Open access peer-reviewed chapter

Cellularity Description of Adipose Depots in Domesticated Animals

Written By

Olaia Urrutia, Leopoldo Alfonso and José A. Mendizabal

Submitted: 30 December 2017 Reviewed: 16 January 2018 Published: 23 February 2018

DOI: 10.5772/intechopen.74109

From the Edited Volume

Adipose Tissue

Edited by Leszek Szablewski

Chapter metrics overview

1,371 Chapter Downloads

View Full Metrics


Cellularity of adipose tissue in domesticated animals varies not only with species, sex, age and management conditions but also with depot. Differences in depots are important in animal production because of the economic and welfare implications and in humans in relation to obesity. The final amount of fat and its composition depends on the differentiation of mesenchymal multipotent precursor cells into mature adipocytes (adipogenesis) capable of fatty acid and triglyceride synthesis (lipogenesis), both processes being regulated by different key adipogenic and lipogenic genes, some of are well known and have been described. Histologically, differences can be classified as hyperplasia (an increase in adipocyte number) and hypertrophy (an increase in adipocyte size), processes that can produce adipocyte size distributions that are not necessarily Gaussian. A detailed description of the type of adipocyte size distribution can help distinguish the different adipocyte populations within depots and characterise each not only in terms of the size but also the number of the constituting cells. This description can help better understand the development and role of the different depots. It can also help when analysing causal relationships with adipogenic drivers and lipogenic enzymes involved in lipid metabolism.


  • adipocyte
  • adipogenesis
  • lipogenesis
  • hyperplasia
  • hypertrophy
  • size
  • number
  • distribution

1. Introduction: cellularity of adipose tissue

Adipose tissue is a specialised connective tissue mesenchymal in origin formed by an association of cells called adipocytes, namely, cells that specialise in accumulating lipids. Adipose tissue plays a fundamental role in maintaining the energy balance in animals, which entails storing energy in the form of fat (triglycerides) during periods of high energy intake and releasing energy through lipolysis during periods of caloric restriction [1]. However, certain discoveries made more than 20 years ago revealed that it is also a highly active endocrine tissue that secretes important substances, namely, adipokines and adipocytokines (leptin, adiponectin, tumour necrosis factor alpha or TNF-α, etc.), enabling it to communicate with different organs and demonstrating its involvement in such biological processes as neuroendocrine function and immune function [2, 3].

There are two main types of adipose tissue, white adipose tissue and brown adipose tissue, formed mainly of white and brown adipocytes, respectively. These tissues are differentiated not just only in terms of their colour but also in terms of their morphology, partitioning, genes and function [4]. The third adipocyte type known as “beige/brite” adipocytes growing in white adipose tissue in response to hormonal stimuli and cold has recently been described in rats and humans [5], as well as in cattle [6] and sheep [7].

Brown adipose tissue consists of small adipocytes (25–40 μm in diameter), which are multilocular, i.e., they contain several lipid droplets surrounded by large numbers of mitochondria; their main functions are thermogenesis, i.e., heat production, and thermal homeostasis [8]. This type of adipose tissue is typical in newborn animals in most species and in adult hibernating mammals [9, 10].

White adipose tissue is the predominant form in adult animals and is the main focus of the considerations set out here. White adipose tissue is composed mainly of adipocytes and a stromal vascular fraction comprising macrophages, capillary endothelial cells, undifferentiated preadipocytes, pluripotent stem cells and fibroblasts. This stroma is responsible for tissue homeostasis [11]. The adipocytes in white adipose tissue are ordinarily large (in cattle they can reach sizes of up to 180 μm in diameter) [12], and they have few mitochondria, their cytoplasm being almost entirely taken up by a single fat vacuole [13, 14]. In addition to its endocrine function, this fatty tissue subtype also plays a structural role, providing mechanical support and protection for certain parts of the body [3, 15, 16].


2. Adipose depots in domesticated animals

Adipose tissue is located in specific deposits, or depots, that are similar in mammals, though the size and composition of the depots vary with species, age, sex, diet, etc. [17]. The main depots are the visceral depots, comprising the kidney knob and channel fat (covering the kidneys and the pelvic cavity), mesenteric fat (surrounding the bowel) and omental fat (surrounding the pre-stomachs), the subcutaneous (SC) depot (covering the outside of the carcass), the intermuscular depot (located in between the muscles) and the intramuscular (IM) depot (infiltrated within the muscles) [18]. For example, Joy et al. [19] observed that omental fat accounted for 10%, kidney knob and channel fat 8.13%, mesenteric fat 8.01% and SC fat 25.3%, with intermuscular fat accounting for 42.6%, in lambs with a live weight of 23 kg fed a concentrate diet. Therefore, the importance of the IM depot is small compared with that of the other depots in quantitative terms.

The relevance of fatty tissue is the effect of the proportions and compositions of the different depots on production efficiency, nutritional value and meat and carcass quality. Fat is present in meat in the form of intermuscular fat, membrane phospholipids and IM and SC fat [20]. From a production standpoint, not all the depots have the same value, with the depositing of IM fat, or marbling, being desirable because of its beneficial effects on sensory attributes and hence its decisive role in determining meat quality [21]. In contrast, the accumulation of excess fat in certain depots has an adverse effect by increasing production costs while decreasing product quality [15, 22]. For instance, visceral fat is mostly discarded at slaughter, while an excess of SC fat lowers the commercial value of carcasses. Subcutaneous fat is also of concern to consumers, because excess SC fat and a high saturated fatty acid (SFA) content are viewed as detrimental to human health [15, 23].

Therefore, the current trend is towards producing lean meats with optimum fatness levels, in particular suitable amounts of IM fat. Additionally, a certain amount of covering, or subcutaneous, fat is necessary to reduce carcass moisture loss and keep the carcass from drying out while also guarding it against potential bacterial contamination [24].

Intramuscular fat composition is especially important because of its impact on meat quality from the standpoint of human health. IM fat is formed mainly by phospholipids, which have a high polyunsaturated fatty acid (PUFA) content, and by triglycerides, consisting mostly of SFAs and monounsaturated fatty acids (MUFAs) [25]. The phospholipid content in the muscle is relatively constant because of phospholipid’s role as structural constituents of the cells, so it is not directly related to fatness. What is more, it is not particularly influenced by species, breed, diet or age, though the n−6/n−3 PUFA ratio may experience some variation according to the diet [26]. Unlike phospholipids, triglycerides in IM tissue may vary considerably from 0.2 to 5%, depending on accumulated fat levels, muscle location and breed [27]. Triglycerides make up most of the lipids (around 90%) in SC adipose tissue, and the PUFA content, especially eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), is low because phospholipids make up just a small proportion [28, 29, 30].

As already indicated, PUFAs are selectively deposited among the phospholipids associated with muscle cell membrane, which may furnish opportunities for modifying muscle PUFA content in ways beneficial to human health without increasing the amount of IM fat [20, 31].


3. Adipogenesis and lipogenesis

Adipogenesis is the process of differentiation by which multipotent mesenchymal precursor cells turn into mature adipose cells [32, 33]. Mesenchymal stem cells are fibroblast cells mesodermal in the origin capable of acting as precursors not only for adipose tissue but also for muscle, bone, cartilage and tendon [34]. There are two stages in the process of adipocyte differentiation: determination and terminal differentiation (Figure 1).

Figure 1.

Schematic representation of the transition process from mesenchymal stem cell to mature adipocyte and transcriptional cascade during adipogenesis. DLK-1 = Delta-like 1 homolog; AP-1 = activator protein-1; GATA2 and 3 = GATA binding proteins 2 and 3; WNT = wingless-type MMTV integration site family members; Zfp423 = zinc finger protein 423; CEBP a, β and δ = CCAAT/enhancer-binding protein a, β and δ; PPARg = peroxisome proliferator-activated receptor gamma; RXRa = retinoic X receptor; LPL = lipoprotein lipase; FABP4 = fatty acid-binding protein; aP2 = adipocyte protein 2 and GLUT4 = glucose transporter type-4.

The first stage encompasses the mechanisms, whereby the pluripotent stem cell takes on the characteristics of the adipocyte lineage and involves transformation of the stem cell into a preadipocyte that is morphologically still the same as the precursor. During this transition, the cell loses its ability to turn into other types of cells [35]. Both positive (zinc finger protein 423 or Zfp423; activator protein-1 or AP-1) and negative (delta-like 1 homologue or DLK1, GATA-binding proteins 2 and 3 and wingless-type MMTV integration site family members or WNTs) regulatory factors are known to take part in the initial stage, which is still poorly understood [36].

In the second stage of terminal differentiation, the preadipocytes acquire the complement of proteins needed for lipid transport and synthesis, insulin sensitivity and the ability to secrete adipokines and turn into mature adipocytes [9, 37]. This is accompanied by the accumulation of fat inside the cell and a change in morphology into a more globular shape [36]. This second stage is viewed as a cascade of transcriptional events in which the first wave consists of induction of CCAAT/enhancer-binding protein (CEBP) β (CEBPβ) and δ (CEBPδ), which activate expression of the central adipogenesis factors peroxisome proliferator-activated receptor gamma (PPARγ) and CEBPα. These transcription factors stimulate expression of genes involved in lipogenesis, such as lipoprotein lipase (LPL), fatty acid-binding protein (FABP4) or adipocyte protein 2 (aP2) and glucose transporter type-4 (GLUT4) [38, 39].

A variety of extracellular factors are capable of acting on the regulation of the above-mentioned transcription factors and are able to determine whether preadipocytes start the process of differentiation or remain quiescent. The activating factors include insulin, glucocorticoids, mono- and polyunsaturated fatty acids (which appear to be PPARγ activators) and prostaglandins [40].

Lipogenesis comprises the processes of fatty acid synthesis and triglyceride formation and takes place primarily in the liver, the white adipose tissue, and in the case of lactating females, the mammary gland. The biochemical mechanisms occurring in these tissues are similar, but the role of each of these processes varies by animal species. Adipose tissue is the main location of fatty acid synthesis in ruminants, and indeed in sheep, this tissue is responsible for over 90% of fatty acid (FA) biosynthesis [41]. As in ruminants, adipose tissue is the main site of fatty acid synthesis in pigs [42]. By contrast, in poultry it is the liver that is most active in de novo lipogenesis, while in rats, mice and rabbits, activity levels are similar in both tissues.

The fatty acids thus formed may be used to make cell membrane-building phospholipids, they may be used as precursors for biologically active metabolites or they may be used in the synthesis of triglycerides to store metabolic energy. This last-mentioned process is carried out by the endoplasmic reticulum in adipocytes and involves esterification of activated fatty acids (acyl-CoA molecules) through the action of acyl-CoA synthetase (ACS) and glycerol-3-phosphate from glucose catabolism.

Fatty acids from de novo synthesis are, as mentioned above, a source of acyl-CoA, but fatty acids ingested in the diet imported by LPL or released by lipolysis may also be used. Glycerol-3-phosphate is obtained by the action of the G3PDH enzyme on dihydroxyacetone phosphate (DHAP), though a small quantity may be produced by the action of glycerol kinase on glycerol [43].


4. Adipocyte hyperplasia and hypertrophy

Adipose tissue plasticity is high, and growth is produced by a combination of two mechanisms: hyperplasia, or an increase in the number of cells, and hypertrophy, or an increase in cell volume. Hyperplasia is a proliferation of preadipocytes and their subsequent differentiation and occurs primarily in the animal foetus and postnatal period. Hypertrophy also takes place during this period, whereby adipocyte volume increases through the accumulation of lipids in the form of triglycerides, formed by the esterification of glycerol-3-phosphate, which is synthesised from glucose and fatty acids (from the diet or synthesised de novo) [4, 13, 44, 45].

After birth and the postnatal period, adipose tissue growth is mainly by hypertrophy of existing adipocytes and the activity of lipogenic enzymes [46]. Nevertheless, adipose tissue size in animals is not necessarily limited by the hyperplasia taking place at an early age, inasmuch as adipocytes can stimulate adipogenesis when a given percentage of adipocytes have reached their maximum volume, thereby inducing an increase in hyperplasia and/or promoting lipid accumulation by preadipocytes which had been quiescent until then [13, 43, 47].

Both hyperplasia and hypertrophy can be influenced by factors such as sex, breed, age, physical condition, diet type and amount, anatomical location of the adipose depots, etc. [48]. As a whole, fat deposition increases with animal weight and age, though development of the different fat depots is not uniform from either a quantitative or a temporal standpoint [49]. Generally speaking, the kidney knob and channel fat depot are the first to grow, followed by the intermuscular and SC depots and, lastly, the IM depot [13, 15]. In sheep, hyperplasia in the kidney knob and channel fat depot is complete approximately 60 days after birth, and further growth is ascribed to hypertrophy. Both processes contribute to growth of the intermuscular depot, with hyperplasia occurring until around 100 days after conception. In contrast, hyperplasia and hypertrophy together may contribute to SC depot growth until lambs turn 12 months of age [13, 44].

IM fat is a late-developing depot, and fat deposition in the IM depot would appear to depend mainly on hyperplasia [23, 50, 51]. In older animals the rate of fat accumulation outpaces muscle growth, in contrast to the situation at younger ages, and as a result, intramuscular fat deposition necessarily takes place later in life [23].

As already mentioned, adipose depots do not all grow at the same time or at the same rate, and adipocyte size therefore varies from one depot to another [22]. As a general rule, mean adipocyte size by depot in sheep [50, 52, 53] and cattle [54, 55, 56, 57] follows this order: omental > kidney knob and channel fat > SC > intermuscular > IM. Furthermore, lipogenic activity in the different fat depots also varies, and this would appear to be related to adipocyte size and thus for the most part follows a similar order [22].

Intramuscular adipose tissue makes up a single depot, because it can be distinguished from other fat depots by its location within the perimysial connective tissue next to the myofibrils, and in addition its metabolism displays certain differences as compared to the other depots. Rates of fatty acid synthesis by intramuscular adipocytes are some 5–10% of the rate observed in the SC depot, which means that the fatty acid synthesising ability of the IM depot is low, as is its ability to break down fatty acids [58]. Gondret et al. [59] carried out proteomic analysis and reported that various lipogenesis, lipolysis, fatty acid oxidation and basal metabolic rate indicators were lower in IM tissue than in SC tissue.

Furthermore, intramuscular adipocytes chiefly use glucose and/or lactate as a carbon source for lipogenesis, whereas SC adipocytes use acetate [60]. Consequently, as Hausman et al. noted [58], there is evidence that the IM and SC tissues are metabolically distinct, with these differences manifesting at the cellular level (preadipocytes and adipocytes), such that, for instance, the proliferation and differentiation potency of bovine intramuscular preadipocytes is lower than that of bovine subcutaneous preadipocytes [61, 62].


5. Adipocyte size distribution

Adipocyte size in domesticated animal populations has been estimated for decades [63]. At the present time, there are different methods of evaluating adipocyte size, by means of electronic measurement following fixation with osmium tetroxide, by means of histological analysis and by means of collagenase digestion. These last two methods are constrained as to the numbers of adipocytes that can be analysed per sample (around 250), but they are less expensive and require less expenditure on equipment [64]. The collagenase digestion method, in which collagenase is used to dissolve the connective tissue surrounding the adipocytes [65], is frequently used in animal studies [48, 66, 67, 68, 69]. Succinctly, samples taken from animals at slaughter are stored in Tyrode’s solution at 39°C, blood vessels and connective tissue are removed and the samples are washed in saline solution and digested using collagenase under incubation at 39°C for 1 and a half hours. The solution is then filtered through an 850 μm filter, microscope slides are prepared and the diameters of a sample of at least 200 of the adipocytes thus obtained are measured using image analysis.

Irrespective of the method employed, for many years now, the size distributions of adipocytes in different animal species have sometimes been observed to fit a normal distribution [48, 53, 70], but this is not always the case. Bimodal distributions have been described [55, 71, 72, 73], and this may, though not always [74], indicate the presence of a heterogeneous cell population [75]. The bimodal distribution of adipocyte size could thus be an indication that both hyperplasia and hypertrophy are taking place at the same time or that factors contributing to heterogeneity during adipocyte hypertrophy are at work. Growth in adipocyte size may increase local hypoxia, resulting in cell death as well as, after a certain critical volume has been reached, in the secretion of new adipocyte recruitment factors [76]. Another possible source of the bimodal distributions could be that the metabolic activity of adipocytes depends on individual cell surface size, since triglyceride exchange with the extracellular milieu occurs across the surface, and consequently differences in lipolytic and lipogenic fluxes could give rise to adipocyte populations that differ in size without hyperplasia necessarily taking place [77].

In any case, proper description of adipocyte size distributions is needed to be able to identify the factors underlying the differences in adipose tissue growth between groups of animals. If a distribution is bimodal, describing adipocyte size on the basis of the mean and the standard error of the mean is not meaningful, and using the mean to compare differences in experimental treatments could lead to erroneous conclusions. Compared with simply calculating mean cell size and the total number of adipocytes, studies that deal with adipocyte distributions are better at explaining the mechanisms involved in fat development [78]. For this reason, various approaches to evaluating adipocyte size distributions have been put forward, e.g., lognormal distribution fitting [72], normal distribution mixtures [79] and combinations of exponential and normal distributions [80]. In any case, before putting any of these approaches to use, it is advisable first to check whether distributions are bimodal [81], since even exploratory data analysis methods like histograms may be misleading [82].

Alfonso and Mendizabal [81] proposed using the bimodality coefficient (BC) [83] and the dip statistic [84] to test the unimodal distribution of the data without having to assume any distribution underlying the bimodal distribution, though other approaches have also been described [85]. If the data prove not to be unimodal, the bimodal adipocyte size distribution can be described by estimating the two distribution modes and the proportion of adipocytes located under the inflexion point (nadir) between the two, thus enabling one population of small adipocytes to be differentiated from a second population of large ones. These three parameter values can be related to the degree of fatness in the animals as illustrated in Figure 2. The relationship needs not be unique but may vary with the species, tissue, age, physical growth, reserve mobilisation, etc. according to the relative importance of hyperplasia and hypertrophy that may be taking place at the time.

Figure 2.

Relationship between the first mode, second mode and percentage small adipocytes and backfat thickness in the outer subcutaneous layer in brood sows (data after Abadía et al. [67]).

Figure 3 depicts what happens to these parameters with an increase in the number of small adipocytes (hyperplasia) or an increase in the size of large adipocytes (hypertrophy). This is a simulated example (as described by Alfonso [86]), which shows that the hypothesis of hyperplasia (an increase in the number of adipocytes) can be tested on the basis of the differences occurring in the percentage of small adipocytes and the hypothesis of hypertrophy (an increase in the diameter of large adipocytes) can be tested on the basis of the differences occurring in the second mode. The effect of both processes combined is also shown, since both changes in adipocyte number and changes in adipocyte size are known to take place simultaneously [87].

Figure 3.

Simulated adipocyte size distribution depicting changes in the bimodal distributions in conditions of hyperplasia and hypertrophy. Arrows indicate the changes observed.

Analysing the data in this way allows differences between groups of animals to be described even in the presence of bimodality. Thus, differences between tissues in cattle [68] and pigs [81, 85], between breeds [85] and between animals grouped by age [88] and sex [85] have been described. Describing these differences is an essential basis for subsequently studying cell multiplication and differentiation both at the genetic level and at the level of expression, the better to explain the mechanisms of adipose tissue growth and be able to act appropriately so as to modulate them with a view to improving animal production systems. These same working methods can be extrapolated to the study of adipose tissue in humans and may help explain the factors involved in obesity and associated health issues.


6. Conclusion

Adipose tissue plays a fundamental role in maintaining the energy balance in animals, storing energy in the form of triglycerides during periods of high energy intake and releasing energy through lipolysis during periods of caloric restriction. In domesticated animals these triglycerides are located in different depots which vary in size and composition with species, sex, age and diet. These same factors in animal production systems condition adipose tissue growth brought about hyperplasia, or the proliferation of adipocytes, and hypertrophy, or the increase in adipocyte size. Both processes are regulated by different factors that stimulate or inhibit the expression of adipogenic and lipogenic genes. A knowledge of these processes enhances our understanding of the genetic basis underlying the growth of fat depots in domesticated animals so as to be able to modify their quantity and composition. Thus, studying adipocyte size distributions helps us understand the different levels of involvement of, on the one hand, hypertrophy and hyperplasia and, on the other, of adipogenic and lipogenic genes and regulatory factors on adipose tissue growth in different groups of animals.



This research was supported by the Instituto Nacional de Investigación y Tecnología Agraria y Alimentaria [National Institute for Agricultural and Food Research and Technology] (INIA project RTA2013-00046-CO3-03).


  1. 1. Pond CM. The evolution of mammalian adipose tissues. In: Symonds M, editors. Adipose Tissue Biology. 2nd ed. Cham: Springer; 2017. p. 1-59. DOI:,031-5_1
  2. 2. Trayhurn P, Beattie JH. Physiological role of adipose tissue: white adipose tissue as an endocrine and secretory organ. Proceedings of Nutrition Society. 2001;60:329–339. DOI:
  3. 3. Kershaw EE, Flier JS. Adipose tissue as an endocrine organ. Journal of Clinical and Endocrinology Metabolism. 2004;89:2548-2556. DOI: 10.1210/jc.2004-0395
  4. 4. Bonnet M, Cassar-Malek I, Chilliard Y, Picard B. Ontogenesis of muscle and adipose tissues and their interactions in ruminants and other species. Animal. 2010;4:1093-1109. DOI: 10.1017/S1751731110000601
  5. 5. Harms M, Seale P. Brown and beige fat: Development, function and therapeutic potential. Natural Medicine. 2013;19:1252-1263. DOI:10.1038/nm.3361
  6. 6. Asano H, Yamada T, Hashimoto O, Umemoto T, Sato R, Ohwatari S, Kanamori Y, Terachi T, Funaba M, Matsui T. Diet-induced changes in Ucp1 expression in bovine adipose tissues. Generative and Comparative Endocrinology. 2013;184:87-92. DOI:10.1016/j.ygcen.2013.01.006
  7. 7. Pope M, Budge H, Symonds ME. The developmental transition of ovine adipose tissue through early life. Acta Physiology. 2014;210:20-30. DOI:10.1111/apha.12053
  8. 8. Rosen ED, Spiegelman BM. Adipocytes as regulators of energy balance and glucose homeostasis. Nature. 2006;444:847-853. DOI: 10.1038/nature05483
  9. 9. Rosen ED, MacDougald OA. Adipocyte differentiation from the inside out. Nature Reviews in Molecular Cellular Biology. 2006;7:885-896. DOI: 10.1038/nrm2066
  10. 10. Klingenspor M, Bast A, Bolze F, Li Y, Maurer S, Schweizer S, Willershäuser M, Fromme T. Brown adipose tissue. In: Symonds M, editor. Adipose Tissue Biology. 2nd ed. Cham: Springer; 2017. pp. 91-147. DOI:
  11. 11. Trujillo ME, Scherer PE. Adipose tissue-derived factors: Impact on health and disease. Endocrine Reviews. 2006;27:762-778. DOI:10.1210/er.2006-0033
  12. 12. Cianzio DS, Topel DG, Whiterhurst GB, Beitz DC, Self HL. Adipose tissue growth and cellularity: Changes in bovine adipocyte size and number. Journal of Animal Science. 1985;60:970-976. DOI:10.2527/jas1985.604970x
  13. 13. Lawrence TLJ, Fowler VR. Tissues: Basic structure and growth. In: Lawrence TLJ, Fowler VR, editors. Growth of Farm Animals. London, UK: CABI Publishing; 2002. pp. 21-83
  14. 14. Gesta S, Kahn CR. White adipose tissue. In: Symonds M, editor. Adipose Tissue Biology. 2nd ed. Cham: Springer; 2017. pp. 149-159. DOI:,031-5_5
  15. 15. Louveau I, Perruchot M-H, Bonnet M, Gondret F. Invited review: Pre- and postnatal adipose tissue development in farm animals: from stem cells to adipocyte physiology. Animal. 2016;10:1839-1847. DOI:10.1017/S1751731116000872
  16. 16. Vigouroux C, Caron-Debarle M, Le Dour C, Magre J, Capeau J. Molecular mechanisms of human lipodystrophies: from adipocyte lipid droplet to oxidative stress and lipotoxicity. International Journal of Biochemistry Cellular Biology. 2011;43:862-876. DOI:10.1016/j.biocel.2011.03.002
  17. 17. Kempster AJ. Fat partition and distribution in the carcasses of cattle, sheep and pigs: A review. Meat Science. 1981;5:83-98. DOI:10.1016/0309-1740(81)90007-3
  18. 18. Robelin J. Growth of adipose tissues in cattle; partitioning between depots, chemical composition and cellularity. A review. Livestock Production Science. 1986;14:349–364
  19. 19. Joy M, Ripoll G, Delfa R. Effects of feeding system on carcass and non-carcass composition of Churra Tensina light lambs. Small Ruminant Research. 2008;78:123-133. DOI:
  20. 20. Scollan ND, Hocquette JF, Nuernberg K, Dannenberger D, Richardson I, Moloney AP. Innovations in beef production systems that enhance the nutritional and health value of beef lipids and their relationship with meat quality. Meat Science. 2006;74:17-33. DOI: 10.1016/j.meatsci.2006.05.002
  21. 21. Savell JW, Branson RE, Cross HR, Stiffler DM, Wise JW, Griffin DB, Smith GC. National consumer retail beef study: Palatability evaluations of beef loin steaks that differed in marbling. Journal of Food Science. 1987;52:517-519. DOI:10.1111/j.1365-2621.1987.tb06664.x
  22. 22. Eguinoa P, Brocklehurst S, Arana A, Mendizabal JA, Vernon RG, Purroy A. Lipogenic enzyme activities in different adipose depots of Pirenaican and Holstein bulls and heifers taking into account adipocyte size. Journal of Animal Science. 2003;71(81):432-440. DOI: 10.2527/2003.812432x
  23. 23. Hocquette JF, Gondret F, Baeza E, Medale F, Jurie C, Pethick DW. Intramuscular fat content in meat-producing animals: development, genetic and nutritional control, and identification of putative markers. Animal. 2010;4:303-319. DOI:10.1017/S1751731109991091
  24. 24. Wood JD. Fat deposition and the quality of fat tissue in meat animals. In: Wisseman J, ed. Fats in Animal Nutrition. London: Butterworths; 1984. p. 407-435
  25. 25. Wood JD, Richardson RI, Nute GR, Fisher A V Campo MM, Kasapidou E, Sheard PR, Enser M. Effects of fatty acids on meat quality: A review. Meat Science. 2004;66:21-32. DOI: 10.1016/S0309-1740(03)00022-6
  26. 26. Enser M, Hallett KG, Hewett B, Fursey GAJ, Wood JD, Harrington G, Hewett B, Fursey GAJ, Wood JD, Harrington G. Fatty acid content and composition of UK beef and lamb muscle in relation to production system and implications for human nutrition. Meat Science. 1998;49:329-341. DOI: 10.1016/S0309-1740(97)00144-7
  27. 27. Raes K, De Smet S, Demeyer DI. Effect of dietary fatty acids on incorporation of long chain polyunsaturated fatty acids and conjugated linoleic acid in lamb, beef and pork meat: A review. Animal Feed Science and Technology. 2004;113:199-221. DOI:
  28. 28. Ashes JR, Siebert BD, Gulati SK, Cuthbertson AZ, Scott TW. Incorporation of n − 3 fatty acids of fish oil into tissue and serum lipids of ruminants. Lipids. 1992;27:629–631
  29. 29. Duckett SK, Wagner DG, Yates LD, Dolezal HG, May SG. Effects of time on feed on beef nutrient composition. Journal of Animal Science. 1993;71(8):2079-2088. DOI: 10.2527/1993.7182079x
  30. 30. Howes NL, Bekhit AEDA, Burritt DJ, Campbell AW. Opportunities and implications of pasture-based lamb fattening to enhance the long-chain fatty acid composition in meat. Comprehensive Reviews in Food Science and Food Safety. 2015;14:22-36. DOI: 10.1111/1541-4337.12118
  31. 31. Shingfield KJ, Bonnet M, Scollan ND. Recent developments in altering the fatty acid composition of ruminant-derived foods. Animal. 2013;7:132-162. DOI: 10.1017/S1751731112001681
  32. 32. Hausman DB, DiGirolamo M, Bartness TJ, Hausman GJ, Martin RJ. The biology of white adipocyte proliferation. Obesity Reviews. 2001;2(4):239-254. DOI: 10.1046/j.1467-789X.2001.00042.x
  33. 33. Kokta TA, Dodson MV, Gertler A, Hill RA. Intercellular signalling between adipose tissue and muscle tissue. Domestic Animal Endocrinology. 2004;27(4):303-331. DOI: 10.1016/j.domaniend.2004.05.004
  34. 34. Chen Y, Shao J-Z, Xiang L-X, Dong X-J, Zhang G-R. Mesenchymal stem cells: a promising candidate in regenerative medicine. International Journal of Biochemical and Cellular Biology. 2008;40:815-820. DOI: 10.1016/j.biocel.2008.01.007
  35. 35. Christodoulides C, Lagathu C, Sethi JK, Vidal-Puig A. Adipogenesis and WNT signalling. Trends in Endocrinology Metabolism. 2009;20(1):16-24. DOI: 10.1016/j.tem.2008.09.002
  36. 36. Lefterova MI, Zhang Y, Steger DJ, Schupp M, Schug J, Cristancho A, Feng D, Zhuo D, Stoeckert CJ, Liu XS, Lazar MA. PPAR gamma and C/EBP factors orchestrate adipocyte biology via adjacent binding on a genome-wide scale. Genes Development. 2008;22:2941-2952. DOI:10.1101/gad.1709008
  37. 37. Lefterova MI, Lazar MA. New developments in adipogenesis. Trends in Endocrinology and Metabolism. 2009;20:107-114 DOI: 10.1016/j.tem.2008.11.005
  38. 38. Wu Z, Xie Y, Bucher NL, Farmer SR. Conditional ectopic expression of C/EBP beta in NIH-3 T3 cells induces PPAR gamma and stimulates adipogenesis. Genes Development. 1995;9:2350-2363. DOI:10.1101/gad.9.19.2350
  39. 39. Wu Z, Bucher NL, Farmer SR. Induction of peroxisome proliferator-activated receptor gamma during the conversion of 3 T3 fibroblasts into adipocytes is mediated by C/EBPbeta, C/EBPdelta, and glucocorticoids. Molecular and Cellular Biology. 1996;16(8):4128–4136
  40. 40. Gregoire FM, Smas CM, Sul HS. Understanding adipocyte differentiation. Physiology Reviews. 1998;78:783-809. DOI: 10.1074/jbc.272.8.5128
  41. 41. Ingle DL, Bauman DE, Garrigus US. Lipogenesis in the ruminant: in vivo site of fatty acid synthesis in sheep. Journal of Nutrition. 1972;102:617–626
  42. 42. Vernon RG. Lipid metabolism in the adipose tissue of ruminant animals. Progress in Lipid Research. 1980;19:23–106
  43. 43. Mersmann HJ, Smith SB. Development of white adipose tissue lipid metabolism. In: Burrin DG, Mersmann HJ, editors. Biology of Metabolism in Growing Animals. Elsevier Limited; 2005. p. 275-302. DOI: 10.1016/S0376-7361(09)70018-4
  44. 44. Mendizabal JA, Theriez M, Bas P, Normand J, Aurousseau B, Purroy A. Fat firmness of subcutaneous adipose tissue in intensively reared lambs. Small Ruminant Research. 2004;53:173-180. DOI:
  45. 45. Vernon RG, Denis RGP, Sorensen A. Signals of adiposity. Domestic Animal Endocrinology. 2001;21:197-214. DOI: 10.1016/S0739-7240(01)00121-7
  46. 46. Robelin J. Cellularity of bovine adipose tissues: developmental changes from 15 to 65 percent mature weight. Journal of Lipid Research. 1981;22:452-457
  47. 47. Dodson MV, Hausman GJ, Guan L, Du M, Rasmussen TP. Lipid metabolism, adipocyte depot physiology and utilisation of meat animals as experimental models for metabolic research. International Journal of Biological Science. 2010;6:691-699. DOI: 10.7150/ijbs.6.691
  48. 48. Mendizabal JA, Soret B, Purroy A, Arana A, Horcada A. Influence of sex on cellularity and lipogenic enzymes of Spanish lamb breeds (Lacha and Rasa Aragonesa). Animal Science. 1997;64:283-289. DOI: 10.1017/S135772980001585X
  49. 49. Arana A, Mendizabal JA, Alzon M, Eguinoa P, Beriain MJ, Purroy A. Effect of feeding lambs oleic acid calcium soaps on growth, adipose tissue development and composition. Small Ruminant Research. 2006;63:75-83. DOI:
  50. 50. Baik M, Jeong JY, Vu T-TT, Piao MY, Kang HJ. Effects of castration on the adiposity and expression of lipid metabolism genes in various fat depots of Korean cattle. Livestock Science. 2014;168:168-176. DOI:
  51. 51. Yang XJ, Albrecht E, Ender K, Zhao RQ, Wegner J. Computer image analysis of intramuscular adipocytes and marbling in the longissimus muscle of cattle. Journal of Animal Science. 2006;84:3251-3258. DOI: 10.2527/jas.2006-187
  52. 52. Arana A, Mendizabal JA, Alzon M, Soret B, Purroy A. The effect of vitamin A supplementation on postnatal adipose tissue development of lambs. Journal of Animal Science. 2008;86:3393-3400. DOI: 10.2527/jas.2008-0889
  53. 53. Soret B, Mendizabal JA, Arana A, Purroy A, Eguinoa P. Breed effects on cellularity and lipogenic enzymes in growing Spanish lambs. Small Ruminant Research. 1998;29:103-112. DOI:
  54. 54. Allen CE. Cellularity of adipose tissue in meat animals. Federation Proceedings. 1976;35:2302–2307
  55. 55. Hood RL, Allen CE. Cellularity of bovine adipose tissue. Journal of Lipid Research 1973;14:605–610
  56. 56. Mendizabal JA, Alberti P, Eguinoa P, Arana A, Soret B, Purroy A. Adipocyte size and lipogenic activities in different adipose tissue in steers of local Spanish breeds. Animal Science. 1999;69:115-121. DOI:
  57. 57. Schiavetta AM, Miller MF, Lunt DK, Davis SK, Smith SB. Adipose tissue cellularity and muscle growth in young steers fed the beta-adrenergic agonist clenbuterol for 50 days and after 78 days of withdrawal. Journal of Animal Science. 1990;68:3614-3623. DOI: 10.2527/1990.68113614x
  58. 58. Hausman GJ, Dodson MV, Ajuwon K, Azain M, Barnes KM, Guan LL, Jiang Z, Poulos SP, Sainz RD, Smith S, Spurlock M, Novakofski J, Fernyhough ME, Bergen WG. Board-invited review: the biology and regulation of preadipocytes and adipocytes in meat animals. Journal of Animal Science. 2009;87:1218-1246. DOI: 10.2527/jas.2008-1427
  59. 59. Gondret F, Guitton N, Guillerm-Regost C, Louveau I. Regional differences in porcine adipocytes isolated from skeletal muscle and adipose tissues as identified by a proteomic approach. Journal of Animal Science. 2008;86:2115-2125. DOI: 10.2527/jas.2007-0750
  60. 60. Smith SB, Kawachi H, Choi CB, Choi CW, Wu G, Sawyer JE. Cellular regulation of bovine intramuscular adipose tissue development and composition. Journal of Animal Science. 2009;87:72-82. DOI: 10.2527/jas.2008-1340
  61. 61. Grant AC, Doumit ME, Tempelman RJ, Buskirk DD. Differentiation of bovine intramuscular and subcutaneous stromal-vascular cells exposed to dexamethasone and troglitazone. American Society of Animal Science. 2015;86:2531-2538. DOI: 10.2527/jas.2008-0860
  62. 62. Wan R, Du J, Ren L, Meng Q. Selective adipogenic effects of propionate on bovine intramuscular and subcutaneous preadipocytes. Meat Science. 2009;82:372-378. DOI: 10.1016/j.meatsci.2009.02.008
  63. 63. Hausman GJ. The comparative anatomy of adipose tissue. In: Cryer A, Van RLR, editors. New Perspectives in Adipose Tissue: Structure, Function and Development. London, UK: Butterworths; 1985. pp. 1-21
  64. 64. Laforest S, Michaud A, Paris G, Pelletier M, Vidal H, Gélöen A, Tchernof, A. Comparative Analysis of three human adipocyte size measurement methods and their relevance for cardiometabolic risk. Obesity. 2017;25:122-131. DOI: 10.1002/oby.21697
  65. 65. Rodbell M. Metabolism of isolated fat cells. Journal of Biological Chemistry. 1964;239:375-380. DOI: 10.1002/cphy.cp050147
  66. 66. Alfonso L, Mourot J, Insausti K, Mendizabal JA, Arana A. Comparative description of growth, fat deposition, carcass and meat quality characteristics of Basque and Large White pigs. Animal Research. 2005;54:33-42. DOI: 10.1051/animres:2.005.001
  67. 67. Abadía S, Mendizabal J, Alfonso L. Relationship between depth of individual backfat layers and size of their constituent adipocytes in sows. Acta Agriculturae Scandinavica, Section A — Animal Science. 2008;58:161–163. DOI:
  68. 68. Martínez del Pino L, Arana A, Alfonso L, Mendizabal JA, Soret B. Adiposity and adipogenic gene expression in four different muscles in beef cattle. PLoS ONE. 2017;12:e0179604. DOI:
  69. 69. Urrutia O, Mendizabal JA, Insausti K, Soret B, Purroy A, Arana A. Effects of addition of linseed and marine algae to the diet on adipose tissue development, fatty acid profile, lipogenic gene expression, and meat quality in lambs. PLoS ONE. 2016;11(6):e0156765. DOI: 10.1371/journal.pone.0156765
  70. 70. Arana A, Soret B, Mendizabal JA, Corroza M, Eguinoa P, Purroy A. Changes in adipose tissue accumulation in “Rasa Aragonesa” breed lambs during growth and fattening. Animal Science. 1998;66:409-413. DOI:
  71. 71. Mersmann HJ, Goodman JR, Brown LJ. Development of swine adipose tissue: morphology and chemical composition. Journal of Lipid Research. 1975;16(4):269-279
  72. 72. Whitehurst GB, Beitz DC, Cianzio D, Topel DG, Johnson DC. Examination of a lognormal distribution equation for describing distributions of diameters of bovine adipocytes. Journal of Animal Science. 1981;53:1236-1245. DOI: 10.2527/jas1981.5351236x
  73. 73. DeMartinis FD, Francendese A. Very small fat cell populations: mammalian occurrence and effect of age. Journal of Lipid Research. 1982;23(8):1107-1120
  74. 74. Baker GA. Transformations of bimodal distributions. Annals of Mathematical Statistics. 1930;1(4):334-344
  75. 75. Knapp TR. Bimodality revisited. Journal of Modern Applied Statistics Methods. 2007;6(1):article 3. DOI: 10.22237/jmasm/1177992120
  76. 76. Sun K, Kusminski CM, Scherer PE. Adipose tissue remodelling and obesity. Journal of Clinical Investigation. 2011;121:2094-2101. DOI: 10.1172/JCI45887
  77. 77. Soula HA, Julienne H, Soulage CO, Géloën A. Modelling adipocytes size distribution. Journal of Theoretical Biology. 2013;332:89-95. DOI: 10.1016/j.jtbi.2013.04.025
  78. 78. Jo J, Shreif Z, Periwal V. Quantitative dynamics of adipose cells. Adipocyte. 2012;1(2):80-88. DOI: 10.4161/adip.19705
  79. 79. Cruz GD, Strathe AB, Rossow HA, Fadel JG. Characterising bovine adipocyte distribution and its relationship with carcass and meat characteristics using a finite mixture model. Journal of Animal Science. 2012;90:2995-3002. DOI: 10.2527/jas.2011-4838
  80. 80. McLaughlin T, Sherman A, Tsao P, Gonzalez O, Yee G, Lamendola C, Reaven GM, Cushman SW. Enhanced proportion of small adipose cells in insulin resistant vs. insulin-sensitive obese individuals implicates impaired adipogenesis. Diabetologia. 2007;50:1707-1715. DOI: 10.1007/s00125-007-0708-y
  81. 81. Alfonso L, Mendizabal JA. Caracterización de la distribución del tamaño de los adipocitos para el estudio del tejido adiposo en producción animal. ITEA. 2016;112(2):147-161. DOI:
  82. 82. Fisher NI, Mammen E, Marron JS. Testing for multimodality. Computational Statistics and Data Analysis. 1994;18:499–512. DOI:
  83. 83. SAS Institute Inc. SAS/STAT User’s Guide Version 6. Fourth Edition, Vol. 1, Cary, NC; 1989. 943 p
  84. 84. Hartigan JA, Hartigan PM. The dip test of unimodality. The Annals of Statistics. 1985;13(1):70-84
  85. 85. Testroet ED, Sherman P, Yoder C, Testroet A, Reynolds C, O'Neil M, Lei SM, Beitz DC, Baas TJ. A novel and robust method for testing bimodality and characterising porcine adipocytes of adipose tissue of 5 purebred lines of pig. Adipocyte. 2017;6:102-111. DOI: 10.1080/21623945.2017.1304870
  86. 86. Alfonso L. Correction for adipocyte size and number in the analysis of differences in gene expression in fat tissues. Journal of Anima Breeding and Genetics. 2017;134:493-504. DOI: 10.1111/jbg.12296
  87. 87. Schoonmaker JP, Fluharty FL, Effect LSC. of source and amount of energy and rate of growth in the growing phase on adipocyte cellularity and lipogenic enzyme activity in the intramuscular and subcutaneous fat depots of Holstein steers. Journal of Animal Science. 2004;82(1):137-148
  88. 88. Soret B, Mendizabal JA, Arana A, Alfonso L. Expression of genes involved in adipogenesis and lipid metabolism in subcutaneous adipose tissue and longissimus muscle in low-marbled Pirenaica beef cattle. Animal. 2016;10:2018-2026. DOI: 10.1017/S175173111600118X

Written By

Olaia Urrutia, Leopoldo Alfonso and José A. Mendizabal

Submitted: 30 December 2017 Reviewed: 16 January 2018 Published: 23 February 2018