We train them. But how much do we know about them. And how do we know what type of muscle fibers we have in the muscle we are training; by knowing what muscle fibers a specific muscle has, can take the guess work out of what type of training that specific muscle requires.
Our muscle tissue consists of fibers (cells) that are highly specialized for the active generation of force for our muscle contraction. Muscle tissue provides motion, maintenance of our posture, and heat production. On the basis of certain structural and functional characteristics, the muscle tissue that our body has is classified into three types: cardiac, smooth and skeletal.
Cardiac muscle tissue forms the bulk of the wall of the heart. Like skeletal muscle tissue, it is striated. Unlike skeletal muscle tissue its contraction is usually not under conscious control and is classed as involuntary.
Smooth muscle tissue is located in the walls of hollow structures such as blood vessels, the stomach, intestines, and the bladder. Smooth muscle fibers are usually involuntary, and they are non-striated (smooth). Smooth muscle tissue, like skeletal and cardiac muscle tissue, can undergo hypertrophy (growth).
Skeletal muscle tissue is attached to our bones. It is striated; that is, the fibers (cells) contain alternating light and dark bands (striations) that are perpendicular to the long axes of the fibers. Skeletal muscle tissue can be made to contract or relax by conscious control (voluntary).
All skeletal muscle fibers are not alike in structure or function. For example, skeletal muscle fibers vary in colour depending on their content of Myoglobin (Myoglobin is found in muscle tissue, where it binds oxygen, helping to provide extra oxygen to release energy to power muscular contractions.) Skeletal muscle fibers contract with different velocities, depending on their ability to split Adenosine Triphosphate (ATP). Faster contracting fibers have greater ability to split ATP. In addition, skeletal muscle fibers vary with respect to the metabolic processes they use to generate ATP. They also differ in terms of the onset of fatigue. On the basis of various structural and functional characteristics, skeletal muscle fibers are classified into three types: Type I fibers, Type II B fibers and type II A fibers.
Type I Fibers
These fibers, also called slow twitch or slow oxidative fibers, contain large amounts of Myoglobin, many mitochondria and many blood capillaries. Type I fibers are red, split ATP at a slow rate, have a slow contraction velocity, very resistant to fatigue and have a high capacity to generate ATP by oxidative metabolic processes. Such fibers are found in large numbers in the postural muscles of the neck.
Type II A Fibers
These fibers, also called fast twitch or fast oxidative fibers, contain very large amounts of Myoglobin, very many mitochondria and very many blood capillaries. Type II A fibers are red, have a very high capacity for generating ATP by oxidative metabolic processes, split ATP at a very rapid rate, have a fast contraction velocity and are resistant to fatigue. Such fibers are infrequently found in humans.
Type II B Fibers
These fibers, also called fast twitch or fast glycolytic fibers, contain a low content of Myoglobin, relatively few mitochondria, relatively few blood capillaries and large amounts glycogen. Type II B fibers are white, geared to generate ATP by anaerobic metabolic processes, not able to supply skeletal muscle fibers continuously with sufficient ATP, fatigue easily, split ATP at a fast rate and have a fast contraction velocity. Such fibers are found in large numbers in the muscles of the arms.
Muscle Fiber Diagram
The diagram above shows the structure of the muscles. The inset picture (brown colored) shows you the fast and slow twitch fibers. Lighter = fast twitch / darker = slow twitch.
Characteristics of Different Muscle Fibers
|Fibre Type||Type I fibers||Type II A fibers||Type II B Type fibers|
|Contraction time||Slow||Fast||Very Fast|
|Size of motor neuron||Small||Large||Very Large|
|Resistance to fatigue||High||Intermediate||Low|
|Activity Used for||Aerobic||Long term anaerobic||Short term anaerobic|
|Force production||Low||High||Very High|
|Major storage fuel||Triglycerides CP||Glycogen CP||Glycogen|
Types of Muscle Fibers
Most skeletal muscles of the body are a mixture of all three types of skeletal muscle fibers, but their proportion varies depending on the usual action of the muscle. For example, postural muscles of the neck, back, and leg have a higher proportion of type I fibers. Muscles of the shoulders and arms are not constantly active but are used intermittently, usually for short periods of time, to produce large amounts of tension such as in lifting and throwing. These muscles have a higher proportion of type I and type II B fibers.
Even though most skeletal muscle is a mixture of all three types of skeletal, all the skeletal muscle fibers of any one motor unit are all the same. In addition, the different skeletal muscle fibers in a muscle may be used in various ways, depending on need. For example, if only a weak contraction is needed to perform a task, only type I fibers are activated by their motor units. If a stronger contraction is needed, the motor units of type II A fibers are activated. If a maximal contraction is required, motor units of type II B fibers are activated as well. Activation of the various motor units is determined in the brain and spinal cord. Although the number of the different skeletal muscle fibers does not change, the characteristics of those present can be altered.
The fast muscle (what the researchers call type IIa) moves 5 times faster than the slow muscle, and the super-fast (called type IIb) moves 10 times faster than the slow muscle fiber.
The average person has approximately 60% fast muscle fiber and 40% slow-twitch fiber (type I). There can be swings in fiber composition, but essentially, we all have three types of muscle fiber that need to trained.
Fiber Type Modification
Various types of exercises can bring about changes in the fibers in a skeletal muscle. Endurance type exercises, such as running or swimming, cause a gradual transformation of type II B fibers into type II A fibers. The transformed muscle fibers show a slight increase in diameter, mitochondria, blood capillaries, and strength. Endurance exercises result in cardiovascular and respiratory changes that cause skeletal muscles to receive better supplies of oxygen and carbohydrates but do not contribute to muscle mass. On the other hand, exercises that require great strength for short periods of time, such as weight lifting, produce an increase in the size and strength of type II B fibers. The increase in size is due to increased synthesis of thin and thick myofilaments. The overall result is that the person develops large muscles.
You can develop your fast-twitch muscle fiber by conducting plyometrics or complex training (combination of plyometrics and weights.) to build the fast muscle (IIa), and performing weight/strength training to build the super-fast (IIb) to the point where you can release exercise-induced growth hormone.
How to find your muscle fiber composition.
The objective of the muscle fiber test is to determine the fiber composition of the muscles being used for a particular exercise. Two test protocols are described: The Dr F. Hatield muscle fiber test and the Charles Poliquin muscle fiber test.
To undertake this test you will require:
- Weight training facilities
- An assistant/spotter
- Selection of exercises
How to conduct the Dr F. Hatfield muscle fiber test:
- Determine your one repetition maximum (1RM) on an exercise
- Rest for 15 minutes
- Perform as many repetitions as possible with 80% of your 1RM
- Less than 7 repetitions – Your fast twitch (FT) dominant
- 7 or 8 repetitions – You have a mixed fiber type
- More than 8 repetitions – You are slow twitch (ST) dominant
If you are FT dominant, then you should use heavier loads and lower repetitions predominantly in your training. ST dominant individuals, on the other hand, will respond better to lighter loads and higher repetitions
How to conduct the Charles Poliquin muscle fiber test:
- Determine your one repetition maximum (1RM) on an exercise
- Rest for 15 minutes
- Perform as many repetitions as possible with 85% of your 1RM
- Less than 5 repetitions – you are fast twitch (FT) dominant
- 5 repetitions – you have mixed fiber type
- More than 5 repetitions – you are slow twitch (ST) dominant
If you are FT dominant, then you should use heavier loads and lower repetitions predominantly in your training. ST dominant individuals, on the other hand, will respond better to lighter loads and higher repetitions.
Tbx15 is highly and specifically expressed in glycolytic myofibers
Tbx15 is a mesodermal gene involved in normal skeletal and muscle development17. In Tbx15LacZ/+ mice, in which the LacZ gene replaced exon 3 of the Tbx15 gene creating a fusion protein allowing tracking of Tbx15 expression17, there was robust X-gal staining in the skeletal muscle with no staining in the control (Fig. 1a). In C2C12 myoblasts, expression of Tbx15 messenger RNA (mRNA) increased ∼12-fold during differentiation, and this was confirmed by western blot analysis (Fig. 1b). Northern blot and quantitative PCR (qPCR) analysis of mouse tissues revealed that Tbx15 expression was at least eight times higher in muscle than adipose tissue, skin and pancreas, with much lower levels in other tissues (Fig. 1c; Supplementary Fig. 1a).
Assessment of individual muscles showed that Tbx15 mRNA was over two-fold higher in glycolytic muscles, such as the extensor digitorum longus (EDL), gastrocnemius and tibialis anterior, than in the oxidative muscle, such as soleus; and this was confirmed by western blot analysis (Fig. 1d). Fluorescent in situ hybridization on quadriceps muscle of wild-type mice demonstrated that Tbx15 message was not uniformly expressed, but in a mosaic pattern of fibre types. Similarly, immunofluorescence for Tbx15 in tibialis anterior muscle, which contains a mixture of oxidative and glycolytic fibres, confirmed that Tbx15 was expressed in a fibre-type-specific manner (Fig. 1e).
To determine which specific fibres exhibited high Tbx15 expression, we stained individual muscles from Tbx15LacZ mice. As shown in Fig. 1f, Tbx15-related X-gal staining was observed only in the glycolytic EDL muscle and not in the oxidative soleus muscle. When sections from tibialis anterior muscle were double stained using antibodies to Tbx15 and myosin IIa, it was clear that Tbx15 was virtually absent from type IIa, fast-twitch, oxidative fibres (Supplementary Fig. 1c). Furthermore, when serial sections were stained for succinate dehydrogenase (SDH), a marker of mitochondrial activity, <20% of the oxidative fibres, but over 80% of the glycolytic muscle fibres, scored positive for Tbx15 expression (Fig. 1g,h). Taken together, these data indicate that Tbx15 is expressed almost exclusively fast-twitch, glycolytic muscle. Interestingly, immunofluorescence also showed that Tbx15 was expressed in both the cytoplasm and nuclei of muscle fibres. These findings were confirmed by cell fractionation studies of skeletal muscle, which showed Tbx15 was present in the cytoplasm, microsomal fraction, and the nucleus (Supplementary Fig. 1b). Antibody specificity was verified by the absence of positive signal in both western blot analysis and immunofluorescence of Tbx15 in Tbx15−/− muscle (Fig. 2a; Supplementary Fig. 1d).
Endurance exercise in both humans and mice has been shown to lead to an increase in oxidative and decrease in glycolytic fibre density22. qPCR analysis revealed that 3 weeks of voluntary wheel cage running reduced mRNA Tbx15 levels by 15% in 6-week-old wild-type (WT) males. This is consistent with a reduction in glycolytic fibres, as myosin IIb mRNA levels were reduced 19%. Other markers of oxidative metabolism, including Pgc-1α, SDH, as well as markers of oxidative fibres, myosin I and myosin IIa, were all increased in exercised muscle (Supplementary Fig. 1e).
Ablation of Tbx15 leads to a decrease in glycolytic myofibers
To define the role of Tbx15 in skeletal muscle, we studied homozygous (Tbx15−/−) and heterozygous Tbx15 knockout (Tbx15+/−) mice. As previously reported, homozygous Tbx15 knockout animals had shortened limbs and other skeletal malformations17, whereas overt developmental abnormalities were noted in the heterozygous knockout animals. qPCR and western blot analysis of quadriceps muscle at 6 weeks of age shows that Tbx15 expression was reduced by about ∼40% in the heterozygous knockout animals and completely abrogated in the homozygous knockout (Fig. 2a). Despite the lack of gross abnormalities, heterozygous ablation of Tbx15 led to a significant ∼10% reduction in muscle mass (even when normalized to body weight) of both the EDL and tibialis anterior muscles, which are comprised mainly of glycolytic fibres. By contrast, there was no change in the weight of the more oxidative soleus muscle. The reduction in muscle mass was even more marked in Tbx15−/− mice with a 20–25% reduction in weight of the EDL and tibialis anterior muscles, again with no changes in the soleus muscle weight (Fig. 2b).
On histological examination of tibialis anterior from 6-week-old animals, there was a moderate increase in the relative density of oxidative, SDH positive fibres in the heterozygous knockout mice and a marked increase in the homozygous knockout compared with controls, in both cases with a corresponding decrease in glycolytic, SDH negative fibres (Fig. 2c). Quantitation of fibres from SDH-stained quadriceps muscle revealed a reduction in the total number of fibres of 11% in heterozygous and 43% in homozygous knockout mice. This reduction was due to a specific and gene dose-dependent reduction in the number of glycolytic muscle fibres of 33% in heterozygous and 72% in homozygous knockout mice (Fig. 2d). On the other hand, the number of oxidative fibres was significantly increased by 28% in heterozygous and was also increased by 12% in the muscle of homozygous knockout mice, although the latter did not quite reach statistical significance (analysis of variance; P=0.1).
Muscle fibre-type assessment by immunofluorescent staining for myosin heavy chain isoforms revealed that ablation of Tbx15 led to the appearance of considerable numbers of type I fibres in Tbx15+/− and Tbx15−/− EDL compared with WT mice, which have virtually no type I fibres in EDL muscle (Fig. 2c). Furthermore, these EDL muscles had a 2–3.5-fold increase in type IIa fibre density compared with controls. This increase in oxidative fibre density is due to a transformation of fibres into oxidative type I and type IIa fibres, as well as a reduction in glycolytic IIb fibres by 12 and 44% in the Tbx15+/− and Tbx15−/− EDL, respectively (Fig. 2e). Interestingly, despite the reduction in muscle weight being caused by a reduction in the number of glycolytic fibres in Tbx15+/− and Tbx15−/− mice, the cross-sectional area (CSA) of the remaining glycolytic fibres was significantly increased by 13 and 46% in tibialis anterior muscles of Tbx15+/− and Tbx15−/− animals, respectively. CSA of oxidative fibres was also increased by 18 and 57% in tibialis anterior muscles of Tbx15+/− and Tbx15−/− animals, respectively (Supplementary Fig. 2a). These changes in fibre size were associated with a change and broadening of distribution, as reflected in the s.d. of fibre size within each animal. This is true for both the glycolytic and oxidative fibres of Tbx15+/− and Tbx15−/− muscles, respectively (Supplementary Fig. 2b–c).
Ablation of Tbx15 leads to a decreased rate of muscle contraction
Since muscle fibre-type controls contraction and relaxation rates of skeletal muscle, with oxidative fibres characterized as slow-twitch and glycolytic fibres as fast twitch, we analysed the impact of loss of Tbx15 on muscle contractile properties. Both twitch and tetanic contraction were tested ex vivo using small fibre bundles of the EDL, normally a fast-twitch muscle. Recordings of single twitches were analysed for peak force, time to peak (TTP), and 50 and 75% relaxation times (t50 and t75%; Fig. 2f; Supplementary Fig. 2d). During single-twitch contraction, peak force was not altered, however. the rate of contraction, as measured by time to peak, was prolonged by ∼30% in Tbx15−/− muscles characteristic of slow, oxidative fibres. Likewise, muscle relaxation, as indicated by increased t50 and t75% values, was retarded in Tbx15−/− mice by ∼35%. After tetanic stimulation, muscle fibres of Tbx15−/− mice also demonstrated a ∼30% slower relaxation rate, as indicated by increase t50 and t75% values, compared with WT, again with no change in peak force (Supplementary Fig. 2d).
Ablation of Tbx15 does not affect skeletal muscle performance
To test whether the ablation of Tbx15 led to changes in muscle performance, we performed several tests of muscle performance both ex vivo and in vivo. In agreement with the results found in Tbx15−/− mice, no differences in maximal twitch force or tetanic force were observed in EDL muscle from Tbx15+/− mice compared with WT controls (Supplementary Fig. 2e–f). There was also no difference in fatigue resistance, as assessed by reduction in maximal tetanic force after repeatedly stimulated (Supplementary Fig. 2g).
In a treadmill exercise paradigm on 8-week-old male mice, there was no significant difference in the running time (13.6±1.5 min for WT versus 13±1.1 min for Tbx15+/−, n=6–8) or the total distance run (126.5±21.5 m versus 122.5±10.7 m; Supplementary Fig. 2h). Likewise, no differences were observed in grip strength (67.3±3.0 versus 62.8±3.1 g, n=12–14; Supplementary Fig. 2i). Finally, although two separate cohorts of Tbx15+/− males tended to show less voluntary wheel running than control mice (8.06±0.70 km per day for WT versus 7.16±0.61 km per day for Tbx15+/−, n=6), this was not statistically significant (Supplementary Fig. 2j). Thus, even though Tbx15 ablation caused both cellular and molecular changes in skeletal muscle, standard tests of muscle performance were not significantly affected.
Tbx15 ablation causes glucose intolerance and obesity
Factors that affect muscle fibre-type distribution and size can lead to alterations in whole-body physiology and metabolism22,23. Since homozygous Tbx15 knockout animals exhibited obvious developmental abnormalities that might also affect metabolic physiology, we focused our assessment on developmentally normal, heterozygous Tbx15 knockout animals. At 5 months of age, Tbx15+/− mice fed a standard chow (21% fat by calories) exhibited no differences from WT in body weight, fed or fasting blood glucose, and circulating insulin levels (Fig. 3a; Supplementary Fig. 3a–b). However, when challenged with an intraperitoneal glucose tolerance test, 5-month-old Tbx15+/− mice showed impaired glucose tolerance with a 43% increase in peak glucose levels (Fig. 3b). This occurred with no change in insulin tolerance, or insulin levels during the intraperitoneal glucose tolerance test (Supplementary Fig. 3c–d). At this age, Dual-energy X-ray absorptiometry (DEXA) analysis revealed that Tbx15+/− mice had no change in lean mass, but a 21% increase in fat tissue mass compared with controls (Fig. 3c). Haematoxylin/eosin staining, Oil Red O staining and biochemical analysis of the livers from 6-month-old Tbx15+/− animals also revealed a threefold increase in hepatic lipid accumulation, with no increase in muscle triglyceride content (Fig. 3d,e).
Indirect calorimetry of 5-month-old mice revealed that Tbx15+/− males had ∼12% decrease in average oxygen consumption during both the light phase (3,386+174 versus 3,038+75 ml kg−1 lean mass per hour in control versus Tbx15+/− mice) and dark phase (3,049+120 versus 2,688±53 ml kg−1 lean mass per hour) of the diurnal cycle (Supplementary Fig. 3d) with no change in respiratory quotient (Supplementary Fig. 3e). Although the differences in oxygen consumption did not quite reach statistical significance (Student’s t-test; P=0.1), similar trends were observed in three separate cohorts of mice. This tendency of decreased oxygen consumption was associated with a significant 25–30% decrease in spontaneous activity of Tbx15+/− mice during both the light phase (93±8 versus 64±5 counts per hour in control versus Tbx15+/− mice per hour) and dark phase (217±21 versus 166±29 counts per hour) of the diurnal cycle (Fig. 3f). Since Tbx15 is almost exclusively expressed in skeletal muscle, with no detectable expression in the brain during any period in development, the reduction in activity after Tbx15 ablation is presumably due to its effects on the skeletal muscle. Thus, although heterozygous deletion of Tbx15 does not impair exercise-induced activity, it does result in decreased spontaneous activity that, at least in part, contributes to an increase in adiposity and hepatosteatosis.
Tbx15 regulates oxidative capacity and AMPK signalling
To further investigate the mechanisms underlying the loss of glycolytic fibres and increase in oxidative fibres observed in the mice with reduced Tbx15, we created C2C12 myoblast cell lines with a stable knockdown of Tbx15 expression (shTbx15 cells) and compared them with C2C12 cells with stable overexpression of Tbx15 (pBABE-Tbx15). qPCR and western blot analysis demonstrated that Tbx15 mRNA and Tbx15 protein were reduced >90% in shTbx15 cells and increased ∼20-fold in pBABE-Tbx15 cells (Fig. 4a).
AMPK signalling has been shown to be both necessary and sufficient to transform glycolytic muscle fibres into oxidative fibres8,10. Western blot analysis of control and shTbx15 myoblasts demonstrated a robust increase in phosphorylation of AMPK on Thr172 and its downstream substrate acetyl-CoA carboxylase (ACC) on Ser79 compared with controls (Fig. 4b), with no change in total levels of AMPK or ACC. This was confirmed in vivo in extracts of muscle from Tbx15+/− mice, which showed an increase in AMPK Thr172 and ACC Ser79 phosphorylation (Fig. 4c).
Activation of AMPK signalling usually leads to an enhancement of oxidative metabolism and increased oxygen consumption rates (OCRs)24. Indeed, although whole-body oxygen consumption tended to be decreased in Tbx15+/− mice, knockdown of Tbx15 in C2C12 cells resulted in a 33±6% increase in basal OCRs, and treatment of the shTbx15 and control myotubes with the AMPK inhibitor, compound C, reduces OCR in both the groups (Fig. 4d; Supplementary Fig. 4a). Conversely, overexpression of Tbx15 in C2C12 myoblasts resulted in a decrease in basal OCR by 34±1% compared with controls (Fig. 4d). Thus, reduction of Tbx15 both in vivo and in vitro leads to an activation of the AMPK signalling pathway. At the cellular level, this leads to an increase in oxidative metabolism in C2C12 myoblasts, but this effect is masked in vivo, as a result of the decreased activity of the Tbx15+/− mice.
AMPK has a well-established role in regulating insulin resistance, and activation of AMPK has been to shown to directly inhibit mTor signalling25,26. To examine the effects of AMPK activation we observed on ablation of Tbx15, we investigated insulin and mTOR signalling in fasted WT and Tbx15+/− muscle 15 min after intravenous administration of insulin. Although insulin stimulation robustly increased phosphorylation of AKT S473 and ERK T202/Y204, no differences in total AKT or ERK proteins or in the phosphorylation of these proteins was observed between WT and Tbx15+/− muscle (Fig. 4e). Likewise, qPCR analysis showed no changes in the glycolytic muscle-specific regulators of Akt activation, Baf60c and Deptor11 (Supplementary Fig. 4b–c) or other regulators of muscle fibre type, including PGC-1α, PGC-1β, RIP140, calcineurin or Ppar-delta (Supplementary Fig. 5). However, after insulin stimulation, levels of phosphor mTor S2448 as well as its downstream phosphorylation target p70S6K1 T389 were significantly reduced in Tbx15+/− muscle, with no changes in levels of total mTor or p70S6K1 (Fig. 4f). Since AMPK has been shown to repress activation of mTor signalling, these results are consistent with the marked activation of the AMPK signalling axis in Tbx15+/− muscle.
Tbx15 regulates Igf2 levels both in vivo and in vitro
To determine possible transcriptional targets of Tbx15, we performed microarray analyses on shTbx15 and pBABE-Tbx15 myoblasts. Differentially regulated genes and pathways were analysed focusing on genes that were oppositely regulated in the Tbx15 knockdown and overexpressing cells. Genes that were significantly downregulated in shTbx15 and upregulated in pBABE-Tbx15 (with fold change >1.5; q<0.10) are found in Supplementary Table 3, and genes that were significantly upregulated in shTbx15 and downregulated in pBABE-Tbx15 are shown in Supplementary Table 4. Gene set enrichment analysis and ingenuity pathway analysis of these data revealed that the Igf pathways were among the most significantly altered in the Tbx15 cellular models. In particular, the expression of the protein hormone Igf2 showed robust and opposite regulation in these Tbx15 cellular models.
qPCR analysis confirmed that the expression of Igf2 was increased over sevenfold in Tbx15 overexpressing cells and was decreased by over 70% in shTbx15 knockdown cells compared with controls (Fig. 5a). Since Igf2 is an important regulator of myogenesis and myoblast differentiation27, we investigated whether shTbx15 myoblasts that exhibit reduced Igf2 levels also had a defect in myogenesis. Indeed, compared with control myotubes, after 4 days of differentiation shTbx15 cells showed a reduced number of myotubes, but those that did form were both shorter and thicker than control differentiated cell (Fig. 5b, left panels). Although markers of myogenesis MyoD, Myf4 and myogenin were normally induced in the knockdown cells, Igf2 levels remain reduced in the differentiated shTbx15 myotubes (Fig. 5c). In further support of a combined role for Tbx15 and Igf2 in myogenesis, in situ hybridization and immunofluorescence studies demonstrate that Tbx15 (Supplementary Fig. 6a) and Igf2 were highly and co-expressed in the developing, myosin-positive muscles of embryonic day 14.5 (E14.5) WT mice (Fig. 5d). In agreement with the cellular models, Tbx15 homozygous embryos demonstrated reduced Igf2 expression in developing muscles (Fig. 5d). This result was confirmed by qPCR of mRNA isolated from dissected limb muscle of E14.5 embryos, which showed 60–65% reductions of Igf2 mRNA in both the Tbx15+/− and Tbx15−/− animals. The levels of Igf2 and Tbx15 were ∼10- and ∼3-fold higher, respectively, at E14.5 than at postnatal day 28 further suggesting a role for Tbx15 during development (Fig. 5e; Supplementary Fig. 6b).
Igf2 mediates Tbx15 action in skeletal muscle
Since Igf2 is regulated by Tbx15 levels and since Igf2 has been implicated to play a role in myogenesis and fibre-type specification28, we examined the muscles of Igf2 knockout mice. As expected, muscle weights of Igf2 knockout mice were smaller (∼60%) than littermate controls29, but there was no obvious change in fibre-type composition. However, like Tbx15+/− and Tbx15−/− animals, muscle fibre size in the Igf2 knockout mice was markedly increased despite the decreased muscle mass (Fig. 5f). Although Igf2 has been thought to be expressed specifically in fast-twitch fibres at E14.5 (ref. 28) and distinct fast- and slow-twitch muscle fibres are found during embryonic development, we found that skeletal muscle of E14.5 embryos expresses fast myosin heavy chain, irrespective of genotype (Supplementary Fig. 6c). However, by day 28, SDH staining revealed clear differences in fibre composition between WT and Tbx15+/− muscles (Supplementary Fig. 6d). Taken together, these data suggest that changes in Igf2 levels during embryogenesis do not lead to changes in fibre type, however, loss of Igf2 at this time may lead to alterations in myogenesis and fibre size.
To test whether the change in morphology and size exhibited by shTbx15 myotubes was due to lack of Igf2 expression, we treated shTbx15 and control myotubes with recombinant Igf2 (10 ng ml−1) in vitro throughout the 4 days of differentiation. Whereas the morphology of control myotubes was not affected by Igf2 treatment, Igf2 treatment partially rescued both the reduced myotube density, as well as the shorter, thicker morphology exhibited by the shTbx15 myotubes treated with vehicle only (Fig. 5b). Thus, Igf2 appears to be one mediator of Tbx15 action in skeletal muscle with specific effects on regulation of myogenesis and fibre size.
Tbx15 regulates IGF-2 transcription via an indirect mechanism
Gene regulation and enhancer elements that regulate IGF-2 expression have been extensively studied. The distal enhancer region (CS9) and the proximal promoter region (P3) that are necessary and sufficient for muscle-specific expression of IGF-2 have been previously been defined30. Luciferase constructs driven by the P3 promoter element or a combination of the P3 promoter and CS9 enhancer region were transiently transfected into control and shTbx15 C2C12 myoblasts. Transcriptional activity from both the P3 promoter element or the P3 promoter and CS9 enhancer region was reduced by ∼27% (P<0.05) in shTbx15 cells compared with control cells in both cases (Supplementary Fig. 6e). To see whether Tbx15 was directly responsible for this effect, we co-transfected the luciferase constructs with different amounts of a Tbx15 expression plasmid into a heterologous cell line, 293FT cells, that do not express Tbx15. In this context, Tbx15 did not transactivate the IGF-2 promoter luciferase constructs (Supplementary Fig. 6f). Taken together, these data suggest that Tbx15 transcriptionally regulates IGF-2, but this effect is most likely indirect.