Although evidence suggests that OTS have a profound impact on skeletal muscle phenotype due to a change in the anabolism/catabolism balance, it remains poorly understood whether OTS induces morphological and contractile properties alterations of skeletal muscle. The purpose of this study was to test the hypothesis that high‐intensity resistance training with insufficient recovery time between bouts, which could potentially lead to overtraining, may result in a decrease of fiber types cross‐sectional area (CSA), alter the muscle contractile activity observed through changes in fiber‐type frequencies and MHC isoform content in rat skeletal muscle. We analyzed the plantaris muscle because it is highly recruited in our model of high intensity resistance training and because it possesses a mixture of slow and fast twitch MHC‐based phenotype. To our knowledge, few studies have investigated contractile changes, specifically MHC expression and skeletal muscle CSA during overtraining.

In recent years, the high incidence of OTS in athletes has attracted interest from researchers to identify the possible causes and effects of this phenomenon (McKenzie, 1999 ; Kentta et al., 2001 ). A variety of hypotheses have been proposed to account for OTS and considerable evidence suggests that muscle injury is frequently the initiator of OTS. Naturally, training and competition are associated with a mild muscle tissue trauma followed by recovery. When adequate recovery is allowed, it results in an “overshoot” phenomenon, an adaptive process often called “adaptive microtrauma” (Smith, 1999 ). However, when intensity/volume training is abruptly increased and rest/recovery time between bouts is insufficient, mild trauma could develop into a more chronic, severe form of tissue trauma. Thus, progression from the benign adaptive microtrauma stage may exacerbate the initial injury (Stone et al., 1991 ) resulting in a subclinical injury in the overtrained athlete. Muscle injury induces a reduction in strength (Fatouros et al., 2006 ) due to extensive muscle damage (Seene et al., 1999 ), swelling in the injured area, soreness, edema (Jamurtas et al., 2000 ; Fatouros et al., 2006 ), and local inflammatory response (Smith, 2004 ). Also, Petibois et al. ( 2003 ) demonstrated that overtrained individuals have higher amino acid and lower protein blood accumulation in response to exercise than well‐trained individuals. This fact suggests a protein catabolism for amino acid supply during exercise in overtrained individuals. Moreover, a decrease in the testosterone/cortisol ratio has also been suggested as a marker of “anabolic‐catabolic balance” and as a tool in OTS diagnosis (Budgett, 1998 ).

Skeletal muscle adaptation to athletic training generally involves applying a progressive overload, which implies workload beyond a comfortable level to optimize performance (Fry et al., 1991 ; Stone et al., 1991 ). Unfortunately, there is a fine line between improved and reduced performance. When the exercise training leads to a repetitive muscle trauma, due to high intensity/volume training, associated with insufficient rest/recovery time between bouts, harmful effects can occur, in a state called overtraining (Kuipers and Keizer, 1988 ). Recovery from overtraining syndrome (OTS) may require weeks to months of absolute rest or greatly reduced exercise training. The term over‐reaching or “short‐time” overtraining describes the less severe form of overtraining, in which recovery generally occurs within days to weeks (Lehmann et al., 1993 ). The line between over‐reaching and OTS is difficult to determine because both can show one or more of the following symptoms: accentuated catabolic state; physiological, immunological, and biochemical alterations; and increased incidence of injury and mood alterations (Armstrong and VanHeest, 2002 ; Meeusen et al., 2006 ).

Skeletal muscle plays an important role in determining success in competitive sports. This tissue shows substantial heterogeneity as a result of distinct fiber types and myosin heavy chain (MHC) isoforms (Schiaffino and Reggiani, 1996 ). Mammalian single muscle fiber analysis revealed the presence of pure (expressing a single MHC isoform) and hybrid fibers (expressing two or more MHC isoforms) in different muscles, which give this tissue a wide functional and metabolic diversity (Pette and Staron, 2000 ). Adult rat limb muscles express four different MHC isoforms: Types I, IIa, IIx/IId, and IIb in fiber Types I, IIA, IIX/IID, and IIB, respectively (Bar and Pette, 1988 ). The differential expression of MHC isoforms in different muscles reflects their functional responses, such as contractile and metabolic behavior. These responses can also be affected by a variety of stimuli, including chronic stimulation, removal of a synergist muscle, endurance exercise, and heavy resistance training (Oakley and Gollnick, 1985 ; Staron et al., 1990 ; Green et al., 1999 ; Sharman et al., 2001 ). It is generally accepted that skeletal muscle can adapt to progressive physical training via both a quantitative mechanism, based on changes in muscle mass and fiber size, and a qualitative mechanism, based on changes in fiber type and MHC content (Schiaffino and Reggiani, 1996 ). In this context, histochemical and biochemical studies have reported that resistance training promotes significant alteration in MHC content, enhancing physical performance (Sharman et al., 2001 ; Campos et al., 2002 ; Harber et al., 2004 ).

Statistical analyses were performed using a software package (SPSS for Windows, version 13.0). To assure that the data were stable, the statistical procedure was performed after the preliminary study of the variable related to normality and equality of variance between the groups, using a statistical power of 80% for the comparisons. Fiber‐type frequency data were analyzed using the Goodman Test for contrasts between and within multinomial populations (Goodman, 1964 , 1965 ) to assess differences between the groups. Statistical comparisons between the groups were made using Analysis of Variance for the two‐factor model (Zar, 1999 ) for each of body weight, food intake, and MHC isoform content values. When significant main effects were revealed, specific differences were assessed using Tukey's post hoc comparisons. Data are expressed as Mean ± Standard Deviation (SD). Differences were considered significant when P < 0.05.

MHC isoform analysis was performed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS‐PAGE) in triplicate (maximum 5% variation). The protocol for analyzing the specimens was based on Talmadge and Roy ( 1993 ) and Mizunoya et al. ( 2008 ) with modifications for single‐fiber analyses (Staron, 1991 ). Eight histological sections (12 μm thick) were collected from each whole muscle sample and placed in a solution (0.5 mL) containing glycerol 10% (w/vol), 2‐mercaptoethanol 5% (vol/vol), and sodium dodecylsulfate (SDS) 2.3% (w/vol) in a Tris/HCl buffer 0.9% (pH 6.8; w/vol). The final solution was shaken for 1 min and heated for 10 min at 60°C (Campos et al., 2002 ). Portions (20 μL) of the extracts were submitted to electrophoresis reaction (SDS‐PAGE 8%) using a 4% stacking gel, for 26 hr at 180 V, where the maximum current was limited to 13 mA. The gels were stained with Coomassie Blue (Bar and Pette, 1988 ) and used to identify the MHC isoforms according to their molecular weight showing bands at the MHCI, MHCIIa, MHCIId, and MHCIIb levels (Fig. 3 ). The gels were photographed, the images captured by VDS Software (Pharmacia Biotech) and their relative percentages were quantified by densitometry using Image Master VDS Software (version 3.0). Identification of plantaris MHC isoforms was accomplished by comigration of EDL muscle samples using a control animal as pattern.

At the end of the experiment, animals were anesthetized with pentobarbital sodium (40 mg/kg IP) and sacrificed by decapitation. Measurements throughout the experiment included weekly animal body weight and food intake. Plantaris muscle was harvested and the middle portion frozen in liquid nitrogen at −156°C. Samples were kept at −80°C until use. Histological sections (12 μm thick) were obtained in a cryostat (JUNG CM1800, Leica Germany) at −24°C to determine muscle fiber‐type frequency and cross sectional area (CSA), using myofibrillar adenosine triphosphatase (mATPase) histochemistry after preincubation at pH 4.2, 4.5, and 10.6 (Guth and Samaha, 1969 ; Brooke and Kaiser, 1970 ). Pure (Types I, IIA, IID, and IIB) and hybrid muscle fibers (Types IIC, IIAD, and IIBD) were identified based on their staining intensities (Staron et al., 1999 ; Fig. 2 ). No attempt was made to delineate subtypes IIDA and IIDB (Staron and Pette, 1993 ). Muscle fiber‐type frequency and CSA of ∼200 fibers from each animal were determined using an Image Analysis System Software (Leica QWin Plus, Germany). Two regions of the plantaris muscle were analyzed. A superficial‐white (SW) portion with a higher percentage of slow fibers (Types I and IIA) and the deep‐red (DR) portion containing a higher percentage of fast fibers (Type IIB; Staron et al., 1999 ).

Trained group was submitted to a 12‐week high‐intensity resistance training program, similar to that described by Cunha et al. ( 2005 ; Table 1 ). Before the initial training program, animals underwent a 1‐week pre‐training (once a day) to familiarize them with the water and exercise. In this phase, rats individually performed jumping bouts into a 38 cm deep vat of water at 28–32°C (Fig. 1 ). Animals jumped into the water and surfaced to breathe without needing any direct stimulus to complete the jumping bouts. The depth was appropriate to allow each animal to breathe on the surface of the water during successive jumps. A jump was counted when the animal reached the surface and returned to the bottom of the vat, a repeatedly performed movement. The adaptation protocol consisted of increasing the number of sets (two to four) and repetitions (five to ten), carrying an overload of 50% body weight strapped to a vest on the animal's chest (Fig. 1 ). After the adaptation period, the Tr group began the progressive high‐intensity resistance training program for five consecutive days per week with 40 sec rests between each set. The volume (sets × jumps) and intensity (overload) of the training protocol are presented in Table 1 . This training emphasizes high workload with insufficient recovery time between bouts, and was designed to induce the muscle to support an overload beyond recommended to promote beneficial effects on muscle tissue. Bouts were performed between 2 and 3 pm.

An animal model was used to test the hypothesis that high‐intensity resistance training associated with insufficient recovery time between bouts could result in harmful effects on skeletal muscle. Training protocol intensity and volume throughout the experiment period were progressive and provided an effective manner of investigating skeletal muscle results. Two days after the end of training, morphometric and biochemical analyses were performed to assess muscle fiber type and MHC isoform content in a single isolated muscle. All experimental procedures were approved by the Biosciences Institute Ethics Committee, UNESP, Botucatu, SP, Brazil (Protocol No. 018/08‐CEEA) and were conducted according to the ethical principles in animal research adopted by the Brazilian College of Animal Experimentation ( www.cobea.org.br ). Male Wistar rats (200–250 g) were obtained from the Multidisciplinary Center for Biological Investigation (CEMIB, UNICAMP, Campinas, São Paulo, Brazil). They were housed in collective polypropylene cages (three animals per cage) covered with metal grids, in a temperature‐controlled room (22–24°C) under a 12:12 hr light–dark photoperiod and provided with unlimited access to standard rat chow and water. Rats were randomly divided into two groups: non‐trained (Co, N = 9) and trained (Tr, N = 9).

An atrophic effect was observed after 12 weeks of high‐intensity resistance training (Table 3 ). In Tr group, plantaris DR portion muscle fiber Types IIA and IID CSA's were significantly lower ( P < 0.05) than the Co group. The decrease in CSA was ∼15.6% for Type IIA and 19.2% for Type IID (Table 3 ). No significant changes in CSA were found in fiber Types I and IIB. However, there was a tendency for Types I and IIB CSA to decrease after the training protocol ( P = 0.08 and P = 0.09, respectively). Plantaris SW portion did not presented muscle fiber atrophy.

The representative SDS‐PAGE gel used to quantify MHC isoforms is shown in Fig. 3 and data from the two groups are summarized in Table 2 . In Tr group, MHCI and MHCIIa percentages decreased and MHCIIb percentage increased ( P < 0.05). These data were supported by alterations in fiber‐type frequency. A representative mATPase histochemical reaction used to measure fiber‐type frequency is shown in Fig. 2 and the corresponding data are presented in Fig. 4 . Tr group showed a significant ( P < 0.05) reduction in Types I and IIA muscle fiber frequency and an increase in Type IIBD fibers in the plantaris DR portion. In the plantaris SW, there was only a reduction in IID fiber‐type frequency.

DISCUSSION

Although it is easy to study resistance training in humans, it is difficult to determine the phenotypic muscle responses to this training. This is primarily due to the invasive nature of muscle biopsies and to the risks inherent in using human subjects. Considering the heterogeneity of muscle fibers in different muscle regions, a small muscle sample cannot accurately reflect total muscle response. To circumvent these problems, Tamaki et al. (1992) suggested a weight‐lifting protocol designed to induce hypertrophy in rat limb muscles. Here, we used a model of resistance training in a liquid medium, suggested as a variation of the model proposed by Tamaki et al. (1992), and a standardized training protocol, which included an imbalance period between exercise bouts and rest. The advantage of our animal model is that it provides the ability to perform analysis on whole muscle preparations, providing a more extensive examination of muscle phenotype adaptations during training. During training, our subjects muscle response was not affected by lifting technique, motivation, food, or any other psychological parameters. With these variables controlled, the purpose of this study was to test the hypothesis that a high‐intensity resistance training protocol with insufficient recovery time, could influence the morphology and MHC expression in rat skeletal muscle. The major findings of this study were: (i) a reduction in IIA and IID plantaris muscle fibers CSA and (ii) fiber‐type frequency change and MHC isoforms content transition from slow to fast.

Training programs can cause specific adjustments in muscle fibers phenotype to supply the body's needs and optimize physical performance (Siu et al., 2004). However, when recovery time, volume, and intensity are inadequate, they can cause a series of hormonal and physiological changes in the organism (Lehmann et al., 1999; Fry et al., 2005), such as alterations in muscle fibers, a decrease in strength and an increase in protein catabolism, leading to a state called overtraining (Fry et al., 1991; Lehmann et al., 1999). Petibois et al. (2003) observed that overtrained individuals presented higher amino acids and lower protein blood accumulation in response to exercise than well‐trained individuals, suggesting that proteins were catabolized for amino acid supply during exercise. This increased requirement for amino acids during hypermetabolism is partly satisfied by an augmentation of muscle proteolysis, the major storage pool of amino acids, and by a concomitant reduction in muscle anabolism (Smith, 1999). In addition, Seene et al. (2004) showed that during overtraining condition, degradation increased and a decreased muscle protein synthesis rate lead to a decrease in muscle mass, particularly in fast twitch muscles. These authors also observed in rat fast twitch muscles during overtraining, that the DNA content per muscle decreased due to a loss of myonuclei consequent to muscle atrophy (Seene et al., 2004). Collectively, the results of these studies suggest that during conditions of overtraining an increase in the catabolism/anabolism ratio could lead to muscle fiber atrophy. In our study, although measurements of the muscle protein and DNA contents were not performed, we did show that high intensity/volume training with insufficient recovery time between bouts, promoted a reduction in muscle fiber CSA's. Compared to the Co group, the Tr group exhibited a significant (P < 0.05) decrease in pure fiber IIA and IID CSA in the plantaris DR portion; there was also a tendency for pure fiber I and IIB CSA to decrease in this portion (P = 0.08 and P = 0.09, respectively). Our results are consistent with several studies which show a predominance of catabolic condition (Seene et al., 1999; Petibois et al., 2000; Seene et al., 2004) in situations with a persistent combination of excessive overload plus inadequate recovery (Jamurtas et al., 2000; Fatouros et al., 2006). Although the molecular events that underlie our findings remain unknown, these observations raise questions as to what signals and cellular conditions initiate muscle mass changes during overtraining conditions. Furthermore, we observed a loss of body weight in the Tr group. Although some studies have reported loss of appetite, due to the arduous training schedule (Mackinnon, 2000; Meeusen et al., 2006), our results showed no differences in food intake between groups (data not shown). Thus, the weight loss observed in the Tr group could also be related to the reduction in plantaris muscle fiber CSA's, although other factors such as loss of motivation, apathy, irritability, and depression could be related to the weight loss (Fry et al., 1991; Mackinnon, 2000).

Animal experiments have shown that overtraining decreases the number of satellite cells (Seene et al., 1999), which are cells, located under the basal lamina of skeletal muscle fibers and when activated, proliferate, differentiate, and fuse with muscle fibers (Rosenblatt et al., 1994). A decrease in activated satellite cells means that new myonuclei are not adding in muscle fibers and muscle atrophy can develops. With skeletal muscle atrophy, myonuclei numbers decrease, decreasing DNA units in overtrained muscle, thereby decreasing synthesis and increasing degradation rate of muscle proteins; this promotes the development of overtraining myopathy (Seene et al., 1999). Although the mechanisms responsible for muscle atrophy are not completely defined, several factors seem to be involved; these include reduced neuromuscular activity, systemic activation of neurohormones and inflammatory cytokines (Dalla Libera et al., 2001; Filippatos et al., 2005), myostatin/follistatin imbalance (Lima et al., 2010), and ubiquitin‐proteasome pathway activation (Schulze and Upäte, 2005). The ubiquitin‐proteasomal pathway, which includes the muscle‐specific E3 ligases, atrogin‐1/muscle atrophy F‐box (MAFbx), and muscle RING Finger 1 (MuRF1), is known to be a powerful contributor to muscle proteolysis (Bodine et al., 2001; Gomes et al., 2001). We did not evaluate the ubiquitin‐proteasomal pathway in our study, but it is possible that this molecular pathway may be involved in the control of gene expression related to skeletal muscle atrophy that occurred in our high‐intensity resistance training and insufficient recovery time model.

In addition, high resistance training has been shown to promote muscle fiber‐type modulation and MHC isoform changes (Campos et al., 2002; Ratamess et al., 2009). Several studies have reported the transition from fast‐to‐slow muscle fiber types and MHC isoforms after resistance training (Roy et al., 1997; Carroll et al., 1998; Sharman et al., 2001). Kesidis et al. (2008) observed a high percentage of fibers expressing MHCIIa and MHCI/IIa in bodybuilders, compared to non‐trained individuals. Similarly, Otis et al. (2007) in a study on tumor‐bearing rats submitted to functional overload, observed a reduction in MHCIIb isoform, associated with increased levels of MHCI isoform. On the basis of these studies, the modulation of fiber types and MHC isoforms toward the slowest contracting fibers, contributes to increase strength, muscle power, and fatigue tolerance. In contrast, in our study, the high‐intensity resistance training with insufficient recovery time between bouts changed MHC isoform content and fiber‐type frequency toward fast contracting activity in rat plantaris muscle. A decrease of MHCI and MHCIIa in the Tr group compared to controls were corroborated with a decrease of Types I and IIA fiber frequencies. Moreover, an increasing in MHCIIb in the Tr group was associated with a significant increase in the Types IIBD fiber frequency, which express more MHCIIb than MHCIId isoforms (Pette and Staron, 2000; Pette and Staron, 2001) in plantaris DR portion. However, the reduction in Type IID fiber frequency in the plantaris SW portion in Tr was not associated with a change in MHCIId content (P > 0.05). Contrary to previous resistance‐training studies that show a MHCIIb‐to‐MHCIIa transition within the fast fiber population (Campos et al., 2002; Harber et al., 2004), we observed a transition of MHCI and MHCIIa toward MHCIIb isoform content in plantaris muscle during high‐intensity/volume training with insufficient recovery time between bouts. This might indicate that the activity patterns of fiber Types I and IIA (MHCI and MHCIIa isoforms), more frequent in plantaris DR portion, which have relatively high oxidative potential, possibly were more susceptible to oxidative damage by reactive oxygen species during high volume exercise in rats plantaris muscle. In addition, Seene et al. (2004) also observed an increase of fast MHC isoforms after 6 weeks of endurance overtraining. These authors reported that overtraining syndrome in skeletal muscle may decrease capillarization impairing oxygen exchange between capillaries and muscle tissue which contributes to exercise intolerance. In this sense, considering that muscle fiber type is also classified according to the type of energy metabolism used (glycolytic or oxidative; Gunawan et al., 2007), our results together with those of Seene et al. (2004), suggest that overtraining situations can result in a decrease in muscle oxidative capacity.

In conclusion, high‐intensity resistance training with insufficient recovery time between bouts promoted muscle atrophy and a shift from slow‐to‐fast contractile phenotype in rat plantaris muscle. Although the molecular events that underlie our findings remain unknown, we think that the stimulation of a transcriptional pathway, for example, the ubiquitin‐proteasomal, which promotes muscle proteolysis (Bodine et al., 2001; Gomes et al., 2001), might occur as a result of overtraining. Alternatively, the decrease in number of satellite cells during overtraining (Seene et al., 1999) could also be a determinant factor in preventing muscle fiber regeneration. Although these possible hypotheses need to be tested, understanding skeletal muscle morphological and biochemical changes following resistance training with appropriate recovery could help physiologists and coaches to improve the supervision of athletic performance.