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Am J Physiol Endocrinol Metab 292: E533-E542, 2007. First published September 26, 2006; doi:10.1152/ajpendo.00229.2006
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Effects of streptozotocin-induced diabetes and physical training on gene expression of titin-based stretch-sensing complexes in mouse striated muscle

T. Maarit Lehti,1 Mika Silvennoinen,1,2 Riikka Kivelä,1,2 Heikki Kainulainen,2 and Jyrki Komulainen1,2

1LIKES Research Center for Sport and Health Sciences; and 2Neuromuscular Research Center, Department of Biology of Physical Activity, University of Jyväskylä, Jyväskylä, Finland

Submitted 16 May 2006 ; accepted in final form 21 September 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
In striated muscle, a sarcomeric noncontractile protein, titin, is proposed to form the backbone of the stress- and strain-sensing structures. We investigated the effects of diabetes, physical training, and their combination on the gene expression of proteins of putative titin stretch-sensing complexes in skeletal and cardiac muscle. Mice were divided into control (C), training (T), streptozotocin-induced diabetic (D), and diabetic training (DT) groups. Training groups performed for 1, 3, or 5 wk of endurance training on a motor-driven treadmill. Muscle samples from T and DT groups together with respective controls were collected 24 h after the last training session. Gene expression of calf muscles (soleus, gastrocnemius, and plantaris) and cardiac muscle were analyzed using microarray and quantitative PCR. Diabetes induced changes in mRNA expression of the proteins of titin stretch-sensing complexes in Z-disc (MLP, myostatin), I-band (CARP, Ankrd2), and M-line (titin kinase signaling). Training alleviated diabetes-induced changes in most affected mRNA levels in skeletal muscle but only one change in cardiac muscle. In conclusion, we showed diabetes-induced changes in mRNA levels of several fiber-type-biased proteins (MLP, myostatin, Ankrd2) in skeletal muscle. These results are consistent with previous observations of diabetes-induced atrophy leading to slower fiber type composition. The ability of exercise to alleviate diabetes-induced changes may indicate slower transition of fiber type.

skeletal muscle; cardiac muscle; microarray; muscle LIM protein; myostatin

IN SKELETAL AND CARDIAC MUSCLE, a sarcomeric noncontractile protein, titin, forms a spring from the Z-disc to the A-band. Titin is the only molecule that extends over half a sarcomere. It maintains the temporal and spatial assembly of the contractile filaments (32, 54). On the basis of its location and interactions with several structural and signaling proteins, titin is thought to be the backbone of the stress- and strain-sensing structure in striated muscle (45). Interestingly, gene expression of some of these proteins is affected by diabetes or insulin resistance (21, 42, 53).

At the Z-disc, titin interacts with telethonin (or T-cap), small ankyrin-1 (sAnk1), and obscurin (32). Telethonin is a linking protein for several signaling and structural proteins, e.g., muscle LIM protein (MLP) and myostatin (16). Interaction of the MLP with myogenic regulatory factors (MRFs) in the nucleus and its effect on expression of brain natriuretic peptide (BNP) and atrial natriuretic factor (ANF) make MLP a possible stretch-regulatory protein (16).

The central I-band of titin contains an N2A region and a cardiac-specific N2B region. N2A interacts with calpain-3 and with three muscle ankyrin repeat proteins (MARPs): cardiac ankyrin repeat protein (CARP), ankyrin repeat domain protein-2 (Ankrd2), and diabetes-related ankyrin repeat protein (DARP). MARPs are induced following different stress conditions (16). N2B of cardiac titin interacts with skeletal muscle LIM protein-3 (SLIM3). Within the M-line region of the A-band, titin contains a kinase domain and a binding site for MuRF1, as well as other binding sites for calpain-3 and SLIM3 (16). Titin kinase regulates the transcriptional activity of the myofiber in response to a mechanical signal (33).

Gene expression of titin as well as some other proteins of the putative stretch-sensing complexes is affected by diabetes or insulin resistance. Titin gene expression increases in parallel with insulin sensitivity during an exercise intervention (53). Lack of myostatin suppresses both hyperglycemia and fat accumulation in the mouse model of type 2 diabetes (42), whereas DARP is upregulated in insulin-resistant animal models (21). The transcription of titin and actin interaction inhibitor S100 calcium-binding protein-A1 (S100A1) is increased in fast skeletal muscle in type 1 diabetes (60).

Moderate physical activity is recommended for the prevention of type 2 diabetes and for the management of both type 1 and type 2 diabetes. Our previous studies have shown physical training to attenuate diabetes-induced changes in the energy metabolism and mRNA expression of extracellular matrix proteins as well as in angiogenic proteins in skeletal muscle (22, 26, 35). Diabetic muscles become atrophic (34) and are more vulnerable to exercise-induced myofiber damage than nondiabetic muscles (8). The ability to increase muscle mass and the rate of protein synthesis depend on the severity of the disease (10, 11). Gene expressions of MLP and MARPs are known to increase and that of myostatin to decrease after physical activity (2, 36). However, the effects of training on titin stretch-sensing complexes are not well known.

We hypothesized that type 1 diabetes as well as training change the mRNA expression of the proteins of putative titin stretch-sensing complexes in skeletal and cardiac muscle. We also hypothesized that training alleviates the effects of diabetes.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals

Male NMRI mice (n = 60, Harlan, The Netherlands) were housed in standard conditions (temperature 22°C, light from 8:00 AM to 8:00 PM) with free access to tap water and food pellets (R36; Labfor, Stockholm, Sweden). The treatment of the animals was in accordance with the European Convention for the Protection of Vertebrate Animals Used for Experimental and Other Scientific Purposes and was approved by the University of Jyväskylä Ethics Committee for Animal Care and Use.

At the age of 10 wk, the animals were randomly assigned to diabetic and control groups. Experimental diabetes, similar to type 1, was induced by a peritoneal injection of streptozotocin (STZ; Sigma-Aldrich, Lyon, France, 180 mg/kg, 0.1 mol/l sodium citrate buffer, pH 4.5). An equal volume of buffer was injected into the control mice. Diabetic mice (urine glucose >200 mg/dl, 72 h after injection of STZ) were not treated with insulin during the study, and they showed symptoms of type 1 diabetes, such as polyuria and weight loss.

Diabetic and healthy animals were randomly assigned to 12 groups (n = 5 per group) that were either sedentary or trained for 1, 3, or 5 wk. Groups were named as follows: sedentary healthy mice (C1, C3, C5), trained healthy mice (T1, T3, T5), sedentary diabetic mice (D1, D3, D5), and trained diabetic mice (DT1, DT3, DT5). The training groups performed treadmill running (21 m/min, 2.5° incline, 1 h/day) for 5 days a week during the dark period. Animals were weighed once a week during the experiment as well as at the beginning and end of the study.

The trained mice were euthanized 24 h after the last training bout by cervical dislocation together with their respective sedentary controls. Blood and muscle samples were taken immediately. Calf muscles (soleus, gastrocnemius, and plantaris) and heart were removed, weighed, and frozen in liquid nitrogen. The proximal part of the left quadriceps femoris muscle was snap-frozen in isopentane (–160°C) cooled with liquid nitrogen.

Procedures

Serum glucose and citrate synthase activities of the calf muscle complex were measured to evaluate the severity of diabetes and the adequacy of exercise, respectively (35). Quadriceps femoris muscles from mice trained for 5 wk and their respective controls were used for measurement of cross-sectional area of muscle fibers (26).

Total RNA was extracted from the left calf muscle complex and cardiac muscle with TRIzol Reagent (Invitrogen, Carlsbad, CA) and further purified with RNeasy columns (Qiagen, Valencia, CA) according to the manufacturer’s protocols. The concentration and purity of RNA were determined by spectrophotometry at wavelengths of 260 and 280 nm. RNA integrity was tested by agarose gel electrophoresis. For the microarray analysis, an equal amount of RNA from each sample was pooled within each group, resulting in 12 arrays each from skeletal and cardiac muscle. Individual RNA samples were used for the real-time PCR.

Pooled RNA samples were analyzed with an Affymetrix Gene Chip MG U74Av2 (Affymetrix, Santa Clara, CA). The microarray analyses were performed by the Finnish DNA Microarray Centre at Turku Centre for Biotechnology according to the manufacturer’s instructions. Arrays were scanned using a GeneArray Scanner G2500A (Agilent, Palo Alto, CA). Image analysis and data processing were performed using Microarray Suite 5.0 (Affymetrix) and GeneSpring 6.1 (Silicon Genetics, Redwood City, CA) software as previously described (35). In brief, all chips were scaled (global scaling) to the target intensity of 50, and the data were subjected to robust normalization. The following comparisons were performed at each time point: trained (T) vs. control (C), diabetic (D) vs. C, trained diabetic (DT) vs. C, and DT vs. D. The complete data set is publicly available in the NCBI Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/; accession nos. GSE1659 and GSE4616).

Total RNA (5 µg) was reverse transcribed using a High-Capacity cDNA Archive kit (Applied Biosystems, Foster City, CA) according to the manufacturer’s instructions. Primers were designed for titin (NM_011652 [GenBank] ) and MuRF2 (XM_355438 [GenBank] ), and primers from the literature were used for CARP, Ankrd2, DARP, and MLP (2). Oligo Explorer software (http://www.uku.fi/~kuulasma/OligoSoftware/index.htm) was used for the primer design (Table 1). A sample of cDNA (4 ng RNA equivalent) was analyzed with SYBR Green PCR Master Mix (Applied Biosystems) and an ABI Prism 7700 Sequence Detection System (Applied Biosystems). All samples were run in triplicate along with the standards. Amplification was performed at the following temperatures: 95°C for 15 min, followed by 40 cycles at 94°C for 15 s, followed by 30 s at annealing temperature, 30 s at 72°C, and finally 15 s at the detection temperature. The annealing and detection temperatures are shown in Table 1. For GAPDH (Mm99999915_g1), telethonin (Mm00495557_g1), serum response factor (Mm00491032_m1), and MuRF1 (Mm01185221_m1), TaqMan probe-based analysis was used. The primer and probe set was purchased from Applied Biosystems, and the conditions that they recommended were used for the real-time PCR. Specific mRNAs in the sample were quantified according to the corresponding gene-specific standard curve. To compensate for variations in RNA quantity, the results were normalized to GAPDH. In the microarray data, GAPDH showed the steadiest expression in all conditions when normally used housekeeping genes were compared. The specificity of the amplified target sequence was confirmed upon observation of a single reaction product of the right size on an agarose gel and a single peak on the DNA melting temperature curve determined at the end of the reaction.


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  		Table 1. Primers for quantitative real-time PCR

 
Statistical methods

Nonparametric Kruskall-Wallis with Mann-Whitney U-tests were used to analyze differences in the quantitative PCR measurements. Statistical analysis of the microarray data was performed as previously described (35). In brief, a one-sided Wilcoxon’s signed rank test was used to determine which genes were expressed above the background (P ≤ 0.04) and to determine significant changes in expression between treatment groups (increased at P ≤ 0.0025 and decreased at P ≥ 0.9975). Calculation of the magnitude of the change in expression was based on differences between corresponding probe pair intensities across the two arrays and one-step Tukey’s biweight estimate statistics.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Body Weight, Serum Glucose, Citrate Synthase, and Myofiber Cross-Sectional Area

The body weight of the mice in groups D and DT decreased during the experiment (P < 0.001, D = –23 ± 10.2%, DT = –19.7 ± 10.6%, C = 0.1 ± 5.1%). At the beginning of the study there was no significant difference in weight between the study groups. The loss of body weight was largely due to muscle atrophy that was described in our previous publication (significant decrease in quadriceps femoris muscle fiber cross-sectional area in D and DT, P < 0.05, D = 2,469 ± 228, DT = 2,025 ± 357, C = 3,524 ± 456 µm2) (26). Serum glucose concentrations in DT and D mice were five times higher than in C mice (P < 0.001; D = 54.9 ± 5.7, DT = 50.5 ± 6.8, C = 10.3 ± 1.5 mmol/l). Serum glucose tended to be lower in DT than in D mice (P = 0.07). T and DT mice had higher citrate synthase activities than the respective untrained mice (P < 0.05; D = 400 ± 74, DT = 501 ± 60, C = 469 ± 99, T = 553 ± 140 nmol·min–1·mg–1), showing the effect of training. Citrate synthase activity was lower in the skeletal muscles of diabetic compared with healthy mice (P < 0.05). The results for body weight, glucose concentration, citrate synthase activity, and myofiber cross-sectional area are given in detail in recent publications (26, 35).

Gene Expression

STZ-induced diabetes affected the mRNA expression of several genes of the proteins that are suggested to be a part of the stretch-sensing system in mouse skeletal and heart muscles. From the complete microarray data (http://www.ncbi.nlm.nih.gov/geo/; accession nos. GSE1659 and GSE4616), genes of particular interest were selected on the basis of the literature. The microarray results are presented in three tables (Tables 2–4). Table 2 shows a list of the proteins that are proposed to form titin-based stretch-sensing complexes, the presence of the probes for the gene on the array, the presence of mRNA in samples, and change in gene expression compared with controls. Titin interaction with these proteins, either directly or indirectly, in the Z-disc, I-band, and M-line is illustrated in Fig. 1. In Tables 3 and 4, significant changes in expression in skeletal and cardiac muscle are shown in detail. The microarray results were verified and supplemented with quantitative PCR measurements of titin, telethonin, MLP, CARP, Ankrd2, DARP, serum response factor (SRF), MuRF1, and MuRF2 mRNA (Fig. 2). In addition, reliability of the microarray results has been shown previously with quantitative PCR measurements of six extracellular matrix (ECM) proteins (26, 35).


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  		Table 2. Observations of genes of putative titin-based stretch-sensing complexes on the microarray

 

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  		Table 4. Significant changes in mRNA levels of putative titin-based stretch-sensing complexes in cardiac muscle

 

Figure 1
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  		Fig. 1. Network of titin interactions. Gene symbols represent corresponding proteins. In addition, titin is separated into 4 segments in the Z-disc (TtnZ), I-band (TtnI), A-band (TtnA), and M-line (TtnM) to clarify the location of interactions on the sarcomere. Edges represent molecular interactions of 2 types: solid line, protein-protein interaction; dashed line, the effect on gene expression. Figure was drawn using Cytoscape software (http://www.cytoscape.org).

 

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  		Table 3. Significant changes in mRNA levels of putative titin-based stretch-sensing complexes in skeletal muscle

 

Figure 2
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  		Fig. 2. Quantitative PCR measurements of titin (Ttn, A), telethonin (T-cap, B), muscle LIM protein (MLP, Csrp3, C), cardiac ankyrin repeat protein (CARP, Ankrd1, D), ankyrin repeat domain 2 (Ankrd2, E), diabetes-related ankyrin repeat protein (DARP, Ankrd23, F), MuRF2 (G), serum response factor (Srf, H), and MuRF1 (I) mRNA presented as fold change from respective control values. D, sedentary diabetic; T, healthy trained; DT, trained diabetic. Values are means ± SD. **P < 0.01; *P < 0.05 vs. controls; #P < 0.05 vs. D; ++P < 0.01; +P < 0.05 vs. DT in skeletal muscle.

 
Titin

There were no probes for titin on the microarray, and no statistically significant changes in titin gene expression in skeletal muscle between experimental groups were found when measured by real-time PCR (Fig. 2).

Z-Disc

Titin interacts with the signaling and structural proteins of the Z-disc mainly through telethonin. These proteins include MLP, myostatin, isk/mink, and calsarcins. Transcription of telethonin as well as that of isk/mink was below the microarray detection limit. Since telethonin mRNA was previously shown to be one of the most abundant sarcomeric protein mRNAs (56), we also measured telethonin mRNA by real-time PCR. There were no differences in expression between groups at any time point (Fig. 2). One of the calsarcins, Myoz1, as measured in this study, showed no change in skeletal and was absent in cardiac muscle. Diabetes induced changes in the transcription of myostatin and MLP in skeletal muscle. A decrease in mRNA synthesis of myostatin was observed in D1, D3, and DT5, and an increase in the transcription of MLP was observed in D1, D3, and D5. Training attenuated the diabetes-induced changes in the level of MLP mRNA in DT1 and DT3 but not in DT5. The microarray results for MLP were further confirmed by real-time PCR (Fig. 2).

Diabetes affected the mRNA expression of MRFs, MyoD, and myf-6 in skeletal muscle differently. Gene expression of MyoD tended to decrease in the D group, but the change was significant only in D3. Transcription of myf-6 was increased in D and DT as well as in T1. Training alleviated the increase in DT1.

Gene expression of BNP and ANF increased in diabetic cardiac muscle in D and DT. BNP mRNA level, however, decreased in skeletal muscle in all of the diabetic groups.

At the Z-disc, titin also interacts with sAnk1, a small isoform of the Ank1 gene. In skeletal muscle, diabetes decreased mRNA expression of Ank1, whereas exercise attenuated the decrease in DT1 and DT3.

I-Band

In the area of the I-band, titin interacts with three MARPs: CARP, Ankrd2, and DARP. In skeletal muscle, gene expression of CARP and Ankrd2 increased in the diabetic sedentary group and physical training attenuated the diabetes-induced effect in DT1 and DT3 but not in DT5. The microarray results of these genes were further confirmed by the real-time PCR (Fig. 2). DARP transcription in skeletal muscle did not differ between groups at any time point when studied by the real-time PCR. In addition to MARPs, the gene expression of another ankyrin repeat protein, ankyrin-3 (Ank3), increased in D and DT as well as in T1 in skeletal muscle. In cardiac muscle, transcription of only one of two detected Ank3 isoforms increased.

The mRNA level of titin and actin interaction-inhibiting protein S100A1 decreased in D and DT in skeletal muscle and slightly increased in D3 and D5 in cardiac muscle.

M-Line

In skeletal muscle, the mRNA expression of titin kinase ligand nbr1 increased in the D group but not in the DT group. The mRNA level of the nbr1 ligand sequestosome-1 (Sqstm1 or p62) increased in D1 and DT1 in both skeletal and cardiac muscle. However, in skeletal muscle, the increase was alleviated by exercise. Gene expression of the Sqstm1 ligand MuRF2 and MuRF2 ligand SRF were measured by real-time PCR in skeletal muscle. The MuRF2 mRNA level decreased in D and DT3, whereas the SRF mRNA level increased in D1 compared with controls (Fig. 2). The transcription of titin-interacting protein SLIM3 decreased in the diabetic control group in both skeletal and cardiac muscle and tended to decrease also in DT. There were no probes for MuRF1 on the array, but real-time PCR measurements showed significant increase in MuRF1 mRNA level in D and DT5 compared with controls (Fig. 2). Training decreased mRNA expression of MuRF1 in T3 and T5 compared with controls but had no statistically significant effect in DT compared with D groups due to high variation.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The results of this study show diabetes-induced changes in mRNA expression of the protein complexes that are proposed to sense stretch in the Z-disc, I-band, and M-line (Fig. 1). However, no significant changes were observed in the transcription of titin, the backbone of these complexes. It is probable that muscle atrophy in the present study was not as extreme as in muscle denervation experiments, in which a decreased titin-actin ratio has been reported (6, 24), although fiber cross-sectional area of quadriceps femoris muscles of the present study mice was decreased nearly 30% after 5 wk of diabetes with or without exercise training (26). Training alleviated the diabetes-induced changes in most mRNA levels in skeletal muscle but only SLIM3 in cardiac muscle, obviously because of their different patterns of normal action (intermittent/constant).

Z-Disc

Titin interaction on the sarcoplasmic reticulum. The NH2-terminal domain of titin interacts with the small isoform of the Ank1 gene sAnk1, which is a transmembrane protein of the sarcoplasmic reticulum (30). The results of this study show that diabetes decreases the mRNA expression of Ank1 in skeletal muscle. Exercise attenuated the decrease in DT1 and DT5. However, it remains to be shown whether the protein level of sAnk1 is affected and whether that, in turn, has an effect on the sarcoplasmic reticulum.

Titin-based stretch-sensing apparatus in the sarcomere. In this study, there were no changes in mRNA expression of the titin-capping protein telethonin (or T-cap), which is a linking protein between titin and signaling proteins as well as structural proteins at the Z-disc. These proteins include MLP, myostatin, isk/mink, and calsarcins (16).

MLP interacts with titin through telethonin and {alpha}-actinin. Because of multiple interactions in the sarcomere (e.g., beta1-spectrin, nebulin-related protein, zyxin) and nucleus (MRFs, myogenin), MLP is thought to be a part of the titin-based stretch-sensing apparatus (45).

MLP mRNA is expressed more in slow (soleus) than in fast (white part of quadriceps femoris) muscle, indicating fiber type specificity (5). A shift toward slow myosin fibers during fasting, low-frequency stimulation, or mechanical overload increases MLP in skeletal muscle (9, 57). However, MLP expression is not downregulated in the transition from slow to faster myosin heavy chain (MHC) isoforms (soleus) by hindlimb suspension (57). MLP mRNA is upregulated after denervation of rat hindlimb muscles (1) and after lengthening contractions (2, 7). It has been proposed that MLP expression is under neural regulation (49). In the cardiomyocytes, a connection between MLP and the calcineurin anchorage to the Z-disc is established (17). In addition, other telethonin-binding proteins, calsarcins, also interact with calcineurin (12).

The novel finding observed in this study was that MLP transcription was increased in diabetic skeletal muscle. The increase in MLP transcription may be a consequence of change in prevalent myofiber type to slower fibers (43) or diabetes-induced neuropathy (52). The ability of exercise to prevent diabetes-induced changes in MLP mRNA expression in DT1 and DT3 may indicate deceleration in the transition to the slower fiber type or in the development of neuropathy.

Titin-based stretch-sensing apparatus in the nucleus. In the nucleus, MLP interacts with MRFs MyoD, Myf-6, and myogenin, thus enhancing their binding to DNA (29). All MRFs are markedly upregulated after denervation (40), and, at least, myogenin and MyoD gene expression increases after lengthening and isometric exercise (7, 19). Distribution of MyoD and myogenin is dependent on fiber type (20). Greater MyoD transcription occurs in fast glycolytic fibers, whereas myogenin is prevalent in slow oxidative fibers (20). Myogenin expression is also increased following endurance training, in parallel with the increase in the transcription of oxidative enzymes (51). In this study, diabetes decreased the mRNA level of MyoD, but that of myogenin was below the detection limit. Our preliminary results from the quadriceps femoris muscle of these mice show a transition from MHC IIb toward MHC IIa in D25 and DT25 (Touvra A, Silvennoinen M, Kivelä R, Lehti TM, Komulainen J, Kontro H, Vihko V, Kainulainen H, unpublished results). The decrease in MyoD mRNA level and the transition in MHC toward slower type myofibers are consistent with the findings of previous studies (43).

myf-6 is important in the maintenance of both mature myogenic and neuromuscular-specific gene expression (40). Abundance of myf-6 mRNA is enriched in the synaptic regions of the muscle fiber (40). In this study, myf-6 transcription was increased after training (T1) and in all diabetic groups. An increase in myf-6 may indicate changes in the maintenance of the neuromuscular junction.

Cardiac markers of mechnical load. BNP and ANF are cardiac markers of mechanical load and the embryonic gene program. In MLP knockout cardiomyocytes, normal stretch-induced enhancement of BNP and ANF mRNA expression is suppressed (27). The increase in mRNA levels of BNP and ANF in the cardiac muscle observed in this study suggest increased MLP activity in the cardiac muscle.

Myostatin, negative regulator of muscle growth. Myostatin is a negative regulator of muscle growth. Lack of myostatin causes hypertrophy and hyperplasia, thus resulting in an increase in muscle mass (16). Myostatin knockout muscles have a faster and more glycolytic phenotype compared with corresponding wild-type muscles (14). However, inhibition of myostatin in adult mice does not induce changes in muscle fiber types or in the expression of MHC isoforms. Lack of myostatin partially suppresses both fat accumulation and the development of hyperglycemia in mouse models of obesity and type 2 diabetes (Ay, Lepob/ob) (42). Endurance training decreases myostatin mRNA content in fast (gastrocnemius, vastus lateralis) but not in slow (soleus) skeletal muscles (36). In muscle atrophies induced by glucocorticoids or muscle unloading, myostatin expression is increased (36).

In this study, transcription of myostatin was decreased in D1, D3, and DT5 in skeletal muscle. Skeletal muscle may try to compensate for atrophy and hyperglycemia by reducing myostatin mRNA expression. Interestingly, myostatin belongs to the transforming growth factor (TGF)-beta superfamily, binds to TGF-beta1 type II receptor (e.g., Smad3 pathway), and inhibits expression of MRFs (4). Our previous study of mRNA expression of ECM genes for the same data set as studied here suggests inhibition of the Smad3 pathway in the diabetic group (35).

I-Band

In the I-band, titin interacts with myopalladin, calpain-3, and three MARPs: CARP, Ankrd2, and DARP (44). In addition to the I-band, MARPs and myopalladin are observed in the nucleus. This N2A-based signaling complex is thought to be stretch-regulated and link stress/strain signals to MARP-regulated gene expression (44).

CARP, marker of cardiac hypertrophy. Normally, CARP is expressed predominantly in cardiac muscle and only a small amount in skeletal muscle. After eccentric exercise (2, 7) and in certain pathological conditions (e.g., spinal muscular atrophy or denervation), expression of CARP is accelerated also in skeletal muscle (47, 55).

During wound healing, CARP is upregulated and activates angiogenesis (50). In vascular smooth muscle cells and endothelial cells, TGF-beta/Smads signaling induces a CARP transcription that might be linked to cell cycle inhibition (3, 23).

The results of this study show that CARP transcription is increased in diabetic skeletal muscle. However, identification of the expressing cells is necessary in determining whether regulation is involved in the stretch sensing or in the activation of angiogenesis as a compensatory mechanism for the reduced capillarization observed in these mice (26).

Ankrd2, the stretch response gene associated with slow muscle function. Ankrd2 is expressed in both skeletal and cardiac muscle (25). In addition to titin, Ankrd2 interacts with other proteins, such as myopalladin, calpain-3, and telethonin (28). Ankrd2 is expressed predominantly in type I skeletal muscle fibers (55). Interestingly, denervation of slow muscle (soleus) decreases the level of Ankrd2 to below the detection limit in 4 wk (41), whereas denervation of fast muscle (gastrocnemius) increases its expression (55). The level of Ankrd2 mRNA also increases after eccentric contraction (2). Gene expression was sensitive only to mode of contraction (eccentric vs. isometric) and not to stress (19).

To our knowledge, this is the first study to show a type 1 diabetes-induced increase in Ankrd2 mRNA expression. It is also noteworthy that training was able to alleviate the effect of diabetes for a period of 3 wk. The increase in Ankrd2 mRNA level in diabetic muscle may be a consequence of transition toward slower fiber type, which is decelerated by exercise.

DARP, insulin resistance-regulated stretch response gene. DARP is expressed in skeletal and cardiac muscle as well as in brown adipose tissue (21). DARP is located in the I-band, nucleus, and intercalated discs in the heart (44). The expression of DARP is elevated after eccentric and isometric contractions as well as after passive stretch in exercised and contralateral legs (2). Stretch increases DARP expression also in cardiac cells (44).

DARP is upregulated in cardiac muscle and downregulated in skeletal muscle in insulin resistance model animals (21). In skeletal muscle, alteration in DARP expression is regulated by energy supply (21). In this study, no changes were observed in the gene expression of DARP in skeletal muscle. This indicates that DARP mRNA expression reacts differently to type 1 than to type 2 diabetes.

Ank3, a likely titin-binding ankyrin repeat protein. In addition to MARPs, the mRNA expression of another ankyrin repeat protein, ankyrin-3 (Ank3), was increased in D and DT in our data. Ank3 (G107 isoform) has a titin-binding domain similar to that of MARPs, according to the sequence (44), and thus may be a part of the stretch-sensing complex. Interestingly, Ank3 isoforms are located in the sarcopalsmic reticulum and sarcolemma, particularly in the postsynaptic membrane (13).

Calcium-dependent titin-actin linkage. In cardiac myocytes, the PEVK segment of titin binds actin, but the linkage is inhibited by S100A1 in a calcium concentration-dependent manner (59). In addition, S100A1 is thought to inhibit microtubule assembly and PKC-mediated phosphorylation as well as to regulate Ca2+ release channel, desmin assembly, and transcription factors (18). In a previous study, STZ-induced diabetes was shown to increase mRNA levels of S100A1 in fast but not in slow skeletal muscle or cardiac muscle. However, the protein content decreases in skeletal but remains unchanged in cardiac muscle (60).

In this study, an increase in S100A1 transcription was observed in D3 and D5 in cardiac muscle but a decrease in D and DT in skeletal muscle (attenuation in DT1). The difference between this and a previous study in skeletal muscle mRNA results (60) may be due to the animal model (mice/rat), muscle (calf muscles/vastus lateralis), or phase of diabetes. Notwithstanding, both studies address the decrease in protein content that may disturb calcium-dependent regulation in the muscle.

M-Line

Mechanically regulated titin kinase domain. At the edge of the M-line, titin contains a catalytic serine-threonine kinase domain (37). In this study, titin kinase activity was not measured. However, mRNA expression of titin kinase ligand nbr1, which is thought to be mechanically induced (33), increased in D but not in DT skeletal muscle. In addition, transcription of Sqstm1, ligand of nbr1 and MuRF2, was increased in D1 in both skeletal and cardiac muscle and also in DT1 in cardiac muscle. These results suggest that diabetes affects signaling through nbr1 and Sqstm1 that may include several signaling pathways (e.g., titin kinase, PKC{zeta}, NF-{kappa}B).

The location of MuRF2 in the M-line or nucleus is regulated by mechanical signals (33). The nuclear ligand of MuRF2 is SRF, a transcription factor of immediate early genes (33). MuRF2 reduces nuclear SRF and increases it in the cytoplasm, thereby repressing the transcriptional activity of immediate early genes that are activated by hypertrophic stimuli such as mechanical stress. Denervation of muscle fibers induces MuRF2 localization to the nucleus (33). In addition to SRF regulation, MuRF2 maintains the order of the microtubular and intermediate filament structures in the muscle (39). In the present study, MuRF2 and SRF mRNA levels were measured in the skeletal muscle with real-time PCR. MuRF2 mRNA level decreased in D1, D3, and DT3, whereas SRF mRNA expression was increased in D1 and surprisingly decreased in T3. These results may indicate activation of SRF-mediated muscle gene expression. However, further studies of MuRF2 and SRF localization are needed to clarify the meaning of these observations in the signaling of diabetic skeletal muscle.

MuRF1, marker of skeletal muscle atrophy. A binding partner and a homologous protein of MuRF2, MuRF1, is involved in ubiquitin-proteasome proteolysis as a ubiquitin ligase. Thus, through ubiquitination, MuRF1 may regulate the turnover of titin and other myofibrillar proteins that interact with it, like nebulin and telethonin (15, 58). In addition to sarcomere, MuRF1 is located in the nucleus, where it interacts with transcription-regulating elements (38). Increased expression of MuRF1 mRNA observed in this study is well in line with Northern blot results of a previous study of several models of atrophy and is probably part of the activation of the ubiquitin-proteasome pathway (34). We also showed that training decreases mRNA expression of MuRF1, although the difference between D and DT groups was not statistically significant, probably due to high variation in the DT group.

SLIM3, marker of cardiogenic cells. SLIM3 links metabolic enzymes, like creatine kinase, to titin at the sarcomere (31). SLIM3 is also a coactivator for the androgen receptor (46) and interacts with the integrin {alpha}7beta1 receptor (48). Previously, SLIM3 mRNA was observed only in cardiac muscle by Northern blotting (46). However, our array results show its mRNA expression also in skeletal muscle. In D and DT, SLIM3 mRNA expression was reduced in both cardiac and skeletal muscle.

Conclusions

We have shown diabetes-induced changes in the mRNA levels of several fiber type-biased proteins (MLP, MyoD, myostatin, Ankrd2). The results of this study support previous observations that fast myofibers are affected first in the course of muscle atrophy (43), leading to the prevalence of slower fiber types. However, some of the proteins studied may also be involved in neuropathy-induced changes (e.g., MLP, myf-6, Ank3) in which neuropathy increases the expression of fast MHC, changing fiber type composition in a faster direction (52). Since we have observed differential changes in the calcineurin-dependent signaling pathway for the same data set as studied here (Silvennoinen M, Touvra AM, Kivelä R, Lehti TM, Vihko V, Kainulainen H, unpublished observation), interactions of Z-line proteins with calcineurin are particularly interesting. It is also tempting to speculate that titin-based stretch-sensing complexes may play a regulative role in diabetes-induced atrophy and the consequences of neuropathy.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
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REFERENCES
 
This study was supported by the LIKES Foundation and the Ministry of Education, Finland.


   ACKNOWLEDGMENTS
 
We thank Dr. S. Koskinen for critically reading the manuscript.


   FOOTNOTES
 

Address for reprint requests and other correspondence: T. M. Lehti, LIKES Research Center for Sport and Health Sciences, Rautpohjankatu 8, Viveca, FIN-40700 Jyväskylä, Finland (e-mail: maarit.lehti{at}likes.fi)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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 RESULTS
 DISCUSSION
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