Thus, epiphyseal plates are lost in mature animals, and the epiphyses become firmly ossified to their diaphyses. However, the factors that regulate the closure of the epiphyseal plate and, hence, the frame size of the animal, are poorly understand. Although regulation is likely to be an interaction between animal age and circulating hormones, there are no obvious hormonal changes when the plate closes (Oberbauer et al., 1989). If whethers are implanted with estradiol the ossification of growth plates is accelerated (Field et al., 1990). Bone growth in mature animals is restricted to the girth or thickness of the bone, and it occurs by the recruitment of periosteal cells to become osteoblasts.

Calcium, calcification and bone resorption

Radioactive calcium (45Ca) may be used to study the uptake of calcium by the skeleton. There is a continuous exchange of calcium between the body fluids (mostly plasma) and approximately 1% of the total bone matrix with direct access to the circulating fluid. Growth by accretion results from small net gains to the matrix (Figure 2-32). When radioactive calcium is injected into a growing animal, the isotope is incorporated into new bone, and the concentration of isotope in the plasma declines (Wasserman, 1977). Both calcium exchange and calcium accretion are more rapid in the epiphysis than in the diaphysis.

Calcium, phosphate and hydroxyl ions are obtained from the extracellular fluid during bone formation. The first stage in ossification is the deposition of a crystal of calcium phosphate, then calcium phosphate is converted to hydroxyapatite. Two histological events associated with ossification have been observed. Firstly, extracellular membrane-bounded vesicles that contain phosphatase begin to accumulate calcium phosphate and hydroxyapatite. Secondly, in vivo crystal nucleation occurs between the ends of tropocollagen molecules.

The supply of calcium and phosphate from the blood is affected by vitamin D. Vitamin D is obtained from the diet or by exposure to ultraviolet light. It is hydroxylated in the liver and then converted to the hormonal form (1,25-dihydroxycholecalciferol) in the kidney. The hormonal form causes the intestine to increase its absorption of calcium and phosphate. The mechanical strength of bone may be used to assess the nutritional availability of minerals (Crenshaw et al., 1981).

The resorption of bone enables bone remodelling in response to local stresses and is coupled with the maintenance of blood calcium and phosphorus levels (Figure 2-33). The organic components of bone are degraded by the lysosomal enzymes of osteoclasts, a process that requires vitamin A. The solubilization of hydroxyapatite in response to parathyroid hormone is probably achieved by a combination of low pH (due to anaerobic glycolysis) and chelation. Bone resorption is inhibited by calcitonin (thyrocalcitonin).

Genetic defects in calcification and bone formation may occur in cattle. Typical signs are lack of calcification of the teeth and hypermobility of joints (Johnston and Young, 1958). In milk fever of cows (parturient paresis), paralysis and unconciousness may occur during parturition or early lactation. The condition is a result of drastically lowered plasma calcium and inorganic phosphorus levels, and may be treated by the injection of calcium borogluconate.

Regulation of skeletal growth

Carcasses from young animals have a relatively high bone content because the skeleton is well developed at birth. As an animal grows to market weight, its proportion of bone decreases on a relative basis, because of the growth of muscle and fat. The long-term control of bone growth is superimposed on the short-term regulation of bone metabolism that occurs in response to changes in blood calcium levels (Figure 2-33), or to remodelling in response to local functional demands.

A number of hormones exert secondary effects on skeletal development. Thyroxine, insulin, growth hormone and gonadal hormones tend to be anabolic. Estrogens may inhibit resorption of bone. Adrenal corticosteroids stimulate resorption of bone and inhibit the formation of new bone. In cattle, castration delays the completion of growth in epiphyseal plates (Brannang, 1971a). This is most noticeable in the distal bones of the limbs and enables the continued longitudinal growth of the legs. In the vertebral column, however, castration reduces bone growth. The reduction is centered on the first thoracic vertebra. Removal of the ovaries from heifers also causes an increase in the longitudinal growth of distal bones (Brannang, 1971b). Growth factors may mediate or augment the activity of the hormones controlling bone growth. Both rapidly growing and adult bones may contain transforming growth factor beta (TGF-beta), beta-2 microglobulin (beta-2 m), and insulinlike growth factor I (IGF I). In adults, these factors may be involved in skeletal remodeling (Canalis et al., 1988).

At localized sites, intrinsic activity in bone (Behari and Andrabi, 1978) probably plays an important role in regulating bone growth, but its current status is uncertain (Moss, 1972). One hypothesis is that loads that are frequently placed on a region of bone cause the transduction of mechanical energy to electrical energy by a piezoelectric effect. In a frequently loaded and negatively charged region, growth is stimulated. The growth of both bone and cartilage may be modified by the application of pulsed magnetic fields (Archer and Ratcliffe, 1983). In electrical fields, osteoclasts migrate towards the positive electrode while osteoblast-like cells migrate towards the negative electrode (Ferrier et al., 1986).

In an unloaded and positively charged region, resorption is stimulated. In bovine bone, it has been suggested that adaptation to changing load patterns occurs by a viscoelastic "creep" at the cement lines that surround individual Haversian systems (Lakes and Saha, 1979). Differences in the arrangement of hydroxyapatite crystallites, in lacunar structure, and in the transition from spongy to compact bone have been observed between the bones of wild and domesticated sheep. These differences may have accompanied a reduction in exercise (Drew et al., 1971). Severe malnutrition of pigs reduces the formation of Haversian bone (Luke et al., 1980).

The gross anatomy of muscles and skeletal units are closely matched, and mutual or interacting control systems probably exist. Because most farm animals are slaughtered in a fairly immature condition, the relationship between muscles and the bony processes that they pull on may not be immediately obvious. But knobs and wrinkles on bone surfaces become more conspicuous with age, and they are readily seen in the carcasses of old bulls. One possible relationship between muscle activity and bone growth may be that isometric contraction, by stopping or slowing the venous blood flow, may stimulate bone growth. Alternatively, by pulling on the periosteum, the effect of muscle activity may be mediated by connective tissue. The importance of local factors is seen in bone transplants, where growth of the transplant almost immediately becomes regulated by the new local conditions.

Joints

When the movement of two adjacent bones is normally restricted or impossible, the joint between them is called a synarthrosis and different types of connective tissue act as a cement between the two adjacent bones. Skull suture joints are cemented by collagen, sternebrae are separated by cartilage, and fibrocartilage occurs in the symphysis pubis and in intervertebral disks. Completely movable joints are called diarthroses. Their articular surfaces are covered by smooth hyaline cartilage lubricated by synovial fluid. Under suitable conditions, diarthroses can be made to develop in organ cultures (Fell, 1956), and development may proceed in the absence of muscular and neural connections, provided that elements of the two diaphyses are present and are separated by undifferentiated tissue. The joint capsule, however, does not develop properly under these conditions, and the proper shaping of articular surfaces only develops with normal use. Lack of normal joint use resulting from lack of exercise might cause leg weakness or stiffness in meat animals. Progressive thinning of articular cartilage with age may be widespread in poultry (Duff, 1987).

Neonatal farm animals are sometimes afflicted by genetic or environmentally-induced joint malformations. Rigid joints are most frequently observed in the distal parts of a limb so that locomotion is difficult or impossible. The name arthrogryposis is frequently used to describe this condition, but is a rather misleading term since arthrogryposis means "crooked-joints" whereas, in reality, joints are often fixed in an extended or rigidly straight position. James (1951) preferred the term "multiple congenital articular rigidity" in human medicine. Since farm animals often have only a single affected joint, the term "multiple" is redundant. Four degrees of immobilization may occur; (1) when the joint can be freed by an externally applied non-destructive force, (2) when muscles and tendons acting around the joint must be severed, (3) when capsular ligaments as well as muscles and tendons must be severed, and (4) when mobilization is impossible due to deformation of the articular surface (Swatland, 1974). The term ankylosis is inappropriate for congenital articular rigidity since it implies that bones are fused across the joint; with one exception (Schmalstieg and Meyer, 1960) this is not a general feature of the condition.

Hereditary congenital articular rigidity in cattle is often associated with a CLEFT PALATE in the skull (Leipold et al., 1969; Greeley et al., 1968; Greeley and Jolly, 1968). However, there are a number of reports of hereditary congenital articular rigidity in cattle without cleft palate but with defects such as abnormal limb shape (Hutt, 1934) or hair defects (Tuff, 1948; Nes, 1953). Hereditary bovine congenital articular rigidity is usually caused by a recessive gene without sex linkage. In Australia, congenital articular rigidity sometimes occurs in conjunction with hydranencephaly (absence or gross reduction of cerebral hemispheres) in cattle (Blood, 1956; Whittem, 1957). The condition is not heritable and may be induced by ingestion of a plant teratogen (Hindmarsh, 1937). In the western states of US, a number of species of the plant Lupinus may cause congenital articular rigidity in cattle (Wagnon, 1960; Shupe et al., 1967; Keeler et al., 1969). For access to the extensive literature on plant teratogens consult Keeler (1984).

In pigs, congenital articular rigidity may be caused by either genetic or environmental factors. Hereditary congenital articular rigidity in pigs probably is caused by an autosomal recessive gene (Hallqvist, 1933) and may be associated with an unusual condition known as the thick-leg syndrome (Morrill, 1947; Koch et al., 1957; Kaye 1962) in which certain bones are thickened by the activity of extra osteoblasts, and fibrous connective tissue may invade muscles and joint capsules. Environmentally induced congenital articular rigidity in pigs may be caused by nutritional deficiencies of vitamin A (Palludan, 1961) or manganese (Miller et al., 1940), or from ingestion of teratogens from plants such as tobacco (Crowe, 1969; Crowe and Pike, 1973), Conium maculatum (Edmonds et al., 1972), Prunus serotina (Selby et al., 1971) or Datura stramonium (Leipold et al., 1973b).

Both genetic (Roberts, 1929; Middleton, 1932, 1934; Zophoniasson, 1929; Morley, 1954) and environmental (Keeler et al., 1967; James et al., 1967; Stamp, 1960; Nisbet and Renwick, 1961) causes of congenital articular rigidity are known in sheep. Judging from the variety of different factors giving rise to the same end result, congenital articular rigidity, the different factors may be acting on the same developmental mechanism - a mechanism that may involves the interaction of nerves, muscles and joints (Chapter 76.

Animal size and bone development

Breeds of cattle with a large mature size usually produce lean meat at a faster rate than early maturing traditional beef breeds with a relatively small adult size. Differences in adult size are produced by differences in skeletal growth, and relationships between the quantitative anatomy of individual bones and meat production traits in beef cattle have been identified (Wilson et al., 1977). Relationships between skeletal and muscular development may involve meat quality because large-framed animals produce leaner meat during their production life span. Large-framed breeds mature late and have a later cessation of linear skeletal growth at their epiphyseal plates. The time of maturation is related to the distribution and amount of adipose tissue in the carcass, particularly marbling fat. Differential bone growth between large and small breeds of cattle is usually established prior to a slaughter weight of 500 kg in males (Jones et al., 1978). But the emphasis that is placed on animal height by many beef breeders may be misplaced. Gilbert et al. (1993) compared present day cattle with those born 20 years ago and found that the faster growing modern animals were longer in body, but not necessarily taller than their predecessors.

Pelvic dimensions in cows of different breeds are related to the incidence of difficult calving or dystocia (Laster, 1974; Neville et al., 1978). Dystocia may be particularly serious when a homozygous double-muscled calf is born. Double-muscling in the calf is caused by increase in the number of muscle fibers so that the shape of the calf is very bulky. The dam, which may be either a heterozygous carrier or completely double muscled, may also exhibit some reduced bone development (Hendricks et al., 1973).

Although one might expect the proportion of bone in a carcass to affect specific gravity measurements, this may not be evident in practice (Preston et al., 1974). Growth promotants may have little or no effect on skeletal development (Ralston et al., 1975), but environmental factors do affect bone development, since certain confinement conditions cause lameness involving skeletal joints (Murphy et al., 1975).

The early research on bone development in beef carcasses is reviewed by Preston and Willis (1974). In the early 1950s, attempts were made to use measurements of isolated carcass bones such as the cannon bone to predict the muscle to bone ratios of carcasses. Although the method worked satisfactorily when applied to a wide range of dissimilar carcasses, it was of little practical value when applied to more uniform commercial carcasses. Muscle to bone ratios improve as animals grow older or fatter, since longitudinal bone growth slows down in older animals and muscles start to accumulate appreciable amounts of intramuscular fat. Animal age is the dominant factor that determines muscle to bone ratios (Preston and Willis, 1974). However, when adjustments are made for animal age and carcass weight, considerable unexplained variation still is found in muscle to bone ratios (Dolezal et al., 1982).

An emphasis on larger, leaner breeds of cattle has obscured the fact that, some years ago, the emphasis was in the opposite direction. The desire to produce small compact animals with bulging muscles favored the survival of dwarf animals with impaired longitudinal bone growth. Although mildly affected animals looked very muscular (Marlowe, 1964), severely affected animals became increasingly common and were poorly suited for beef production. Dwarfism from impaired longitudinal growth of bones is a recessive trait that affects males more strongly than females (Bovard and Hazel, 1963).

From research on allometric skeletal growth in pigs (Doornenbal, 1975; Davies, 1975), it appears that a relationship between mature body size and the potential for lean meat production also may exist in pigs. In other words, selection for leaner pork may favor greater skeletal dimensions in the mature animal, due to later maturation.

Lameness and locomotor disorders are a particularly serious problem in pigs. Optimum dietary levels of calcium and phosphorus, and the required levels of vitamin D, vitamin A, copper and manganese have been established (Pond et al., 1975). Connective tissue metabolism is affected by levels of iron (Prockop, 1971), manganese (Leach, 1971), copper (Carnes, 1971) and zinc (Westmorland, 1971). Since bones contain collagen fibers, factors that affect collagen synthesis also act on bone growth (Bengtsson and Hakkarainen, 1975). Although exercise levels do not produce much noticeable effect on bone growth (Murray et al., 1974), there is evidence that certain confinement conditions do affect the breaking strength of bone (Elliot and Doige, 1973). The pathogenesis of malfunctioning joints in the absence of any readily apparent infection remains a difficult problem (Hogg et al., 1975). A predisposition to such conditions as a result of selective breeding for meat production has been proposed, and pathological changes have been found in the skeletal cartilage of pigs prior to, or without evidence of lameness (Thurley, 1969). However, the incidence of pathological conditions affecting the joints is about the same in slow-growing pigs as in fast-growing pigs so that factors other than simply animal growth rate must be involved (Nakano et al., 1984).

The avian epiphyseal plate has blood vessels that penetrate deeply into the zones of cellular proliferation whereas, in mammals, the epiphyseal plate is almost devoid of such vessels. There is considerable sexual dimorphism in the longitudinal growth of bones in chickens and turkeys, and males have longer bones with a later completion of growth (Latimer, 1927; Sullivan and Al-Ubaidi, 1963). Wise (1977) found that the bones of broiler-type chickens tend to be shorter and thicker than those of layer-type birds. A relationship between mature body size and the potential for lean meat production also has been observed in poultry. Thus, at equal body weights, the skeletal system may be less mature in broilers selected on the basis of meat yield, than in layers. As well as skeletal defects caused by nutritional deficiencies (vitamin D, vitamin A, phosphorus, manganese, choline, biotin, nicotinic acid, folic acid, zinc and pyridoxine), poultry are afflicted by a number of genetic skeletal defects. Asymmetrical or abnormal development of the sternum may occur in birds at market weight and is a cause for downgrading.

THE MICROSTRUCTURE AND DEVELOPMENT OF ADIPOSE TISSUE

Introduction

In the overall balance sheet for energy in agriculture, relatively large amounts of feed energy are used when animals deposit fat in their bodies, but much of this fat is removed and wasted after the animal is slaughtered. About 6% of the live weight of a steer may be removed as fat in the abattoir, and an equal amount may be trimmed from the dressed carcass by the butcher. Adipose tissue only serves its proper function when an animal uses the energy and insulation provided by its adipose tissue to survive a period of inadequate feed intake or cold weather. Adipose tissue in meat, however, is not altogether undesirable and wasteful. It is desirable in moderation to give a "finished" appearance to a carcass and, without at least some subcutaneous fat, a carcass is judged to be unattractive by traditional standards. When nutrition has an effect on meat quality, the effect may be mediated via changes in fat, leading to changes in tenderness and juiciness (Berge et al., 1993).

Fat that is deposited within muscles (intramuscular adipose tissue) appears as a delicate pattern of wavy lines in the meat - hence its common name, marbling fat. It is traditionally maintained that marbling fat contributes to the juiciness of cooked meat because it melts away from between bundles of muscle fibers to make the meat more tender and more succulent. However, it is difficult to find much scientific evidence in support of this traditional view (Breidenstein et al., 1968), except in poultry where subcutaneous and intermuscular fat baste the meat as it is being roasted. Berry and Leddy (1990) found that steaks with little marbling had desirable palatability after rapid, high temperature cooking. However, the steaks had been properly conditioned or aged (21 days). Unscientific as it may be, there is considerable reluctance to allow completely lean beef onto the market as top quality meat, unless there is some redeeming factor such as extreme youth (heavy baby beef) or excellent conditioning (21 days or more).

Much of the characteristic flavor associated with different types of meat originates from carbonyl compounds concentrated in the adipose tissue (Sink, 1979; Wasserman, 1979). The flavor of meat is sometimes modified by the animals' diet (Park et al., 1972). Not all the fat in the carcass is macroscopically visible. Many muscle fibers, particularly those in postural muscles, contain large numbers of microscopic fat droplets (Bullard, 1912). The storage of fat droplets within muscle fibers is related to the overall metabolism of the animal. For example, the deposition of lipid in the postural muscles of cows reaches a peak at about one week after calving when lipid stores in the liver also are at their highest (Roberts et al., 1983).

The ancestors of present-day farm animals once were used to feed populations of people, many of whom were manual workers who expended large amounts of energy in tasks that we now perform by machine. Fat contains more stored energy than lean, and the high fat content of meat once supplied much of the energy that people expended in their daily work. Now this extra energy is undesireable, and a reduction of the fat content of meat is a major goal in the continued improvement of meat animals. Most of the meat eaten in the 19th century was derived from older animals than are marketed now, and most of these animals had already given long service in the production of milk or wool, or had been used to pull plows or wagons. The presence of marbling fat would have greatly improved the palatability of the tough and strong tasting meat derived from these mature animals. Also, large families were more common a hundred years ago, and large joints of meat cut from large carcasses were well suited to domestic requirements.

To understand how meat may be made leaner, we need to look at the origin, metabolism and proliferation of adipose cells (Allen et al., 1976; Leat, 1976; Garton, 1976; Evans, 1977).

White adipose cells

A mature adipose cell or adipocyte may have a diameter of about 100 microns and is filled with triglyceride. Thus, its nucleus and cytoplasm are restricted to a thin layer under the cell membrane. The adipose cells of species of large mammals are larger than those of small mammals. Relative to other mammals, carnivores and ruminants tend to have adipose tissue that is characterized by large numbers of small cells (Pond and Mattacks, 1985a). When isolated from their surroundings, adipose cells are rounded in shape but, when packed together in adipose depots, they are compressed with flattened sides. Pockets of very small adipose cells sometimes appear between normal-sized cells and this may bias measurements of mean cell size, depending on how well the small cells are detected (Kirtland et al., 1975; Ashwell et al., 1975). When making histological measurements on frozen sections of adipose tissue, it is important to ensure that each cell has a nucleus. Large adipose cells are easily fragmented and, when a section is thawed, the cell fragments may become rounded like oil droplets. Some of these fragments may appear to be bounded by part of the original cell membrane, and they can only be distinguished from real, small adipose cells by their lack of a nucleus.

Mature adipose cells have very little cytoplasm and contain few organelles. The golgi complex is small, there are only a few ribosomes and mitochondria, and the endoplasmic reticulum is sparse. The large triglyceride droplet that fills the bulk of each cell is not directly bounded by a membrane, although it may be restrained in position by a delicate meshwork of very thin filaments approximately 10 nm in diameter. These filaments are most conspicuous in the adipose cells of poultry (Wood, 1967).

In many locations in the body, large numbers of adipose cells are grouped together to form adipose depots. Adipose cells are kept in place by a meshwork of fine reticular fibers (Motta, 1975; Swatland, 1975) responsible for the weak fluorescence of adipose tissue. Large adipose depots usually are subdivided into layers or lobules by partitions or septa of fibrous connective tissue. In the layered subcutaneous fat of a pork carcass, the septa may follow the body contours and may create a weak boundary layer echo in the ultrasonic estimation of fat depth. Adipose depots are well supplied by blood capillaries (Gersh and Still, 1945).

Brown adipose cells

Brown adipose cells may occur in cold-adapted, hibernating or newborn mammals. Apart from their brown color, caused by greater vascularity and a high cytochrome concentration, brown adipose cells have a different appearance microscopically. They are are small, they have abundant cytoplasm and mitochondria, and their triglyceride is subdivided into a number of small droplets (Figure 2-34). At the ultrastructural level, the mitochondria of brown adipose cells may have a distinct appearance caused by parallel arrays of long cristae (internal partitions), most of which extend across the full width of the mitochondrion. Brown adipose cells generate heat by a process called non-shivering thermogenesis. Instead of releasing fatty acids into the blood stream, like white adipose cells, brown adipose cells oxidize their own fatty acids to release heat instead of synthesizing ATP. Cold animals also may generate heat by shivering, using their muscles to convert chemical energy to heat. When animals with brown fat are placed in a thermoneutral environment, non-shivering thermogenesis is increased by the intravenous infusion of norepinephrine (noradrenaline). When stimulated by acute cold, sympathetic axons release norepinephrine that reaches the beta-adrenergic receptors on the surface of brown adipose cells (Nicholls, 1983).

Unfortunately, the study of brown adipose tissue in farm mammals is complicated by the fact that, relative to some other mammalian species, the distinctions between brown adipose cells and immature cells of ordinary white adipose tissue are seldom distinct. During their development, white adipose cells pass through a multilocular stage with many small triglyceride droplets before they reach their final unilocular stage of development with one large triglyceride droplet. Thus, it is often difficult to decide on morphological grounds alone whether a multilocular adipose cell in a neonatal animal is a brown adipose cell or an immature white adipose cell. In newborn farm animals, it is possible that immature white adipose cells exhibit some of the heat-generating properties of brown adipose cells. In hamsters, multilocular brown adipose cells may develop either from unilocular adipose cells or by proliferation from endothelial cells (Nechad and Barnard, 1979).

In lambs (Thompson and Jenkinson, 1969; Gemmel et al., 1972), calves (Alexander et al., 1975) and kids (Thompson and Jenkinson, 1970), brown adipose cells have been identified on the basis of their mitochondrial morphology and norepinephrine response, although other expected features such as brown color and a multilocular condition were indistinct. In calves, Alexander et al. (1975) found that brown fat formed approximately 2% of the body weight, but none of the adipose tissue located subcutaneously exhibited the properties of brown adipose tissue. In farm mammals, any brown adipose tissue present at birth to warm the newborn animal is seldom maintained for very long. Dauncey et al. (1981) identified brown adipose tissue in neonatal pigs, and found that small amounts persisted until at least three weeks after birth. Where brown fat does occur in newborn farm animals, it is probably converted to ordinary white adipose tissue later in development. The transition from brown to white adipose cells has been observed among cells in culture (Dyer and Pirie, 1978). However, when white adipose cells lose their triglyceride, they do not revert back to being brown adipose cells (Cox et al., 1978).

Origin of adipose cells

Speculation concerning the origin of adipose cells has continued since the end of the 19th century, by which time most of the feasible possibilities had already been proposed (Clark and Clark, 1940). In certain locations in the body of a fetus, where adipose tissue will develop later in life, mesenchyme cells congregate in lobules resembling glandular tissue. In an embryo, the mesenchyme is a diffuse tissue composed of cells that may differentiate to form the connective tissues of the body, as well as blood and lymphatic vessels. The mesenchyme cells in gland-like lobules begin to accumulate small droplets of triglyceride. The droplets coalesce as they are crowded together and, finally, they form a single large mass in the center of each cell. In other locations where adipose tissue is about to develop, the adipose cell precursors (sometimes called preadipocytes) may resemble fibroblasts. In this case, cells are spindle shaped with an oval nucleus, and the cytoplasm contains rough endoplasmic reticulum, microtubules, microfilaments and spherical mitochondria (Slavin, 1979). In pigs, presumptive adipose cells contain one to three prominent nucleoli in their otherwise pale nuclei (Hausman and Martin, 1981).

It is difficult to prove that the gland-like cells and the fibroblast-like cells that give rise to adipose cells are distinct types of cells with a rigid pattern of development. However, the possibility that cells are rigidly programmed to develop exclusively into adipose cells is supported by the behavior of transplanted cells. Cells that have been transplanted from precursor adipose lobules to future low-fat regions will store triglyceride and become adipose cells, even in an inappropriate location (Le Gros Clark, 1945). Similarly, precursor adipose cells from certain sites will continue their programmed development into adipose cells, even if they are removed from the body and cultured in vitro (Van and Roncari, 1977, 1978; Poznanski et al., 1973; Van et al., 1976; Dardick et al., 1976). However, the trigger to differentiation appears to be hormonal because it is influenced by insulin, triiodothyronine and insulin-like growth factor (IGF-1; Hausman, 1989). The trigger or inducer acts upon a commiting gene for the differentiation pathway (Chen et al., 1989).

In neonatal pigs, there are four types of cells, any or all of which could be adipose cell precursors (Mersmann et al., 1975). One source of adipose cells may be the recruitment of fibroblasts. Fibroblasts can increase in number by mitosis. When recruited to become adipose cells, they give up the shape and activities of a fibroblast, and pass through a multilocular stage of triglyceride accumulation, finally to become adipose cells resembling those derived from precursor cells (Clark and Clark, 1940). Adipose cells sometimes originate from the endothelial cells of the vascular system (Tavassoli, 1976) and there is evidence of this in pigs (Wright and Hausman, 1990). Although extra adipose cells may be recruited when animals become obese, the extra cells are retained when an animal returns to its normal level of fatness (Faust et al., 1978). Macrophages also have been suggested as a source of new adipose cells (McCullough, 1944; Smith et al., 1952), and pluripotent mesenchymal stem cells capable of forming a variety of cell types may exist within connective tissues (Young et al., 1993).

Adipose tissue distribution

Adipose depots range in size from small groups of adipose cells located between muscle fiber bundles, to the vast numbers of adipose cells that are located subcutaneously and viscerally. It is important to distinguish between anatomical sites and systemic locations. Specific muscles or regions of the carcass are anatomical sites. Intermuscular, intramuscular, visceral and subcutaneous are systemic locations. For example, fat from a specified anatomical site such as the shoulder may be separated into different systemic depots (subcutaneous, intermuscular and intramuscular). The distinction between anatomical sites and systemic locations is important commercially. For example, bovine intramuscular marbling fat sometimes first becomes noticable in rump and loin muscles, where it adds to their value. In the same systemic location, but at a different anatomical site such as the brisket, marbling fat confers no economic advantage and is wasteful. In cattle, the relative growth of subcutaneous fat is similar in both the forequarter and the hindquarter. However, the relative growth of intermuscular fat is higher in the forequarter than in the hindquarter (Berg et al., 1979). There is no guarantee that systemic fat depots are homogeneous, even at a single anatomical site. For example, the backfat seen on pork carcasses is subdivided into three layers that differ in their composition and pattern of growth (Moody and Zobrisky, 1966). Similarly, there may be considerable differences between the adipose cells from different anatomical locations. Ramsay et al. (1989) found that preadipocytes from the shoulder and ham responded to hydrocortisone whereas preadipocytes from the perirenal region did not.

The systemic deposition of fat in a carcass influences commercial indices of carcass composition such as the dressing percentage. Intramuscular fat present in meat at the time of cooking is mostly retained within the meat (Renk et al., 1985). The total of omental fat, mesenteric fat and kidney fat may constitute about 30% of the total fat in a beef steer (Cianzo et al., 1982).

CARCASS WEIGHT

DRESSING % = ---------------- x 100

LIVE WEIGHT

Thus, most of the fat deposited around the viscera is removed with the viscera at slaughter, and this reduces the dressing percentage. Fat that is deposited between or within carcass muscles increases the dressing percentage.

In cattle, it was traditionally maintained that fat deposition followed three systemic phases (Hammond et al., 1971). In the first phase, fat was thought to be deposited around the viscera and kidneys, and within the caul and mesenteries. Caul fat is a thin sheet of adipose tissue contained in a large fold of connective tissue over the stomach and adjacent organs. In the second phase, fat was thought to be deposited subcutaneously and intermuscularly. The third phase was thought to be the deposition of marbling fat within muscles. Sometimes, however, no simple chronological separation of the three phases of fat deposition is detectable, and the relative amounts of fat in the main systemic locations may remain constant (Berg and Butterfield, 1976). On a high energy ration, cattle may deposit subcutaneous fat at a greater rate than they deposit intermuscular fat. However, this difference does not appear when animals are on a low energy ration (Fortin et al., 1981). Breeds of cattle differ in the way that they develop their systemic fat depots. For example, Herefords produce more subcutaneous fat and less perirenal and pelvic fat than Angus, Friesian and Charolais crossbred cattle (Charles and Johnson, 1976). However, when adipose growth at different anatomical sites is examined, relative to total fat, only minor differences may be found between breeds (Berg et al., 1978).

In pigs, subcutaneous fat grows at the same rate as total body fat, intermuscular fat grows more slowly, and visceral fat grows faster (Kempster and Evans, 1979). The separation of phases in adipose deposition is complicated by the fact that, while the experimenter usually works in terms of calendar days and weeks, the experimental animal is following its own physiological calendar. The animal's physiological calendar is based on events that mark the progress through its life cycle (Chapter 8). For example, skeletal and reproductive development follow an orderly sequence of events, and different breeds may progress through this sequence at different rates. Measured in calendar weeks and months, early-maturing breeds deposit noticeable amounts of marbling fat before late-maturing breeds. Thus, the introduction of late-maturing breeds with a large adult size may be used to delay fat deposition and to enhance lean growth in cattle populations.

Sex hormones may produce large and economically important differences in the overall fatness of beef, mutton and pork carcasses. Provided that comparisons are made at equal fatness, however, the distribution of fat within beef carcasses may be similar for both sexes (Berg et al., 1979). Bulls, rams and boars generally produce leaner carcasses than steers, wethers and barrows, respectively. Steers and wethers generally produce leaner carcasses than cows and ewes, respectively. In pigs the situation is reversed, and gilts generally produce leaner carcasses than barrows. The deposition of fat tends to occur at a lighter weight in heifers than in steers, and at a lighter weight in steers than in bulls.

Adipose tissue metabolism

Although adipose tissue depots are primarily for energy storage or for mechanical and thermal insulation, they are far from passive. Anabolic processes are very active when an animal is depositing fat, while catabolic processes are very active when an animal is living off its energy reserves, either during prolonged starvation or between periods of feeding. The metabolism of adipose cells is altered in cows after calving, due to the mobilization of adipose reserves for milk production: lipogenesis is reduced and lipolysis is increased (Pike and Roberts, 1980). Metabolic water may be produced by oxidation of fat when animals are deprived of water. Thus, the depth of backfat and the total weight of subcutaneous fat may be reduced in pigs on a restricted water intake (Skipitaris, 1981).

Energy is stored in molecules of triglyceride (Tables 2-4 and 2-5). The long-chain fatty acids found in animal triglycerides (R1, R2 and R3 in Table 2-4) vary in length, as shown by the values of "n" in the general formula for fatty acids. The value for "n" in animal fat usually ranges from 5 to 20. These fatty acids are insoluble in water, like the triglycerides that are formed from them. Adjacent carbon atoms along a chain may be linked together with an unsaturated double bond. Unsaturated bonds produce a bend in the chain and this lowers the melting point. Unless complicated by the presence of unsaturated bonds, melting points are proportional to chain length.

Table 2-4. The Structure of Triglyceride

G - FATTY ACID

L

Y

C - FATTY ACID

E

R

O

L - FATTY ACID

H O

¦ ¦

H - C - O - C - R1

¦

¦ O

¦ ¦

H - C - 0 - C - R2

¦

¦ O

¦ ¦

H - C - O - C - R3

¦

H

Table 2-5. The Structure of Fatty Acids

General structure CH3-(CH2)n-COOH

Names for values of "n"

2 - butyric

4 - caproic

6 - caprylic

8 - capric

10 - lauric

12 - myristic

14 - palmitic

16 - stearic

18 - arachidic

20 - behenic

The degree of saturation of carcass fat is affected by a number of factors. Within a breed, steers may have more saturated fat than heifers (Terrell et al., 1969) but differences may also exist between breeds, as in comparisons of Japanese Wagyu cattle with Angus (May et al., 1993). About 12 months after birth, cattle normally start to fatten rapidly, and progressively more unsaturated fatty acids are deposited (Leat, 1975, 1977). Genetically obese pigs have a greater degree of saturation of their fatty acids than normal pigs (Scott et al., 1981).

Most plant oils contain a high proportion of unsaturated fatty acids with low melting points, but animals have a high proportion of saturated fatty acids and their fat is solid or semi-solid at room temperature. Thus, when a beef carcass is cooled from body temperature down to nearly 0oC after slaughter, subcutaneous fat passes from a liquid to a solid state. In beef carcasses, a greater proportion of fatty acids may be saturated in the summer than in the winter (Link et al., 1970). Seasonal changes of this type are quite marked in wild sheep, where they may help the animal to adapt to climatic changes (Turner, 1979). Fat tends to be soft and yellow in ram carcasses relative to wethers, and fat also tends to be soft and yellow in lambs fed a high-energy diet relative to those on a low-energy diet (Busboom et al., 1981).

In general, triglycerides located subcutaneously where they may be relatively cool in the live animal have a low melting point. Conversely, perirenal or suet fat in the beef carcass is brittle at room temperature since it comes from a warm place in the body and has a high melting point. In the abattoir, shrouds are often pinned over beef carcasses so that the molten subcutaneous fat solidifies with a smooth surface.

In animal fat, most fatty acids contain an even number of carbon atoms, and unsaturated bonds are either absent or few in number. In meat, the most common unsaturated fatty acids are oleic, linoleic and linolenic acids with 18 carbons and with 1, 2, and 3 double bonds, respectively.

Systems involved in triglyceride deposition

The integration of adipose metabolism with that of the rest of the body is rather complicated. At a greatly simplified level, the major systems involved are shown in Figure 2-35. At the top of the diagram is an adipose cell, complete with a diagramatic trapdoor to represent the countless membrane sites by which glucose gains access to the cell. In the diagram, two short lengths of tube represent the blood vascular system and the small intestine. The liver is also shown diagramatically. Figure 2-35 also contains a diagramatic representation of a reticuloendothelial cell. The reticuloendothelial system is formed from a heterogeneous collection of individual cells that are widely dispersed through the body. Although the cells of the reticuloendothelial system may differ in appearance and location, they all share certain common properties, many of which are related to defence against infectious microorganisms. Reticuloendothelial cells are involved in adipose tissue metabolism. Figure 2-35 contains some letter "E" symbols that represent glycerol, fatty acids and triglycerides.

Dietary intake

In the case of a pig that has some triglyceride in its diet, the triglyceride reaches the small intestine and is hydrolyzed in the presence of pancreatic lipase (Figure 2-36A). Colipase, a protein secreted by the pancreas, binds to the surface of fat droplets in the presence of bile salts from the liver and facilitates the attachment of lipase (Patton and Carey, 1979). The partially or completely separated fatty acids may pursue a number of different pathways, one of which involves absorption by an intestinal cell and reassembly into a new triglyceride inside the cell. Large numbers of reassembled triglyceride molecules may be wrapped in protein and phospholipid to form a small globule about one micron in diameter called a chylomicron. The triglycerides contained in the chylomicra then pass to the lacteals (Figure 2-36B). Lacteals are blind-ending lymphatic capillaries in the axes of the intestinal villi. The lymphatic system conducts chylomicra to the venous system where, after passing into the general circulation, they are made available to adipose cells (Figure 2-36C). However, chylomicra do not enter directly into adipose cells. Their triglyceride is first hydrolyzed to fatty acids and glycerol by lipoprotein lipase (clearing factor lipase) located on the surfaces of adipose cells or capillary cells (Figure 2-36D). The transfer of the fatty acids to the adipose cell is facilitated by the continuity of intercellular connections with intracellular channels (Blanchette-Mackie and Scow, 1981). Yet again, triglycerides now are reassembled, this time within the adipose cell (Figure 2-36E). Lipoprotein lipase activity in adipose tissue is linearly related to cell size. In young obese rats, lipoprotein lipase activity is deficient in red muscle fibers (Hartman, 1980). There is also a hormone-sensitive lipase in skeletal muscle that may be involved in intramuscular lipolysis (Oscai et al., 1990).

Most of the glycerol released by lipoprotein lipase outside the adipose cell returns to the blood stream (Figure 2-36F). Inside the adipose cell, a new carbon backbone for the triglyceride is formed from glycerophosphate that has been derived from glucose (Figure 2-36G). The entry of glucose into a cell is facilitated by insulin. In porcine adipose tissue the enhancement of carbohydrate metabolism by insulin is antagonized by porcine growth hormone (Walton and Etherton, 1986). This may explain the leaner carcasses and greater lean growth of pigs treated with growth hormone.

By a simple route, such as that outlined above, ingested fatty acids may be deposited in an unaltered form in their new "host". Thus, in pigs, the degree of saturation of carcass fat is influenced by the nature of the fatty acids that are ingested. This may affect meat quality because fat with a low melting point appears greasy. Fat with a relatively large number of unsaturated bonds also is more likely to undergo auto-oxidation with the formation of unpleasant odors (Lawrie, 1974). Commercial problems often arise when highly unsaturated fats such as linseed oil or fish oil are fed to monogastric animals, since these oils may cause unpleasant odors in the meat. Unpleasant odors may not become apparent until the meat has been cooked and its unsaturated fatty acids have been oxidized (Reineccius, 1979). Such problems are rare in ruminants since unsaturated fats are reduced in the rumen. However, prior treatment of the feed with formaldehyde inhibits this reduction and enables unsaturated fatty acids to pass through the digestive system relatively unchanged (Cook et al., 1970). In cattle and sheep, dietary lipids are extensively modified in the rumen. Esterified fatty acids are release by hydrolysis, the residual glycerol and galactose are fermented, and unsaturated fatty acids are hydrogenated (Garton, 1967). Large amounts of stearic acid are formed from oleic, linoleic and linolenic acids. These, together with other fatty acids derived from the microflora, are absorbed in the intestine. In pigs, a high linoleic intake in the diet is notorious for softening the subcutaneous fat of the carcass and causing problems for meat cutters. Soft pork fat may be detected texturally (Irie and Ohmoto, 1982; Dransfield and Jones, 1984), ultrasonically (Miles et al., 1985), and by fiber optic spectrophotometyry (Irie and Swatland, 1992).

There is also the possibility of using the direct dietary incorporation of fatty acids to some advantage, as in the case of 20-carbon omega-3 fatty acids from marine sources. These have acquired a reputation for being a desireable component of the human diet. The basic problem is that the inclusion of marine oils into animal feeds causes off-flavors in the product. A search is currently underway for terrestrial sources of omega-3 fatty acids, coupled with more favorable refining methods (Hargis and Van Elswyk, 1993).

Alternative inputs and lipogenesis

An alternative carbon input is available to adipose cells from very-low-density lipoproteins (VLDL) from the liver (Figure 2-37 and Table 2-6). In VLDL, a central core of triglyceride is packaged by a layer of protein, phospholipid and cholesterol. The leaching of triglyceride by lipoprotein lipase near to adipose cells (Figure 2-37A) leads to a reorganization of VLDL components, and a low-density lipoprotein (LDL) is produced. The cholesterol that remains in LDL is released into the blood stream by cells of the reticuloendothelial system (Figure 2-37B). In the liver, cholesterol, triglyceride and circulating high-density lipoproteins (HDL) are assembled into new VLDL (Figure 2-37C).

Table 2-6. Percent composition of components used for triglyceride transport.

---------------------------------------------------------------

Composition Chylomicron VLDL LDL HDL

---------------------------------------------------------------

Triglyceride 85 50 10 4 Cholesteryl ester 3 12 37 15 Unesterified cholesterol 1 7 8 2

Protein 2 10 23 55

Phospholipid 9 18 20 24

Overall density <0.95 <1.006 1.019 1.063

to 1.063 to 1.21

---------------------------------------------------------------

Meat animals may form new fat by lipogenesis, as shown in the classical experiments of Lawes and Gilbert (Hall, 1905). In pigs, most of the new triglyceride is synthesized from glucose, and the fatty acids that are produced are dominated by stearic acid and its desaturated form, oleic acid. In ruminants, acetate is the main substrate, and stearic acid is the dominant fatty acid. In poultry, lipogenesis occurs in the liver rather than in adipose cells, and triglyceride reaches the adipose cells as VLDL in the blood. Oleic and palmitic acids are dominant when the dietary intake of triglyceride is low (Evans, 1977).

LOW-FAT MEAT

The genetic and nutritional factors that regulate the amount and distribution of adipose tissue can be described in terms of cellular hyperplasia (an increase in cells numbers) and cellular hypertrophy (an increase in cell size). An early stimulus to research on these topics was provided by the hopeful hypothesis that adipose cell numbers might be genetically regulated, while adipose cell size might be nutritionally regulated. Had this been the case in meat animals, selection for animals with low numbers of adipose cells might have provided a method for enhancing the production of lean meat. It was also suggested that a low plane of nutrition early in life might reduce the numbers of fat cells, so that animals would be less likely to deposit fat in the final stages of growth to market weight.

Once adipose cells start to store triglyceride, they are no longer capable of mitosis (Desnoyers et al., 1980), and insulin causes increased adiposity by increasing the size rather than the number of adipose cells (Salans et al., 1972). It has also been proposed that adipose cell size is involved in the regulation of feed intake and triglyceride deposition (Faust et al., 1977).

The basic cellular problem that has daunted hopes of reducing the number of adipose cells by a low level of nutrition early in life is that populations of morphologically identical adipose cells in adult animals are formed from an initial population of specific precursor cells which is variably supplemented by the recruitment of fibroblast-like cells. Thus, the apparently limitless possibilities for recruitment of extra adipose cells may allow compensatory growth to offset any reduction in the initial population of specific precursor cells.

However, some possibilities do remain. In double-muscled cattle, adipose cells are greatly reduced in number (cellular hypoplasia). Since double-muscling is a genetic condition, there could be a simple genetic mechanism that regulates adipose cell numbers in these animals. Adipose cell number and size can be modified by the selective breeding of mice for either high or low postweaning growth rate (Martin et al., 1979). Regulation of adipose cell numbers is affected by thyroid hormones but not by growth hormone (Ramsay et al., 1987).

Passive immunization of meat animals with antibodies developed against adipose cell membranes offers a novel approach to the age-old problem of fat reduction in meat animals. Nassar and Hu (1991) showed that perirenal fat and subcutaneous fat may be reduced in lambs by this method without a detrimental effect on the efficiency of carcass production. The lambs in the experiment were given three daily intraperitoneal injections. The technique also works on pigs and has the added advantage of increasing muscle growth (Kestin et al., 1993).

The possibility of repartioning the flow of energy into parts of the body that are growing has been investigated with beta-agonists such as clenbuterol (Rick et al., 1984; Baker et al., 1984). The fatty acids that are released by beta-adrenergic stimulation of adipose tissue appear to be diverted towards the provision of energy for protein synthesis. Thus, muscle growth is enhanced and adipose tissue growth is reduced. The effect may be enhanced by a simultaneous reduction in protein degradation and an increase in serum STH levels. Clenbuterol and ractopamine both directly increase protein synthesis in muscle (Maltin et al., 1987; Anderson et al., 1990), and an increased association of ribosomes with myofibrils may occur with clenbuterol (Horne and Hesketh, 1990). On the other side of the balance between anabolism and catabolism in muscle, cimaterol and clenbuterol also reduce myofibrillar protein degradation (Béchet et al., 1990; Benson et al., 1991). Another approach to reducing adipose accumulation is the suppression of fatty acid synthase by inhibition of gene expression (Clarke, 1993), but how this might affect cellularity in terms of adipose cell numbers and size remains to be seen.

Pork

Newborn pigs have very little fat relative to other mammals. Between birth and 4 weeks of age, the percentage of fat increases from about 1% to about 18% of empty live weight (Manners and McCrea, 1963). Adipose tissue lobules in regions that will later deposit large amounts of fat contain few cells, relative to neonatal ruminants, and they are separated by areas of loose mesenchyme or undifferentiated tissue. Triglyceride deposition in adipose cells first becomes microscopically detectable at about the third month of gestation. After this time and for the remainder of gestation, the numbers of cells in each lobule show no great increase, although the numbers of lobules and the total numbers of adipose cells do increase (Hermans, 1973a). In pigs, fibroblast-like cells and gland-like cells are not the only source of adipose cells, since perirenal adipose cells may originate from endothelial cells of the vascular system (Desnoyers and Vodovar, 1977). Fat is deposited very rapidly around the kidneys, and perirenal fat has high enzyme activity, large adipose cells and a low connective fiber content (Anderson et al., 1972). The activities of lipoprotein lipase and hormone sensitive lipase increase markedly after birth (Steffen et al., 1978), and the pig then starts its life-long process of triglyceride accumulation. However, the partitioning of lipoprotein lipase activity between muscle and adipose tissue may have already been genetically determined before birth so that genetically lean pigs can divert more energy to their musculature (McNamara and Martin, 1982). If a pregnant sow is diabetic, its offspring may have an increased amount of body fat (Ezekwe and Martin, 1980).

The heritability of adipose deposition is quite high in pigs, typically about 50% for back fat thickness. Commercial selection for decreased backfat thickness produces a real reduction in total carcass fatness, not simply a redistribution of adipose tissue (Fortin and Elliot, 1985). However, the backfat of pigs is divided into two or three layers (depending on the position of measurement) and the different layers exhibit some degree of independence in their depth changes in response to selection. The deepest layer may be most responsive (Mersmann and Leymaster, 1984). Pigs also respond readily to selection for increased adipose deposition in medical experiments on obesity.

Lean breeds of pigs may have a greater ability than fat breeds to mobilize their fat, perhaps because of a greater release of norepinephrine from their sympathetic nerve endings (Gregory and Lister, 1981). Metz and Dekker (1981), however, have shown that genetic differences in the rate of fat deposition may occur without marked differences in fat mobilization. Obese pigs, for example, have about the same rate of lipolysis as normal lean pigs (Mersmann, 1985). Thus, plasma lipid concentration is a poor indicator of adiposity in pigs (Mersmann and MacNeil, 1985). Another possible mechanism for the genetic control of adiposity is the activity of lipoprotein lipase. Hausman et al. (1983) showed that fetal adipose cells of obese pigs react positively for lipoprotein lipase whereas cells from lean fetuses do not.

Anderson and Kauffman (1973) found that, up to 5 months after birth, subcutaneous adipose tissue grew by a combination of hyperplasia and hypertrophy. Growth after this time was almost completely from adipose cell hypertrophy. This was confirmed by Hood and Allen (1977) for both perirenal and extramuscular (basically subcutaneous) adipose tissue. Measurements of adipose cell numbers and diameters in this type of research are usually made on cells that have been released from their connective tissue framework. In histological sections, it is difficult to measure adipose cell diameters since most cells are cut off-center and have reduced cross sectional areas. Adipose cells are easily fragmented, and the extent of fragmentation may vary with experimental conditions. Fragments may round-up and form small globules, and without cytological examination these are difficult to distinguish from small adipose cells. Similarly, cells have to be captured once they have been released from their connective tissue framework, and different techniques vary in their efficiency. The methodological problems associated with measuring adipose cell size and number are reviewed by Mersmann and MacNeil (1986).

Bimodality in frequency histograms of adipose cell diameters may suggest that there are two populations of cells - large ones and small ones. The large ones might be mature adipose cells filled to capacity with triglyceride, and the small ones might be young adipose cells in the process of filling-up. On the basis of these possibilities, bimodality is often regarded as evidence for the formation of new adipose cells. Although this may be a correct assumption in many cases, other possibilities cannot be automatically excluded. A unimodal population might be continuously supplemented by new cells which are too few in number at any point in time to form their own peak in the frequency distribution of cell size. Over the animal's lifetime, however, this might be a major source of cells. Similarly, a bimodal population might be static, and the populations of large and small adipose cells might be related to an unrecognized factor. For example, adipose cells near to blood vessels might be larger than distant cells, or vice versa.

When Anderson and Kauffman (1973) measured the diameters of adipose cells from the back fat of Chester White barrows, they found unimodal populations in the frequency distribution of different sized cells. When Mersmann et al. (1973) attempted the same measurements with cells isolated from the neck region of crossbred pigs, they found that cell diameters formed bimodal populations. Wood et al. (1975) found that the rate of adipose cell hypertrophy in subcutaneous shoulder fat depended on the depth at which cells were taken from the fat. Carpenter et al. (1961) found considerable variation in adipose distribution along the length of the longissimus dorsi muscle. Enser et al. (1976) found differences in growth patterns between shoulder and midback adipose depots; hyperplasia persisted for longer in the shoulder depot, and new cells appeared to originate from near the muscle surface. Subcutaneous fat from the shoulder region of Large White pigs exhibits both hypertrophy and hyperplasia to at least 188 days of age (Wood et al., 1978). Etherton (1980) reported bimodal distributions for adipose cell diameters as late as one year of age in both lean and obese breeds. Histological differences exist between breeds, as in the case of Large Whites and Pietrains (Moody et al., 1978).

Kirtland and Gurr (1980) used a thymidine label to assess the extent of DNA synthesis associated with the formation of new cells in fat depots. Formation of new cells proceeded rapidly between 2 and 40 days of age, but beyond this time the growth of backfat layers was primarily due to the filling of pre-existing empty adipose cells.

Despite this confusion over the relative contributions of hyperplasia and hypertrophy, the case for the genetic regulation of adipose cell numbers and diameters is quite strong. Steele et al. (1974) measured adipose cell numbers and diameters in strains of the Duroc breed with either a high or a low backfat thickness, and found considerable differences, particularly in the contribution from hyperplasia. Breeds and strains of pigs differing in backfat thickness may also exhibit hormonal differences in fat mobilization (Standal et al., 1973; Steele and Frobish, 1976; Wood et al., 1977), but whether these differences can be meaningfully related to differences in cell numbers and diameters is a difficult question. What does it mean if the concentration of lipolytic enzymes is high, when expressed on the basis of tissue weight? In rats for example, the volume of cytoplasm per adipose cell decreases from 8% to 0.8% during cellular hypertrophy (Goldrick, 1967). Adipose cell cytoplasm is restricted to a thin layer beneath the plasma membrane and, as an adipose cell grows in size, the plasma membrane increases in area at a slower rate than the volume of the cell. Thus, if geometrical proportionality is maintained, cytoplasmic components will decline in their overall concentration in the tissue as cells grow in size.

The development of intramuscular adipose tissue in pork may depend on muscle structure and fascicular arrangement. Kauffman and Safanie (1967) proposed that muscles with long parallel fasciculi may deposit intramuscular fat more readily than pennate muscles with short fasciculi inserted at an angle to their tendon. The deposition of intramuscular fat is later than in subcutaneous, visceral and intermuscular locations, and it does not occur to any marked extent until 16 weeks in trapezius and semitendinosus muscles of Duroc and Hampshire pigs (Lee and Kauffman, 1974a). Thus, growth from this time onwards involves the formation of new adipose cells in these locations. The small mean diameters typically found among intramuscular adipose cells (Lee and Kauffman, 1974b) may be from recruitment of new small diameter cells while older cells continue to grow by hypertrophy. Lee et al. (1973a, 1973b) examined the effect of an early low plane of nutrition on adipose tissue development, and they compared animals at a standard age and at a standard weight. At 24 weeks, the reduced amount of intramuscular fat in pigs on a low plane of nutrition was caused by a decrease in cell size and cell number. Reduction in subcutaneous fat, however, was from a decrease in cell size but not in cell number. On a weight constant basis (80 kg), an early low plane of nutrition had little effect on subcutaneous adipose tissue growth, but there were reductions in adipose cell numbers and diameters intramuscularly. Similar results also were obtained by Campbell and Dunkin (1983) and it now appears unlikely that early postnatal nutrition can be used to manipulate or alter the amount of carcass fat in pigs reared under commercial conditions.

Beef

An extensive study of adipose tissue in bovine fetuses was undertaken by Bell in 1909 and, although his work is dated technically, many of his observations and conclusions are still valid. The morphological distinction between mesenchyme cells and adipose precursor cells was then, and still is, a rather subjective separation. Bell (1909) identified the first adipose precursor cells around the kidneys of 4.7 cm fetuses (approximately 50 days gestation). By 30 cm length (approximately 125 days gestation), masses of adipose cells were found in close association with blood vessels. By this time, adipose cells also had developed in sternal intermuscular and subcutaneous depots. An interesting point observed by Bell (1909) was that, in different anatomical sites, there was a considerable variation in the delay between the time of appearance of precursor cells and the time of triglyceride deposition. Perirenal adipose precursor cells showed a long delay while sternal cells showed a short delay. Bell (1909) sketchily followed adipose tissue development into the postnatal period, and he found that thin cattle had smaller subcutaneous adipose cells than fat cattle. He concluded that an increase in adipose cell numbers was partly responsible for the postnatal accumulation p carcass fat. Much the same view still prevails today (Allen, 1976).

In cattle, as in pigs, visceral adipose tissue develops before intramuscular adipose tissue. Adipose precursor cells make their greatest contribution to cell numbers viscerally, while the recruitment of fibroblast-like cells is more important intramuscularly. Hood and Allen (1973) found that hyperplasia was nearly complete in subcutaneous and perirenal depots by approximately 8 months. Intramuscularly, progressive cellular recruitment, as revealed by bimodality of cell diameters, still occurred at 14 months. Hood and Allen (1973) also found smaller diameter subcutaneous adipose cells in lean Holstein cattle, than in well finished Hereford x Angus crossbreds. Truscott et al. (1983) found that growth in the perirenal fat depots of Hereford and Fiesian steers was caused exclusively by cellular hypertrophy. In subcutaneous fat, however, hypertrophy was responsible for growth upto 13 months but, after this time, both hypertrophy and hyperplasia occurred together. A similar result was obtained by Cianzio et al. (1985) at a number of systematic locations in cross-bred beef steers. Adipose cell hypertrophy was primarily responsible for adipose tissue growth until 19 months, but an increase in cell numbers was found from 11 to 15 months in intramuscular fat and after 15 months in brisket fat.

Also it appears that the balance between adipose cell hypertrophy and hyperplasia might be influenced by animal nutrition. Prior (1982) found that steers fed alfalfa hay produced less total adipose tissue with larger cells than steers fed a high-energy ration. The animals on the high-energy ration produced more total adipose tissue but with smaller cells. It is likely that the difference created by this nutritional treatment is simply one of a static adipose cell population versus an actively growing population. Mean cell size might be kept low by the formation of new cells in the active population.

The subjective assessment of marbling fat is very important in the commercial grading of beef carcasses. However, it is difficult to relate the histological frequency and distribution of intramuscular adipose cells to marbling scores (Cooper et al., 1968; Moody and Cassens, 1968). Lipoprotein lipase distribution in bovine adipose cells is described by Plaas et al. (1978).

Lamb

In fetal lambs, Wensvoort (1967) found adipose cell precursors in 5 to 8 cm fetuses (approximately 44 to 52 days gestation). The initial storage of triglyceride occurred at 9 cm (approximately 55 days gestation). According to Wensvoort (1968), adipose tissue formation in fetal lambs differs from that in fetal calves. In fetal lambs, two extra types of cells are involved, pleo-protoplasmic and plurivacuolar cells. Pleo-protoplasmic cells are large polygonal cells which have central nuclei and abundant granulated cytoplasm without lipid droplets. Possibly, a morphological overlap between the developmental stages of white and brown adipose cells may account for this. Lambs have little prenatal development of subcutaneous fat, although it develops rapidly immediately after birth (Vezinhet and Prud'hon, 1975).

Burton et al. (1974) found that the amount of perirenal fat increased between 50 and 70 kg live weight in Suffolk ewes, and was caused by adipose cell hypertrophy without evidence of new cells being formed. Nougues and Vezinhet (1977) found the same in the perirenal fat of Merino sheep. Haugebak et al. (1974) examined the effect of maintenance and ad libitum early diets combined with either of two protein levels in a finishing diet. In general, adipose cell hypertrophy was found in all conditions that allowed fat deposition. An interesting exception was that sternal intermuscular adipose cells reached their final diameters at a much earlier time than in the rest of the carcass. In well fed animals, hyperplasia was complete by the time that the finishing diet was fed, whereas an early low plane of nutrition prolonged hyperplasia into the finishing period. Thus, in the growth of adipose tissue, it appears that the prolongation of hyperplasia may contribute to the mechanism of compensatory growth or catch-up growth. However, comparable studies on compensatory growth mechanisms by Burton et al. (1974) yielded no statistically valid evidence of changes in cell sizes and numbers. Nougues and Vezinhet (1977) found that hypertrophy coupled with hyperplasia persisted to 100 days in intermuscular and subcutaneous fat of Merino sheep. After 100 days, adipose tissue growth was solely due to hypertrophy. somatotropic hormone has a strong effect on adipose accumulation and distribution in sheep. Hypophysectomy causes an increase in fat deposition, and this can be reversed by administering STH (Vezinhet et al., 1974).

Poultry

Avian adipose cells differ from those of farm mammals since they have only a limited capacity for lipogenesis. Thus, they rely mainly on the capture of circulating lipids that have been synthesized in the liver or released by digestion in the gut. Brown fat is absent or at least difficult to locate in birds (Johnston, 1971). By day 11 or 12 of incubation, precise anatomical sites of future adipose tissue may be identified in the chick embryo (Liebelt and Eastlick, 1952) and the future distribution of subcutaneous fat depots is established (Liebelt and Eastlick, 1954). Evans (1977) concluded that hyperplasia continued after hatching until the onset of sexual maturity and that, beyond this time, adipose growth was due to hypertrophy. In ducks, Evans (1977) found a nine-fold increase in adipose cell volume between hatching and 8 weeks of age.

Among different species of birds, those with a large body size tend to have larger adipose cells than those with a smaller body size (Pond and Mattacks, 1985b). There are marked differences in adipose cell numbers and diameters between layer-type and broiler-type fowl, with the lean broiler-type birds having fewer and smaller cells (March and Hansen, 1977). These authors also found that an early restriction of nutrient intake, although it inhibited hypertrophy, had only a slight effect on adipose cell hyperplasia. Thus, adequate cell numbers for fat deposition were present in birds once they were returned to an ad libitum diet.

Growth of the abdominal fat pad in chickens is from a combination of adipose cell hyperplasia and hypertrophy, up to about 12 to 14 weeks, then it continues mainly by hypertrophy (Hood, 1982; March et al., 1984). Ballam and March (1979) found that restricted feed intake had little effect on the numbers of retroperitoneal adipose cell in broilers, but that there were many small cells and few large cells. Adipose cell numbers along the sartorius muscle were unaffected.

Different anatomical sites of adipose tissue sites may differ in their priority to obtain circulating lipid. In broilers, Cahaner et al. (1986) found that the order was mesenterial > abdominal > gizzard > sartorial = neck. Adipose growth in broilers may be modified with beta agonists such as cimaterol and Merkley and Cartwright (1989) found that adipose cell diameters were more sensitive to cimaterol treatment than were cell numbers (Figure 2-38). Selection for growth rate may have increased visceral adipose cell hypertrophy with little change in the number of cells (Cartwright et al., 1986).

PROBLEMS WITH ADIPOSE TISSUE

Boar taint

Pork from boar carcasses sometimes yields an unpleasant odor called boar taint. However, this is only detectable by a small percentage of consumers (Walstra, 1974). The odor is most often detected in meat from old boars that have been used for breeding, and it need not be a problem in boars which have been slaughtered at a relatively light weight. This enables pork producers in some countries to take advantage of the rapid growth of young males that have not been castrated. Entire males may have a 10 to 15% advantage in feed efficiency and carcasses may be several percent leaner. On the other hand, problems may be encountered with soft fat, skin damage from fighting and low curing yields.

Boar taint becomes noticeable when pork is cooked or when carcass fat is tested with a hot iron, and is caused by the concentration of sex steroids in the fat (Patterson, 1968; Beery et al., 1969). Patterson (1968) identified the major factor as 5 - androst - 16 - ene - 3 - one. This is commonly called androstenone and has an intense odor of urine. There are other testicular steroids in the 16-androstene family and these have a musk odor. Androstenone passes into the blood stream and is accumulated by adipose tissue and by parotid and submaxillary salivary glands (Gower, 1972). Androstenone is transmitted in the boar's breath or saliva to the sow during mating and acts as a chemical messenger or pheromone (Sink, 1967). The enthusiasm with which sows root out truffles from deep in the ground may be due to the fact that these underground mushrooms also produce androstenone (Claus et al., 1981). Jonsson and Wismer-Pedersen (1974) found that the intensity of boar taint was a heritable trait (h&S'2 = 0.54). Claus (1975) showed that boar taint may be supressed immunologically and this method has been advocated for inhibiting boar odor while still utilizing the greater leaness and growth efficiency of intact males (Brooks and Pearson, 1986; Brooks et al., 1986). Skatole and indole produced from the amino acid tryptophan in the gut also contribute to boar taint (Hansson et al., 1980) since they have a strong fecal odor.

Abnormal development of adipose tissue

Replacement of muscle fibers by adipose cells is a common end result of a number of pathological conditions that affect skeletal muscle. In most of these conditions there is usually some evidence of muscle fiber regeneration. In meat animals, however, it is not uncommon to find that muscle fibers have been replaced by adipose cells without any evidence of muscle fiber regeneration and with no decrease in overall muscle volume. The most appropriate name for this condition is muscular steatosis (Hadlow, 1962).

Muscular steatosis is most frequent in cattle and pigs, but it may also occur in sheep (Hartman and Shorland, 1957). Sometimes the occurrence of muscular steatosis is indicated before slaughter by an abnormal gait (Leipold et al., 1973a), but usually the condition is not found until a carcass is butchered. The dividing line between excessive marbling fat and muscular steatosis is sometimes difficult to establish, and it is often only the restriction of muscular steatosis to a single muscle or muscle group in an otherwise poorly marbled carcass that makes it conspicuous. Muscular steatosis sometimes occurs in conditions which suggest that it has been caused by muscle damage or denervation. Strenuous muscle exertion may cause extensive muscle damage, particularly in those muscles that are used when an animal rears up on its hindlegs. Link et al. (1967) found that muscular steatosis sometimes occurred in muscles from which biopsy samples had been taken. Naturally occurring muscular steatosis also has been linked to vascular abnormalities in muscle (MacKenzie, 1912; Nowicki and Zajac, 1964). Another possibility is that muscular steatosis is the end result of a lipid accumulation myopathy, similar to those which occur in man (Harriman and Reed, 1972; Johnson et al., 1973). In pigs, there is evidence that the onset of muscular steatosis is accompanied by lipid accumulation in muscle fibers (Allen et al., 1967; Montroni and Testi, 1965). However, steatosis can sometimes be a major problem when it affects a high proportion of pigs in a herd, and we do not yet have a proper scientific understanding of the problem.

Pigs sometimes develop nodular lesions in their subcutaneous and visceral adipose tissue. Nodular lesions of fat necrosis may also be found in the pancreas and in other visceral organs. Ito (1973) suggested that this condition is caused by the release of esterase from degenerating pancreatic cells, and elevated levels of esterase may be found in the adipose tissue of pigs with "yellow fat disease" (Danse and Steenbergen-Botterweg, 1974). Adipose degeneration in pigs usually is accompanied by a marked increase in yellow pigmentation, probably due to accumulation of ceroid pigment (Hermans, 1973b). In cattle, yellow carotenoid pigment normally accumulates in the healthy adipose tissues of grass-fed animals. The depth of coloration increases as animals become older. Yellow coloration of lamb fat is a normal result of certain types of nutrition and is caused by lutein (Kruggel et al., 1982), an alternative name for which is xanthophyll. Yellow discoloration of subcutaneous fat in lambs may also be caused by bilirubin. Brown discoloration has been attributed to a combination of translucency, heme pigment and peroxidation of unsaturated fats (Prache et al., 1990).

REFERENCES

Alexander,G., J.W. Bennett, and R.T. Gemmell. 1975. J.Physiol., 244:223.

Allen, C.E. 1976. Fed.Proc., 35:2302.

Allen, C.E., D.C. Beitz, D.A. Cramer, and R.G. Kauffman. 1976. "Biology of Fat in Meat Animals," North Central Regional Research Publication 234. University of Wisconsin, Madison.

Allen,E., R.G. Cassens, and R.W. Bray. 1967. J.Food Sci., 32:146.

Anderson, D.B. and R.G. Kauffman. 1973. J.Lipid Res., 14:160.

Anderson, D.B., R.G. Kauffman, and L.L. Kastenschmidt. 1972. J. Lipid Res., 13:593.

Anderson, P.T., W.G. Helferich, L.C. Parkhill, R.A. Merkel and W.G. Bergen. 1990. J. Nutr., 120:1677.

Archer, C.W., A. Hornbruch, and L. Wolpert. 1983. J. Embryol. Exp. Morphol., 75:101. Archer, C.W. and N.A. Ratcliffe. 1983. J. Exp. Zool., 225:243.

Ashwell, M.A., P. Priest, and C. Sowter. 1975. Nature, 256:724.

Bailey, A.J. and T.J. Sims. 1977. J. Sci. Food Agric., 28:565.

Bailey, A.J., G.B. Shellswell, and V.C. Duance. 1979. Nature 278:67.

Baker, P.K., R.H. Dalrymple, D.L. Ingle, and C.A. Ricks. 1984. J. Anim. Sci., 59:1256.

Ballam, G.C. and B.E. March. 1979. Poultry Sci., 58:940.

Barrineau, L.L., C.B. Rich, and J.A. Foster. 1981. Connect. Tissue Res., 8:189. Béchet, D.M., A. Listrat, C. Deval, M. Ferrara and J.F. Quirke. 1990. Am. J. Physiol. 259:E822. Beery, K.E., J.D. Sink, S. Patton, and J.H. Ziegler. 1969. J.Am.Oil Chem.Soc., 46:439A.

Behari, J. and W.H. Andrabi. 1978. Connect.Tissue Res., 6:181.

Bell, E.T. 1909. Amer.J.Anat., 9:412.

Bengtsson, S.G. and R.V.J. Hakkarainen. 1975. J. Anim. Sci., 41:106.

Benson, D.W., T. Foley-Nelson, W.T. Chance, F-S. Zhang, J.H. James and J.E. Fischer. 1991. J. Surg. Res., 50:1.

Bentz, H., H.P. Bachinger, R. Glanville, and K. Kuhn. 1978. Eur. J. Biochem., 92:563.

Berg, R.T., B.B. Andersen, and T. Liboriussen. 1978. Anim. Prod., 27:63.

Berg, R.T. and R.M. Butterfield. 1976. New Concepts of Cattle Growth, New York: John Wiley & Sons. Berg, R.T., S.D.M. Jones, M.A. Price, R. Fukuhara, R.M. Butterfield, and R.T. Hardin. 1979. Can. J. Anim. Sci., 59:359.

Berge, P., J. Culioli, M. Renerre, C. Touraille, D. Micol, and Y. Geay. 1993. Meat Sci., 35:79.

Berge, S. 1948. J. Anim. Sci., 7:233.

Berry, B.W. and K.F. Leddy. 1990. J. Anim. Sci., 68:666.

Birk, D.E. and R.L. Trelstad. 1986. J. Cell Biol., 103:231.

Blanchette-Mackie, E.J. and R.O. Scow. 1981. J. Ultrastruct. Res., 77:277.

Blood. D.C. 1956. Aust.Vet. J., 32:125.

Bock, P. 1977. Mikroskopie, 33:332.

Bock, P. 1978. Histochemistry, 55:269.

Bovard, K.P. and L.N. Hazel. 1963. J. Anim. Sci., 22:188.

Brannang, E. 1971a. Swedish J. Agric. Res., 1:69.

Brannang, E. 1971b. Swedish J. Agric. Res., 1:79.

Breidenstein, B.B., C.C. Cooper, R.G. Cassens, G. Evans, and R.W. Bray. 1968. J.Anim.Sci., 27:1532.

Brissie, S.S. Spicer, and N.T. Thompson. 1975. Anat.Rec., 181:83.

Brooks, R.I. and A.M. Pearson. 1986. J. Anim. Sci., 62:632.

Brooks, R.I., A.M. Pearson, M.G. Hogberg, J.J. Perstka, and J.I. Gray. 1986. J. Anim. Sci., 62:1279.

Bullard, H.H. 1912. Am. J. Anat., 14:1.

Burgeson, R.E., F.A. El Adli, I.I. Kaitila, and D.W. Hollister. 1976. Proc. Natl. Acad. Sci., 73:2579.

Burson, D.E. and M.C. Hunt. 1986a. Meat Sci., 17:153.

Burson, D.E. and M.C. Hunt. 1986b. J. Food Sci., 51:51.

Burton, J.H., M. Anderson, and J.T. Reid, 1974. Br.J.Nutr., 32:515.

Busboom, J.R., G.J. Miller, R.A. Field, J.D. Crouse, M.L. Riley, G.E. Nelms, and C.L. Ferrell, 1981. J. Anim. Sci., 52:83.

Cahaner, A., Z. Nitsan, and I. Nir. 1986. Poultry Sci., 65:215.

Campbell, R.G. and A.C. Dunkin. 1983. Br. J. Nutr., 49:109.

Canalis, E., T. McCarthy, and M. Centrella. 1988. Calcif. Tissue Int., 43:346.

Carnes, W.H. 1971. Fed.Proc., 30:995.

Carpenter, Z.L., R.W. Bray, E.J. Briskey, and D.H. Traeder. 1961. J.Anim.Sci., 20:603.

Cartwright, A.L., H.L. Marks, and D.R. Campion. 1986. Poultry Sci., 65:1021.

Charles, D.D. and E.R. Johnson. 1976. J. Anim. Sci., 42:332.

Chen, S., L.C. Teicher, D. Kazim, R.E. Pollack, and L.S. Wise. 1989. Science, 244:582.

Cianzio, D.S., D.G. Topel, G.B. Whitehurst, D.C. Beitz, and H.L. Self. 1982. J. Anim. Sci., 55:305.

Cianzio, D.S., D.G. Topel, G.B. Whitehurst, D.C. Beitz, and H.L. Self, 1985. J. Anim. Sci., 60:970.

Clark, E.R. and E.L. Clark. 1940. Amer.J.Anat., 67:255.

Clarke, S.D. 1993. J. Anim. Sci., 71:1957.

Claus, R. 1975. "Neutralization of pheromones by antisera in pigs," in Immunization with Hormones in Reproductive Research, E. Nieschlag, ed., North Holland Publishing Company, Amsterdam, pp. 189-197. Claus, R., H.O. Hoppen, and H. Karg. 1981. Experientia, 37:1178.

Cook, L.J., T.W. Scott, K.A. Ferguson, and I.W. McDonald, 1970. Nature, Lond. 228:178.

Cooper, C.C., B.B. Breidenstein, R.G. Cassens, G. Evans, and R.W. Bray. 1968. J.Anim.Sci., 27:1542.

Cooper, S. and M.H. Gladden. 1974. Quart.J.Exp.Biol.Physiol., 59:367.

Cox, R.W., W.M.F. Leat, D. Chauca, M.A. Peacock, and J. Bligh. 1978. Res. Vet. Sci., 25:58.

Craig, A.S. and D.A.D. Parry. 1981. Proc. Roy. Soc. Lond. B., 212:85.

Crenshaw, T.D., E.R. Peo, A.J. Lewis, and B.D. Moser. 1981. J. Anim. Sci., 53:827.

Crowe, M.W. 1969. Med. Vet. Pract., 50(13):54.

Crowe, M.W. and H.T. Pike. 1973. J.Am. Vet. Med. Assoc., 162:453.

Danse, L.H.J.C. and W.A. Steenbergen-Botterweg. 1974. Vet.Path., 11:465.

Dardick, I., W.J Poznanski, I. Waheed, and G. Setterfield. 1976. Tissue and Cell, 8:561.

Dauncey, M.J., F.B.P. Wooding, and D.L. Ingram. 1981. Res. Vet. Sci., 31:76.

Davies, A.S. 1975. Anim.Prod., 20:45.

Desnoyers, F., G. Durand, and N. Vodovar. 1980. Biol. Cell., 38:195.

Desnoyers, F. and N. Vodovar. 1977. Biol.Cell., 29:177.

Detwiler, S.R. 1934. J. Exp. Zool., 67:395.

Dolezal, H.G., C.E. Murphey, G.C. Smith, and Z.L. Carpenter. 1982. Meat Sci., 6:55.

Doornenbal, H. 1975. Growth, 39:427.

Dransfield, E. and R.C.D. Jones. 1984. J. Food Technol., 19:181.

Drew, I.M., D. Perkins, and P. Daly. 1971. Science 171:280.

Duance, V.C., D.J. Restall, H. Beard, F.J. Bourne, and A.J. Bailey. 1977. FEBS Letters, 79:248.

Duance, V.C., S.F. Wotton, C.A. Voyle, and A.J. Bailey. 1984. Biochem. J., 221:885.

Duff, S.R.I. 1987. J. Comp. Pathol., 97:41.

Dyer, H. McM. and B.J.S. Pirie. 1978. J. Anat., 125:519.

Edmonds, L.D., L.A. Selby, and A.A. Case. 1972. J. Am. Vet. Med. Assoc., 160:1319. Edwards, D.A. 1946. J. Anat., 80:147.

Elliot, J.I. and C.E. Doige. 1973. Can. J. Anim. Sci., 53:211.

Enser, M.B., J.D. Wood, D.J. Restall, and H.J.H. MacFie. 1976. J.Agric.Sci.,Camb., 86:633.

Etherington, D.J. and T.J. Sims. 1981. J. Sci. Food Agric., 32:539.

Etherton, T.D. 1980. Growth 44:182.

Evans, A.J. 1972. Br.Poultry Sci., 13:615.

Evans, A.J. 1977. "The Growth of Fat," in Growth and Poultry Meat Production, K.N. Boorman and B.J. Wilson, eds., Edinburgh: British Poultry Science, pp.29-64.

Ezekwe, M.O. and R.J. Martin. 1980. Horm. Metab. Res., 12:136.

Faust, I.M., P.R. Johnson, and J. Hirsch. 1977. Science, 197:393.

Faust, I.M., P.R. Johnson, J.S. Stern, and J. Hirsch. 1978. Am. J. Physiol., 235:E279.

Fell, H.B. 1956. "Skeletal Development in Tissue Culture," in The Biochemistry and Physiology of Bone, G.H. Bourne, ed., New York: Academic Press, chapter 14, pp. 401-441. Ferrier, J., S.M. Ross, J. Kanehisa, and J.E. Aubin. 1986. J. Cell Physiol., 129:283.

Field, R.A., G. Maiorano, F.C. Hinds, W.J. Murdoch, and M.L. Riley. 1990. J. Anim. Sci., 68:3663.

Finerty, M. 1981. J. Theor. Biol., 93:279.

Floridi, A., E. Ippolito, and F. Postacchini. 1981. Connect. Tissue Res., 9:95. Fortin, A. and J.I. Elliot. 1985. J. Anim. Sci., 61:158.

Fortin, A., J.T. Reid, A.M. Maiga, D.W. Sim, and G.H. Wellington. 1981. J. Anim. Sci., 53:982.

Franzen, L. and K. Norrby. 1980. Cell Tissue Kinet., 13:635.

Furth, A. 1988. New Scientist, 117 (1602): 58.

Garton, G.A. 1967. World Rev. Nutr. Dietetics, 7:225.

Garton, G.A. 1976. "Physiological Significance of Lipids," in Meat Animals: Growth and Productivity, D.N. Rhodes, V.R. Fowler and M.F. Fuller, eds., New York: Plenum Press, pp. 159-176.

Gay, S. and E.J. Miller. 1978. Collagen in the Physiology and Pathology of Connective Tissue, Stuttgart: Gustav Fischer Verlag. Gemmell, R.T., A.W. Bell, and G. Alexander. 1972. Amer.J.Anat., 133:143.

Gersh, I. and M.A. Still. 1945. J.Exp.Med., 81:219.

Gilbert, R.P., D.R.C. Bailey, and N.H. Shannon. 1993. J. Anim. Sci., 71:1712.

Goldrick, R.B. 1967. Am. J. Physiol., 212:777.

Gotoh, T., Y. Sugi, and R. Hirakow. 1983. J. Electron Microsc., 32:213.

Gotta-Pereira, G., F.G. Rodrigo, and J.F. David-Ferreira. 1976. Stain Tech., 51:7.

Gower, D.B. 1972. J. Steroid Biochem., 3:45.

Greeley, R.G., C.L. Boyd, and D.G. Jolly. 1968. SWest Vet., 21:277.

Greeley, R.G. and D.G. Jolly. 1968. SWest Vet., 21:189.

Gregory, N.G. and D. Lister. 1981. Proc. Nutr. Soc., 40:11A.

Gregory, N.G. and O.P. Whelehan. 1983. J. Sci. Food Agric., 34:1397.

Hadlow, W.J. 1962. "Diseases of Skeletal Muscle," in Comparative Neuropathology, J.R.M. Innes and L.Z. Saunders, eds., New York: Academic Press, pp. 147-243. Hall, A.D. 1905. The Book of the Rothamsted Experiments. John Murray, London.

Hall, J.B. and M.C. Hunt. 1982. J. Anim. Sci., 55:321.

Hallqvist, C. 1933. Hereditas, 18:215.

Hammond, J., I.L. Mason, and T.J. Robinson. 1971. Hammond's Farm Animals, 4th Edition, London: Edward Arnold, p. 89. Hansson, K-E., K. Lundstrom, S. Fjelkner-Modig, and J. Persson. 1980. Swedish J. Agric. Res., 10:167.

Harcourt, R.A. and R. Bradley. 1973. Vet. Rec., 92:233.

Hargis, P.S. and M.E. Van Elswyk. 1993. World's Poultry Sci. J., 49:251.

Harriman, D.G. and R. Reed. 1972. J.Path., 106:1.

Harrison, T.J. 1975. J. Anat., 120:625.

Hartman, A.D. 1980. Am. J. Physiol., 241:E108.

Hartman, L. and F.B. Shorland. 1957. J.Sci.Food Agric., 8:428.

Haugebak, C.D., H.B. Hedrick, and J.M. Asplund. 1974. J.Anim.Sci., 39:1016.

Hausman, G.J. 1989. J. Anim. Sci., 67:3136.

Hausman, G.J., D.R. Campion, and G.B. Thomas. 1983. J. Lipid Res., 24:223.

Hausman, G.J. and R.J. Martin. 1981. J. Anim. Sci., 52:1442.

Heine, H. and F.J. Forster. 1974. Acta Anat., 89:387.

Helliwell, T.R., O. Gunhan, and R.H.T. Edwards. 1990. J. Neurol. Sci., 98:267. Hendricks, H.B., E.D. Aberle, D.J. Jones, and T.G. Martin. 1973. J. Anim. Sci., 37:1305.

Hermans, P.G.C. 1973a. Tijdschr.Diergeneesd., 98:662.

Hermans, P.G.C. 1973b. Tijdschr.Diergeneesd., 98:668.

Hindmarsh, W.L. 1937. Vet. Res.Rep.,NSW., 7:58.

Hiner, R.L., E.E. Anderson, and C.R. Fellers. 1955. Food Technol., 9:80.

Hogg, D.A. 1982. J. Anat., 135:501.

Hogg, A., R.F. Ross, and D.F. Cox. 1975. Am. J.Vet. Res., 36:965.

Hood, R.L. 1982. Poultry Sci., 61:117.

Hood, R.L. and C.E. Allen. 1973. J. Lipid Res., 14:605.

Hood, R.L. and C.E. Allen. 1977. J. Lipid Res., 18:275.

Horne, Z., and J. Hesketh. 1990. Biochem. J. 272:831.

Howlett, C.R. 1980. J. Anat. 130:745.

Hutt, F.B. 1934. J.Hered., 25:41.

Inoue, S. and C.P. Leblond. 1986. Am. J. Anat., 176:121.

Inoue, S. and C.P. Leblond. 1988. Am. J. Anat., 181:341.

Irie, M. and K. Ohmoto. 1982. Jap. J. Swine Sci., 19:165.

Irie, M. and H.J. Swatland. 1992. Asian Australasian J. Anim. Sci., 5:753.

Ito, T. 1973. Jap.J.Vet.Sci., 35:299.

James, L.F., J.L. Shupe, W. Binns, and R.F. Keeler. 1967. Am. J.Vet. Res., 28:1379.

James, T. 1951. Edinb.Med. J., 58:565.

John, H.A. and H. Lawson. 1980. Cell Biol. Int. Rep., 4:841.

Johnson, M.A., J.J. Fulthrope, and P. Hudgson. 1973. Acta Neuropath., 24:97.

Johnston, D.W. 1971. Comp. Biochem. Physiol., 40A:1107.

Johnston, W.G. and G.B. Young. 1958. Vet. Rec., 70:1219.

Jones, S.D.M., M.A. Price, and R.T. Berg, 1978. Can. J. Anim. Sci., 58:151.

Jonsson, P. and J. Wismer-Pedersen. 1974. Livestock Prod. Sci., 1:53.

Kauffman, R.G. and A.H. Safanie. 1967. J.Food Sci., 32:283.

Kaye, M.M. 1962. Can. J. Comp. Med., 26:218.

Keeler, R.F. 1984. J. Anim. Sci., 58:1029.

Keeler, R.F., W. Binns, L.F. James, and J.L. Shupe. 1969. Can. J. Comp. Med., 33:89.

Keeler, R.F., L.F. James, W. Binns, and J.L. Shupe. 1967. Can. J. Comp. Med., 31:334.

Keene, D.R., L.Y. Sakai, H.P. Bachinger, and R.E. Burgeson. 1987. J. Cell Biol., 105:2393.

Keene, D.R., E. Engvall, and R.W. Glanville. 1988. J. Cell. Biol., 107:1995.

Kempster, A.J. and D.G. Evans. 1979. J. Agric. Sci., Camb., 93:349.

Kestin, S., R. Kennedy, E. Tonner, M. Kiernan, A. Cryer, H. Griffin, S. Butterwith, S. Rhind, and D. Flint. 1993. J. Anim. Sci., 71:1486.

King, J.W.B. and R.C. Roberts. 1960. Anim. Prod., 2:59.

Kirtland, J. and M.I. Gurr. 1980. J. Agric. Sci., Camb., 95:325.

Kirtland, J., M.I. Gurr, G. Saville, and E.M. Widdowson. 1975. Nature, 256:723.

Koch, P., H. Fischer, and H. Schumann. 1957. Erbpathologie der Landwirtschaftlichen Haustiere, Berlin and Hamburg: Paul Parey, pp.189-190. Kuhn, K. and R.W. Glanville. 1980. "Molecular Structure and Higher Organization of Different Collagen Types," in Biology of Collagen, A. Viidik and J. Vuust, eds., New York: Academic Press, pp. 1-14.

Lakes, R. and S. Saha. 1979. Science, 204:501.

Laster, D.B. 1974. J. Anim. Sci., 38:496.

Latimer, H.B. 1927. Am. J. Anat., 40:1.

Laurent, G.J. 1982. Biochem. J., 206:535.

Laurie, G.W., C.P. Leblond, I. Cournil, and G.R. Martin, 1980. J. Histochem. Cytochem., 28:1267.

Lawrie, R.A. 1974. Meat Science., Oxford: Pergamon Press.

Le Gros Clark, W.E. 1945. The Tissues of the Body., Oxford: Clarendon Press.

Leach, R.M. 1971. Fed.Proc., 30:991.

Leat, W.M.F. 1975. J. Agric. Sci., Camb., 85:551.

Leat, W.M.F. 1976. "The Control of Fat Absorption, Deposition and Mobilization in Farm Animals," in Meat Animals: Growth and Productivity, V.R. Fowler and M.F. Fuller, eds., New York: Plenum Press, pp. 177-193.

Leat, W.M.F. 1977. J. Agric. Sci., Camb. 89:575.

Lee, Y.B. and R.G. Kauffman. 1974a. J.Anim.Sci., 38:532.

Lee, Y.B. and R.G. Kauffman. 1974b. J.Anim.Sci., 38:538.

Lee, Y.B., R.G. Kauffman, and R.H. Grummer. 1973a. J.Anim.Sci., 37:1312.

Lee, Y.B., R.G. Kauffman, and R.H. Grummer. 1973b. J.Anim.Sci., 37:1319.

Leipold, H.W., B. Blaugh, K. Huston, C.G.M. Edgerly, and C.M. Hibbs. 1973a. Vet.Med.Small Anim.Clin., 68:645.

Leipold, H.W., Cates, W.F., Radostits, O.M. and W.E. Howell. 1969. Can. Vet. J., 10:268.

Leipold, H.W., F.W. Oehme, and J.E. Cook. 1973b. J.Am. Vet. Med. Assoc., 162:1059. Liebelt, R.A. and H.L. Eastlick. 1952. Anat. Rec., 112:422.

Liebelt, R.A. and H.L. Eastlick. 1954. Poultry Sci., 33:169.

Light, N. and A.E. Champion. 1985. Biochem. J., 219:1017.

Light, N., A.E. Champion, C. Voyle, and A.J. Bailey. 1985. Meat Sci., 13:137.

Link, B.A., R.W. Bray, R.G. Cassens, and R.G. Kauffman. 1970. J.Anim.Sci., 30:726.

Link, B.A., R.G. Cassens, R.W. Bray, and T. Kowalczyk. 1967. J. Anim. Sci., 26:694.

Lipton, B.H. 1977. Dev. Biol., 61:153.

Luke, D.A., C.H. Tonge, and D.J. Reid. 1980. J. Anat., 130:859.

Lwebuga-Mukasa, J.S., S. Lappi, and P. Taylor. 1976. Biochemistry, 15:1425.

MacKenzie, L.E. 1912. Virchows Arch.Path.Anat.Physiol., 210:57.

Maltin, C.A., S.M. Hay, M.I. Delday, F.G. Smith, G.E. Lobley and P.J. Reeds. 1987. Biosci. Rep. 7:525.

Mandl, I., J. Cantor, M. Osman, and G.M. Turino, 1986. Conn. Tiss. Res., 15:9. Manners, M.J. and M.R. McCrea. 1963. Brit. J. Nutr., 17:495.

March, B.E. and G. Hansen. 1977. Poultry Sci., 56:886.

March, B.E., C. MacMillan, and S. Chu. 1984. Poultry Sci., 63:2207.

Marlowe, T.J. 1964. J. Anim. Sci., 23:454.

Martin, R., J. White, J. Herbein, and M.O. Ezekwe. 1979. Growth, 43:167.

Martin, G.R., R. Timpl, P.K. Muller, and K. Kuhn. 1985. Trends Biochem. Sci., July. pp. 1-3.

May, S.G., C.A. Sturdivant, D.K. Lunt, R.K. Miller, and S.B. Smith. 1993. Meat Sci., 35:289.

Mayne, R. and K.R. Strahs. 1974. J. Cell Biol., 63:212a.

McCullough, A.W. 1944. J. Morphol., 75:193.

McNamara, J.P. and R.J. Martin. 1982. J. Anim. Sci., 55:1057.

Merley, J.W. and A.L. Cartwright. 1989. Poultry Sci., 68:762.

Mersmann, H.J. 1985. J. Anim. Sci., 60:131.

Mersmann, H.J. and K.A. Leymaster. 1984. Growth, 48:321.

Mersmann, H.J. and M.D. MacNeil. 1985. J. Anim. Sci., 61:122.

Mersmann, H.J. and M.D. MacNeil. 1986. J. Anim. Sci., 62:980.

Mersmann, H.J., J.R. Goodman, and L.J. Brown. 1975. J.Lipid Res., 16:269.

Mersmann, H.J., M.C. Underwood, L.J. Brown, and J.M. Houk. 1973. Am.J.Physiol., 224:1130.

Metz, S.H.M. and R.A. Dekker. 1981. Anim. Prod., 33:149.

Meyer, W., K. Neurand, and B. Radke. 1982. J. Anat., 134:139.

Michna, H. 1988. J. Anat., 158:1.

Middleton, D.S. 1932. Edinb.Med. J., 39:389.

Middleton, D.S. 1934. Edinb.Med. J., 41:401.

Miles, C.A., G.A.J. Fursey, and R.C.D. Jones. 1985. J. Sci. Food Agric., 36:215. Miller, A. 1982. Trends Biochem. Sci., 7(1):13.

Miller, E.J. 1985. "Recent Information on the Chemistry of the Collagens," in: The Chemistry and Biology of Mineralized Tissues, W.T. Butler, ed., Birmingham: EBSCO Media.

Miller, R.C., T.B. Keith, M.A. McCarty, and W.T.S. Thorp. 1940. Proc. Soc. Exp. Biol. Med., 45:50.

Montagna, W. 1945. J. Morphol., 76:87.

Montroni, L. and F. Testi. 1965. Acta Med.Vet.,Napoli., 11:505.

Moody, W.G. and R.G. Cassens. 1968. J.Food Sci., 33:47.

Moody, W.G., M.B. Enser, J.D. Wood, D.J. Restall, and D. Lister. 1978. J.Anim.Sci., 46:618.

Moody, W.G. and S.E. Zobrisky. 1966. J. Anim. Sci., 25:809.

Morley, F.H.W. 1954. Aust.Vet. J., 30:237.

Morrill, C.C. 1947. N.Am. Vet., 28:738.

Moss, M.L. 1972. "The Regulation of Skeletal Growth," in Regulation of Organ and Tissue Growth, R.J.Goss, ed., New York: Academic Press, pp.127-142. Motta, P. 1975. J.Microscopie, 22:15.

Munz,K. and C. Meves. 1974. Histochemistry, 40:181.

Murphy, P.A., E.D. Weavers, and J.N. Barrett. 1975. Vet. Rec., 97:445.

Murray, D.M., J.P. Bowland, R.T. Berg, and B.A. Young. 1974. Can. J. Anim. Sci., 54:91.

Nakano, T., F.X. Aherne, J.J. Brennan, and J.R. Thompson. 1984. Can. J. Anim. Sci., 64:139.

Nakano, T., J.R. Thompson, and F.X. Aherne. 1985. Can. Inst. Food Sci. Technol., J. 18:100. Nassar, A.H. and C.Y. Hu. 1991. J. Anim. Sci., 69:578.

Nechad, M. and T. Barnard. 1979. Biol. Cellulaire, 36:43. Nes, N. 1953. Nord.VetMed., 5:869.

Neville, W.E., B.G. Mullinix, J.B. Smith, and W.C. McCormick. 1978. J. Anim. Sci., 47:1080.

Nicholls, D.G. 1983. Biosci. Rep., 3:431.

Nisbet, D.I. and C.C. Renwick. 1961. J. Comp. Pathol., 71:177.

Nougues, J. and A. Vezinhet. 1977. Ann. Biol. Anim. Biochem. Biophys., 17:799.

Nowicki, L. and H. Zajac. 1964. Medycyna Wet., 20:279.

Oberbauer, A.M., W.B. Currie, L. Krook, and M.L. Thonney. 1989. J. Anim. Sci., 67:3124.

Odetti, P.R., A. Borgoglio, and R. Rolandi. 1992. Metabolism, 41:655.

Odetti, P., M.A. Pronzato, G. Noberasco, L. Cosso, N. Traverso, D. Cottalasso and U. Marinari. 1994. Lab. Invest., 70: 61. Oghiso, Y., Y-S. Lee, R. Takahashi, and K. Fujiwara. 1977. Jap. J. Vet. Sci., 39:101.

Oka, M., T. Miki, H. Hama, and T. Yamamuro. 1979. Clin. Orthop., 145:264.

O'Neill, I.K., M.L. Trimble, and J.C. Casey. 1979. Meat Sci., 3:223.

Oscai, L.B., D.A. Essig, and W.K. Palmer, 1990. J. Appl. Physiol., 69:1571.

Palludan, B. 1961. Acta Vet. Scand., 2:32.

Palsson, H. 1940. J. Agric. Sci., Camb., 30:1.

Palsson, H. and J.B. Verges. 1965. J. Agric. Sci., Camb., 64:247.

Park, R.J., R.A. Spurway, and J.L. Wheeler. 1972. J. Agric. Sci., Camb., 78:53. Parry, D.A.D., G.R.G. Barnes, and A.S. Craig. 1978a. Proc.R.Soc.Lond.B., 203:305. Parry, D.A.D., A.S. Craig, and G.R.G. Barnes. 1978b. Proc.R.Soc.Lond.B., 203:293.

Partridge, S.M. 1966. "Elastin," in The Physiology and Biochemistry of Muscle as a Food, E.J.Briskey, R.G.Cassens and J.C. Trautman, eds., Madison: University of Wisconsin Press, pp.327-339. Patten, B.M. 1931. The Embryology of the Pig. 2nd Editition, P. Blakiston's Son, Philadelphia.

Patterson, R.L.S. 1968. J.Sci.Food Agric., 19:31.

Patton, J.S. and M.C. Carey. 1979. Science, 204:145.

Pike, B.V. and C.J. Roberts. 1980. Res. Vet. Sci., 29:108.

Pimental, E.R. 1981. Acta Histochem. Cytochem., 14:35.

Plaas, H.A.K., R. Harwood, and A. Cryer. 1978. Biochem. Soc. Trans., 6:596.

Pond, C.M. and C.A. Mattacks. 1985a. J. Morphol., 185:183. Pond, C.M. and C.A. Mattacks. 1985b. J. Morphol., 185:195.

Pond, W.G., E.F. Walker, and D. Kirtland. 1975. J. Anim. Sci., 41:1053.

Poznanski, W.J., I. Waheed, and R. Van. 1973. Lab.Invest., 29:570.

Prache, S., B. Aurousseau, M. Theriez, and M. Renerre. 1990. INRA Prod. Anim., 3:275.

Preston, R.L., R.D. Vance, V.R. Cahill, and S.W. Kock. 1974. J. Anim. Sci., 38:47.

Preston, T.R. and M.B. Willis. 1974. Intensive Beef Production, Oxford: Pergamon Press.

Prior, R.L. 1983. J. Anim. Sci., 56:483.

Prockop, D.J. 1971. Fed.Proc., 30:984.

Puchtler, H., S.N. Meloan, and G.R. Pollard. 1976. Histochemistry, 49:1.

Puchtler, H. and F.S. Waldrop. 1978. Histochemistry, 57:177.

Puchtler, H., F.S. Waldrop, and L.S. Valentine. 1973. Histochemie, 35:17.

Ralston, A.T., W.H. Kennick, and T.P. Davidson. 1975. J. Anim. Sci., 40:1211.

Ramsay, T.G., G.J. Hausman, and R.J. Martin. 1987. J. Anim. Sci., 64:735.

Ramsay, T.G., M.E. White, and C.K. Wolverton. 1989. J. Anim. Sci., 67:2222.

Ramshaw, J.A.M. 1986. Connective Tiss. Res., 14:307.

Reale, E., F. Benazzo, and A. Ruggeri. 1981. J. Submicrosc. Cytol., 13:135.

Reineccius, G.A. 1979. J.Food Sci., 44:12.

Renk, B.Z., R.G. Kauffmann, and D.M. Schaefer. 1985. J. Anim. Sci., 61:876.

Ricks, C.A., R.H. Dalrymple, P.K. Baker, and D.L. Ingle. 1984. J. Anim. Sci., 59:1247.

Roberts, C.J., B.A. Turfrey, and A.P. Bland. 1983. Vet. Pathol., 20:23.

Roberts, J.A.F. 1929. J.Genet., 21:57.

Romhanyi, G. 1983. Histochemistry, 77:133.

Ross, R. and P. Bornstein. 1971. Sci.Am., 224:44.

Rowe, R.W.D. 1978. Meat Sci. 2:275.

Rowe, R.W.D. 1986. Meat Sci. 17:293.

Rucker, R.B. and M. Lefevre. 1980. "Chemical Changes in Elastin as a Function of Maturation," in Chemical Deterioration of Proteins, J.R. Whitaker and M. Fujmaki, eds., Amer. Chem. Soc., Washington, DC. Symp. Ser. 123: 63-82.

Sage, H. 1983. Comp. Biochem. Physiol., 74B:373.

Salans, L.B., M.J. Zarnowski, and R. Segal. 1972. J. Lipid Res. 13:616.

Sánchez-Mejorada, G. and F. Alonso de Florida. 1992. Muscle Nerve, 15:716.

Sanes, J.R. and J.M. Cheney. 1982. J. Cell Biol., 93:442.

Sasse, J., H. von der Mark, U. Kuhl, W. Dessau, and K. von der Mark. 1981. Dev. Biol., 83:79.

Schmalstieg, R. von and H. Meyer. 1960. Dt.tierarztl.Wschr., 67:41.

Scott, R.A., S.G. Cornelius, and H.J. Mersmann. 1981. J. Anim. Sci., 53:977.

Selby, L.A., R.W. Menges, E.C. Houser, R.E. Flatt, and A.A. Case. 1971. Arch.Environ.Health, 22:496.

Sellier, P. and R. Boccard. 1971. Ann. Genet. Sel. Anim., 3:433.

Selye, H. 1965. The Mast Cells., Butterworths, London.

Serafini-Fracassini, A., J.M. Field, and J. Hinnie. 1978. J. Ultrastruct. Res., 65:190.

Shupe, J.L., W. Binns, L.F. James, and R.F. Keeler. 1967. J.Am. Vet. Med. Assoc., 151:198.

Shaw, A.M. 1929. Sci. Agr., 10:29.

Shaw, A.M. 1930. Sci. Agr., 10:690.

Sidman, R.L. 1956. Anat. Rec., 124:581.

Simpson, J. W. and A.C. Taylor. 1974. Proc. Soc. Exp. Biol. Med., 145:42.

Sink, J.D. 1967. J. Theor. Biol., 17:174.

Sink, J.D. 1979. J.Food Sci., 44:1.

Sisson, S. and J.D. Grossman. 1953. The Anatomy of the Domestic Animals. 4th Edition, Philadelphia: W.B. Saunders.

Skipitaris, C.N. 1981. J. Agric. Sci., Camb., 97:83.

Slavin, B.G. 1979. Anat.Rec. 195:63.

Check this Smith reference: ?????

Smith, C., E.H. Samson, H.A. Padykula, M.G. Laskey, and S.H. Smith, and M.D. Judge. 1991. J. Anim. Sci., 69:1989. Loewenthal, L.A. 1952. J. Morphol. 90:103.

Stamp, J.T. 1960. J. Comp. Pathol., 70:296.

Standal, N., E. Vold, O. Trygstad, and I. Foss. 1973. Anim.Prod., 16:37.

Steele, N.C. and L.T. Frobish. 1976. Growth, 40:369.

Steele, N.C., L.T. Frobish, and M. Keeney. 1974. J.Anim.Sci., 39:712.

Steffen, D.G., L.J. Brown, and H.J. Mersmann. 1978. Comp. Biochem. Physiol., 59B:195.

Stingl, J. and O. Stembera. 1974. Lymphology, 7:160.

Sullivan, T.W. and Y.Y. Al-Ubaidi. 1963. Poul. Sci., 42:46.

Svoboda, E.L.A., T.P. Howley, and D.A. Deporter. 1983. Connect. Tissue Res., 12:43.

Swatland, H.J. 1974. Vet. Bull., 44:279.

Swatland, H.J. 1975. J.Anim.Sci., 41:78.

Swatland, H.J. 1978. Zbl. Vet. Med. A., 25:556.

Swatland, H.J. 1987a. J. Anim. Sci., 64:1038.

Swatland, H.J. 1987b. J. Anim. Sci., 65:158.

Swatland, H.J. 1987c. J. Food Sci., 52:865.

Swatland, H.J. 1987d. Histochem. J., 19:276.

Swatland, H.J. 1987e. Meat Sci., 19:277.

Swatland, H.J. 1989. J. Food Sci., 55:305.

Swatland, H.J. 1990. J. Comput. Assist. Microsc., 2:125.

Tavassoli, M. 1976. Acta Anat., 94:65.

Terrell, R.N., G.G. Suess, and R.W. Bray. 1969. J.Anim.Sci., 28:449

Thompson, G.E. and D.M. Jenkinson. 1969. Can.J.Physiol.Pharmacol., 47:249.

Thompson, G.E. and D.M. Jenkinson. 1970. Res.Vet.Sci., 11:102.

Thorngren, K-G. and L.I. Hansson. 1981. Acta Anat., 110:121.

Thurley, D.C. 1969. Path.Vet., 6:217.

Trelstad, R.L. 1982. Cell, 28:197.

Trelstad, R.L. and K. Hayashi. 1979. Develop. Biol., 71:228.

Truscott, T.G., J.D. Wood, and H.R. Denny. 1983. J. Agric. Sci., Camb., 100:271. Tuff, P. 1948. Skand.VetTidskr., 38:379.

Turner, J.C. 1979. Comp.Biochem.Physiol., 62A:599.

Tzaphlidou, M. 1986. Micron Microscop., Acta, 17:201.

Van, R.L.R., C.E. Baylisss, and D.A.K. Roncari. 1976. J.Clin.Invest., 58:699.

Van, R.L.R. and D.A.K. Roncari. 1977. Cell Tissue Res., 181:197.

Van, R.L.R. and D.A.K. Roncari. 1978. Cell Tissue Res., 195:317.

Vezinhet, A. and M. Prud'hon. 1975. Anim. Prod. 20:363.

Vezinhet, A., M. Prud'hon, and M. Benevent. 1974. Ann. Biol. Anim. Biochem. Biophys., 14:117. Wagnon, K.A. 1960. J.Range Mgmt, 13:89.

Walstra, P. 1974. Livestock Prod. Sci., 1:187.

Walton, P.E. and T.D. Etherton. 1986. J. Anim. Sci., 62:1584.

Wasserman, A.E. 1979. J.Food Sci., 44:6.

Wasserman, R.H. 1977. "Bones," in Duke's Physiology of Domestic Animals. 9th Editition, M.J.Svenson, ed., Ithaca: Comstock, pp. 413-432. Wensvoort, P. 1967. Path.Vet., 4:69.

Wensvoort, P. 1968. Path.Vet., 5:270.

Westmorland, N. 1971. Fed.Proc., 30:1001.

Whittem, J.H. 1957. J.Path.Bact., 73:375.

Wilson, L.L., H.B. Roth, J.H. Ziegler, and J.D. Sink. 1977. J. Anim. Sci., 44:932.

Wise, D.R. 1977. "The Growth of the Skeleton," in Growth and Poultry Meat Production, K.N.Boorman and B.J. Wilson, eds., Edinburgh: British Poultry Science Ltd., pp. 65-78. Wood, E.M. 1967. Anat. Rec., 157:437.

Wood, J.D., M.B. Enser, and D.J. Restall. 1978. Anim.Prod., 27:1.

Wood, J.D., M.B. Enser, and D.J. Restall. 1975. J.Agric.Sci.,Camb., 84:221.

Wood, J.D., N.G. Gregory, G.M. Hall, and D. Lister. 1977. Br.J.Nutr., 37:167.

Wright, J.T. and G.J. Hausman. 1990. J. Anim. Sci., 68:1170.

Wu, J.J., C.L. Kastner, M.C. Hunt, D.H. Kropf, and D.M. Allen. 1981. J. Anim. Sci., 53:1256.

Young, O. and T.J. Braggins. 1993. Meat Sci., 35:213.

Young, H.E., E.M. Ceballos, J.C. Smith, M.L. Mancini, R.P. Wright, B.L. Ragan, I. Bushell, and P.A. Lucas. 1993. In Vitro Cell. Dev. Biol. 29A:723. Young, O.A., B.W. Hogg, B.J. Mortimer, and J.E. Waller. 1993. N. Z. J. Agric. Res., 36:143.

Zophoniasson, P. 1929. Nord.JordbrForsk., 11:327.

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