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. 2010 Mar 2;24(4):790–799. doi: 10.1210/me.2009-0345

The Transcription Factor ATF4 Promotes Skeletal Myofiber Atrophy during Fasting

Scott M Ebert 1, Alex Mas Monteys 1, Daniel K Fox 1, Kale S Bongers 1, Bridget E Shields 1, Sharon E Malmberg 1, Beverly L Davidson 1, Manish Suneja 1, Christopher M Adams 1
PMCID: PMC2852358  PMID: 20197309

Abstract

Prolonged fasting alters skeletal muscle gene expression in a manner that promotes myofiber atrophy, but the underlying mechanisms are not fully understood. Here, we examined the potential role of activating transcription factor 4 (ATF4), a transcription factor with an evolutionarily ancient role in the cellular response to starvation. In mouse skeletal muscle, fasting increases the level of ATF4 mRNA. To determine whether increased ATF4 expression was required for myofiber atrophy, we reduced ATF4 expression with an inhibitory RNA targeting ATF4 and found that it reduced myofiber atrophy during fasting. Likewise, reducing the fasting level of ATF4 mRNA with a phosphorylation-resistant form of eukaryotic initiation factor 2α decreased myofiber atrophy. To determine whether ATF4 was sufficient to reduce myofiber size, we overexpressed ATF4 and found that it reduced myofiber size in the absence of fasting. In contrast, a transcriptionally inactive ATF4 construct did not reduce myofiber size, suggesting a requirement for ATF4-mediated transcriptional regulation. To begin to determine the mechanism of ATF4-mediated myofiber atrophy, we compared the effects of fasting and ATF4 overexpression on global skeletal muscle mRNA expression. Interestingly, expression of ATF4 increased a small subset of five fasting-responsive mRNAs, including four of the 15 mRNAs most highly induced by fasting. These five mRNAs encode proteins previously implicated in growth suppression (p21Cip1/Waf1, GADD45α, and PW1/Peg3) or titin-based stress signaling [muscle LIM protein (MLP) and cardiac ankyrin repeat protein (CARP)]. Taken together, these data identify ATF4 as a novel mediator of skeletal myofiber atrophy during starvation.

Fasting induces ATF4 expression in skeletal muscle, which in turn promotes myofiber atrophy.

Expression of the basic leucine zipper (bZIP) activating transcription factor 4 (ATF4) (also known as CREB-2) is tightly regulated and leads to the activation of many genes involved in the cellular response to stress (1,2). The best-characterized pathway to ATF4 expression is mediated by phosphorylation of the eukaryotic translation initiation factor 2α (eIF2α) on serine residue 51, which occurs in response to various intracellular stress signals such as depletion of free intracellular amino acids (3,4,5,6). eIF2α phosphorylation reduces the amount of eIF2 · GTP · Met-tRNAi complex required for high levels of mRNA translation initiation. However, eIF2α phosphorylation also increases the level of ATF4 mRNA and preferentially increases ATF4 synthesis by reducing the inhibitory effects of upstream open reading frames in the ATF4 transcript. Thus, as a result of eIF2α phosphorylation, global protein synthesis is reduced, but levels of ATF4 mRNA and ATF4 protein are increased (7,8).

In mammals, adult skeletal muscle is the principal repository of amino acids and protein, and gross alterations in skeletal muscle amino acid and protein content (reflected as changes in the size of skeletal myofibers) underlie common conditions such as skeletal muscle atrophy. A classical cause of skeletal muscle atrophy is food deprivation, which reduces free intracellular amino acid levels, increases eIF2α phosphorylation, increases ATF4 mRNA levels, decreases global protein synthesis, and promotes proteolysis and autophagy in skeletal muscle (9,10,11,12). However, the consequences of ATF4 expression in skeletal muscle have not been examined. In the current studies, we tested the hypothesis that ATF4 might promote skeletal myofiber atrophy during food deprivation.

Results

ATF4 is required for skeletal myofiber atrophy during fasting

We used quantitative real-time RT-PCR (qPCR) to assess the effect of fasting on ATF4 mRNA in mouse tibialis anterior (TA) muscle. Relative to levels in nonfasted control mice, skeletal muscle ATF4 mRNA levels in fasted mice were increased (Fig. 1A). These data are consistent with the results of a previous oligonucleotide microarray study showing that fasting increases the level of ATF4 mRNA in mouse gastrocnemius muscle (11). With the goal of reducing fasting-mediated ATF4 expression, we inserted a small interfering RNA sequence known to knock down ATF4 expression (13) into the hairpin of a miR-30-based micro-RNA shuttle (14), thereby generating a plasmid encoding miR-ATF4. We then transfected miR-ATF4 plasmid into mouse TA muscles via electroporation. In each mouse, one TA muscle received miR-ATF4 plasmid, and the contralateral TA muscle received an equivalent amount of empty vector. Two weeks later, we subjected the mice to a 24-h fast and then compared ATF4 mRNA levels between the two TA muscles of each animal. Under nonfasting conditions, miR-ATF4 had little effect on levels of ATF4 mRNA (Fig. 1B). However, miR-ATF4 prevented the fasting-induced elevation of ATF4 mRNA (Fig. 1B).

Figure 1.

Figure 1

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An artificial micro-RNA targeting ATF4 (miR-ATF4) reduces skeletal muscle ATF4 mRNA and myofiber atrophy during fasting. A, Mice were allowed ad libitum access to food or fasted for 24 h, and then ATF4 mRNA levels in the TA muscles were determined by qPCR. Data are means ± sem from four experiments, each using pooled RNA from three mice. *, P < 0.01. B–E, Mouse TA muscles were transfected on d 0. On d 14, mice were subjected to nonfasting or fasting conditions for 24 h, as indicated, before the TA muscles were harvested. B, The left TA was transfected with 20 μg empty pU6 vector, and the right TA was transfected with 20 μg pU6-miR-ATF4. TA muscle ATF4 mRNA levels were determined by qPCR and normalized to levels in the left TA of nonfasted mice. Data are means ± sem from four experiments and a total of seven transfected muscles per condition. *, P < 0.02. C and D, TA muscles were transfected with 5 μg pCMV-GFP plus either 20 μg empty pU6 vector (left TA) or 20 μg pU6-miR-ATF4 (right TA). C, Fiber size distribution curves from a representative experiment. D, Mean diameters of transfected myofibers ± sem from five experiments each using one mouse. *, P < 0.01. E, TA muscles were transfected as described in B. TA muscle mRNA levels were determined by qPCR, and levels under fasting conditions were normalized to levels in TA muscles transfected with empty pU6 vector and harvested under nonfasting conditions, which are indicated by the dashed line. Data are means ± sem from four experiments and a total of seven transfected muscles per condition. *, P < 0.02. Additional micro-RNA controls may be found in Supplemental Fig. 1 (published on The Endocrine Society’s Journals Online web site at http://mend.endojournals.org).

To determine whether ATF4 is required for skeletal myofiber atrophy during fasting, we cotransfected miR-ATF4 plasmid and a plasmid encoding green fluorescent protein (GFP), which served as a transfection marker. Previous studies demonstrated that when a mixture of two plasmids (each expressing a distinct cDNA) is electroporated into skeletal muscle, more than 90% of transfected myofibers coexpress both cDNAs (15,16). Moreover, transfection of markers such as GFP (Supplemental Fig. 2) or enhanced GFP (eGFP) (17) does not alter myofiber size. Two weeks after transfection, we examined the effect of fasting on the size of transfected myofibers. In the absence of fasting, miR-ATF4 did not significantly alter myofiber size (Fig. 1, C and D). In contrast, under fasting conditions, miR-ATF4 reduced atrophy of transfected myofibers. This was reflected as a rightward shift of the fiber size distribution curve (Fig. 1C) and an increased average myofiber diameter (Fig. 1D). As a biochemical correlate of myofiber atrophy, we examined levels of mRNA transcripts encoding the E3 ubiquitin ligases atrogin-1/MAFbx and MuRF1, which are induced by a broad range of conditions that promote muscle atrophy (18,19). Figure 1E shows that miR-ATF4 reduced levels of atrogin-1/MAFbx and MuRF1 mRNAs under fasting conditions. These data suggest that increased ATF4 expression is required for skeletal myofiber atrophy during fasting.

eIF2α phosphorylation is known to increase ATF4 mRNA levels (7,8), and transfection of cDNA encoding a phosphorylation-resistant form of eIF2α (eIF2α-S51A) inhibits the effects of eIF2α phosphorylation in cultured cell assays (20). Thus, as a second means of reducing ATF4, we transfected TA muscles with plasmid DNA encoding eIF2α-S51A. As in the previous studies, the contralateral TA muscle was transfected with empty vector and served as an intra-subject negative control. Ten days after transfection, we examined the effect of fasting. The immunoblot in Fig. 2A confirmed expression of eIF2α-S51A construct, and Fig. 2B shows that eIF2α-S51A reduced the level of ATF4 mRNA under fasting conditions. Under nonfasting conditions, eIF2α-S51A did not significantly alter myofiber size (Fig. 2, C and D). However, under fasting conditions, eIF2α-S51A reduced myofiber atrophy (Fig. 2, C and D) as well as levels of atrogin-1 and MuRF1 mRNAs (Fig. 2E). These effects of eIF2α-S51A mimicked the effects of miR-ATF4 (Fig. 1) and provide a second line of evidence that increased ATF4 expression is necessary for skeletal myofiber atrophy during fasting.

Figure 2.

Figure 2

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A phosphorylation-resistant form of eIF2α (eIF2α-S51A) reduces skeletal muscle ATF4 mRNA and myofiber atrophy during fasting. A–E, Mouse TA muscles were transfected on d 0. On d 10, mice were subjected to nonfasting or fasting conditions for 24 h, as indicated, before the TA muscles were harvested. A and B, The left TA was transfected with 25 μg pcDNA3, and the right TA was transfected with 25 μg pCMV-eIF2α-S51A-FLAG; A, TA muscle protein extracts were subjected to immunoprecipitation with anti-FLAG monoclonal IgG, followed by SDS-PAGE and immunoblot analysis with anti-eIF2α polyclonal antiserum; B, TA muscle ATF4 mRNA levels were determined by qPCR and normalized to levels in the left TA of nonfasted mice, which were set at 1. Data are means ± sem from five experiments each using pooled RNA from three mice. *, P < 0.01. C and D, Mouse TA muscles were transfected with 2 μg pCMV-eGFP plus either 25 μg pcDNA3 (left TA) or 25 μg pCMV-eIF2α-S51A-FLAG (right TA); C, fiber size distribution curves from a representative experiment; D, mean diameters of transfected myofibers ± sem from three experiments using a total of seven mice. *, P < 0.01. E, TA muscles were transfected as described in A and B. TA muscle mRNA levels were determined by qPCR, and levels under fasting conditions were normalized to levels in TA muscles transfected with pcDNA3 and harvested under nonfasting conditions, which are indicated by the dashed line. Data are means ± sem from five experiments, each using pooled RNA from three mice. *, P < 0.01.

ATF4 overexpression reduces skeletal myofiber size in the absence of fasting

To determine whether ATF4 is sufficient to reduce myofiber size, we transfected TA muscles with plasmid encoding ATF4 and then examined transfected myofibers in the absence of fasting. The immunoblot in Fig. 3A confirmed heterologous expression of ATF4. In the presence of wild-type ATF4, many transfected fibers had the shrunken appearance characteristic of atrophy (Fig. 3B), and the fiber size distribution curve was shifted to the left (Fig. 3C). As expected, ATF4 overexpression was inhibited by miR-ATF4 (Fig. 3D), which shifted the fiber size distribution curve back to the right (Fig. 3E). These data indicate that increased ATF4 expression is sufficient to reduce skeletal myofiber size.

Figure 3.

Figure 3

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Overexpression of ATF4 reduces myofiber size in the absence of fasting. A, Mouse TA muscles were transfected with 20 μg empty pcDNA3 vector (left TA) or 20 μg pCMV-ATF4-FLAG (right TA) and then harvested under nonfasting conditions 11 d later. Skeletal muscle protein extracts were subjected to immunoprecipitation with anti-ATF4 polyclonal antiserum, followed by SDS-PAGE and immunoblot analysis with anti-FLAG monoclonal IgG. B and C, Mouse TA muscles were transfected with 5 μg pCMV-GFP plus either 20 μg empty pcDNA3 vector (left TA) or 20 μg pCMV-ATF4-FLAG (right TA) and then harvested under nonfasting conditions 11 d later; B, representative images in the absence or presence of pCMV-ATF4-FLAG (bar, 100 μm); C, representative fiber size distribution curves from one of eight experiments. D, Mouse TA muscles were transfected with 15 μg empty pcDNA3 vector or pCMV-ATF4-FLAG with or without 20 μg pU6 vector or pU6-miR-ATF4, as indicated. Muscles were harvested under nonfasting conditions 9 d after transfection, and skeletal muscle protein extracts were analyzed as in A. E, Mouse TA muscles were transfected with 5 μg pCMV-GFP plus 15 μg pCMV-ATF4-FLAG plus either 20 μg pU6 vector (left TA) or 20 μg pU6-miR-ATF4 (right TA) and then harvested under nonfasting conditions 12 d after transfection. Data are representative fiber size distribution curves from one of three experiments.

As an additional control, we transfected TA muscles with plasmid encoding a transcriptionally inactive ATF4 construct that contains a series of point mutations in the bZIP domain (ATF4ΔbZIP) (7,21). The immunoblot in Fig. 4A confirmed expression of the full-length mutant ATF4 construct. Whereas wild-type ATF4 reduced the average diameter of transfected myofibers by more than 25% (Fig. 4C), ATF4ΔbZIP did not reduce fiber size (Fig. 4, B and C). These results suggest that increased ATF4 expression reduces myofiber size by altering gene expression.

Figure 4.

Figure 4

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ATF4-mediated reduction in myofiber size requires an intact bZIP domain. A, Mouse TA muscles were transfected with 20 μg empty pcDNA3 vector (left TA) or 20 μg pCMV-ATF4ΔbZIP-FLAG (right TA) and then harvested under nonfasting conditions 11 d later. Skeletal muscle protein extracts were subjected to immunoprecipitation with anti-ATF4 polyclonal antiserum, followed by SDS-PAGE and immunoblot analysis with anti-FLAG monoclonal IgG. B and C, Left-sided mouse TA muscles were transfected with 5 μg pCMV-GFP plus 20 μg empty pcDNA3 vector. Right-sided TA muscles were transfected with 5 μg pCMV-GFP plus either 20 μg pCMV-ATF4-FLAG or 20 μg pCMV-ATF4ΔbZIP-FLAG, as indicated. Muscles were harvested under nonfasting conditions 11 d later. B, Representative fiber size distribution curves in the presence of pCMV-ATF4-FLAG or pCMV-ATF4ΔbZIP-FLAG from one of six experiments. C, Mean fiber diameters ± sem from at least six experiments per ATF4 construct. *, P < 0.01 using paired t tests.

ATF4 overexpression reproduces the effect of fasting on five skeletal muscle mRNAs

To investigate the effects of ATF4 on gene expression, we first examined the effect of ATF4 overexpression on atrogin-1 and MuRF1 mRNA levels, which were decreased by both miR-ATF4 and eIF2α-S51A (Figs. 1 and 2). However, in multiple experiments, we were unable to find any evidence of increased atrogin-1 or MuRF1 mRNA levels at early or late time points after ATF4 overexpression (Supplemental Fig. 3). Thus, although ATF4 was sufficient to reduce myofiber size (Figs. 3 and 4), it was not sufficient to induce atrogin-1 and MuRF1 mRNAs. This suggested that ATF4 might regulate other skeletal muscle mRNAs that were increased or decreased by fasting.

To test this idea, we used exon expression arrays to compare the effects of fasting and ATF4 on global TA muscle mRNA expression. First, to determine the effect of fasting, we compared TA mRNA levels in mice that were subjected to fasting or nonfasting conditions for 24 h. Second, to determine the effect of ATF4, we compared TA mRNA levels in mice whose left TA was transfected with empty plasmid vector and whose right TA was transfected with plasmid encoding ATF4. In both experiments, 16,756 transcripts were measured, and we defined significantly altered mRNAs as those mRNAs whose levels changed by 1.4-fold with a P value ≤0.01. Compared with ATF4 overexpression, fasting had a broader effect on skeletal muscle mRNA expression, increasing 146 mRNAs and decreasing 100 mRNAs. In contrast, ATF4 overexpression increased 25 mRNAs and decreased three mRNAs. A complete list of mRNAs whose levels were altered by fasting or ATF4 overexpression may be found in Supplemental Tables 1 and 2.

By comparing mRNAs that were altered by fasting with mRNAs that were altered by ATF4, we identified five mRNAs that were significantly increased by both conditions: CDKN1A (encoding p21Cip1/Waf1), CSRP3 [encoding muscle LIM protein (MLP)], PEG3 (encoding PW1/Peg3), GADD45A (encoding GADD45α), and ANKRD1 [encoding cardiac ankyrin repeat protein (CARP)]. Interestingly, four of these mRNAs (CDKN1A, CSRP3, PEG3, and GADD45A) were among the 15 mRNAs most highly increased by fasting (Fig. 5A).

Figure 5.

Figure 5

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ATF4 increases a subset of five fasting-responsive skeletal muscle mRNAs. A, To determine the effect of fasting on skeletal muscle mRNA expression, mice were allowed ad libitum access to food or fasted for 24 h before TA muscle RNA was harvested. RNA from three mice per group was pooled and then subjected to exon expression array analysis. mRNA levels under fasting conditions were normalized to levels under nonfasting conditions, which were set at 1. To determine the effect of ATF4 overexpression on skeletal muscle mRNA expression, mouse TA muscles were transfected with 25 μg empty pcDNA3 vector (left TA) or 25 μg pCMV-ATF4-FLAG (right TA). TA muscle RNA was harvested under nonfasting conditions 11 d later and then subjected to exon expression array analysis. mRNA levels in the presence of ATF4 (right TA) were normalized to levels in the absence of ATF4 (left TA), which were set at 1. Both experiments were performed four times to generate the final data. The figure shows the 15 mRNAs most increased by fasting as well as the effect of ATF4 on these mRNAs. Data are the mean fold changes induced by fasting or ATF4. For all mRNAs shown, the effect of fasting was statistically significant (P ≤ 0.01). The red boldface denotes transcripts that were also increased by ATF4 (P ≤ 0.01). B, Mice were subjected to nonfasting or fasting conditions for 24 h, and then TA muscle mRNA was harvested and analyzed by qPCR. mRNA levels under fasting conditions were normalized to mRNA levels under nonfasting conditions, which were set at 1 and are indicated by the dashed line. Data are the means ± sem from four experiments. P ≤ 0.03 for all mRNAs shown. C and D, Mouse TA muscles were transfected with 25 μg empty pcDNA3 vector (left TA) or 25 μg pCMV-ATF4-FLAG (right TA). TA muscles were harvested under nonfasting conditions 11 d later. C, TA muscle mRNA was analyzed by qPCR. mRNA levels in the presence of ATF4 (right TA) were normalized to mRNA levels in the absence of ATF4 (left TA), which were set at 1 and are indicated by the dashed line. Data are the means ± sem from seven experiments. P ≤ 0.01 for all mRNAs shown. D, An equal amount (100 μg) of total muscle protein extract from each condition was subjected to SDS-PAGE, followed by immunoblot analysis with anti-p21 monoclonal IgG or anti-MLP polyclonal antiserum. The membranes were stained with Ponceau S to confirm equal sample loading. The graph shows average densitometry signals ± sem from three experiments. Levels in the presence of ATF4 (right TA) were normalized to levels in the absence of ATF4 (left TA), which were set at 1 and are indicated by the dashed line.

We used qPCR to confirm fasting- and ATF4-mediated increases in these five mRNAs. As predicted by the exon expression arrays, fasting and ATF4 increased levels of CDKN1A, CSRP3, PEG3, GADD45A, and ANKRD1 mRNAs (Fig. 5, B and C). To determine whether the ATF4-mediated increase in CDKN1A and CSRP3 mRNAs led to increased levels of p21 and MLP, we performed immunoblot analysis of total skeletal muscle protein extracts in the absence or presence of ATF4 overexpression. Figure 5D shows that ATF4 increased p21 and MLP protein levels by more than 3-fold. These data suggest that ATF4 promotes myofiber atrophy by inducing a small but important subset of fasting-responsive genes.

Discussion

ATF4 is an evolutionarily ancient regulator of amino acid and protein metabolism and the cellular response to stress (1,2,3,4,7,8), but its effects in skeletal muscle were not known. In the current studies, we found that fasting increased the level of skeletal muscle ATF4 mRNA, which in turn was required for atrophy of skeletal myofibers. Moreover, increased ATF4 expression reduced myofiber size in the absence of fasting. These data identify ATF4 as a novel, inducible mediator of skeletal myofiber atrophy during starvation.

We also found that a phosphorylation-resistant eIF2α construct (eIF2α-S51A) reduced ATF4 mRNA and myofiber atrophy under fasting conditions. This result suggests that fasting increases ATF4 expression via eIF2α phosphorylation and is consistent with a recent report of increased skeletal muscle eIF2α phosphorylation during food deprivation (10). The downstream mechanisms of ATF4-dependent atrophy appear to be complex. ATF4 is a bZIP transcription factor capable of activating or repressing genes in a cell-type-specific manner (22). Our data indicate that an intact bZIP domain is required for ATF4-mediated atrophy and that ATF4 functions primarily as a transcriptional activator in skeletal muscle.

Interestingly, when we reduced ATF4 levels with miR-ATF4 or eIF2α-S51A, it reduced fasting-mediated induction of mRNAs encoding atrogin-1 and MuRF1, which catalyze proteolytic events during skeletal muscle atrophy (19,23,24,25,26). However, ATF4 was not sufficient to increase atrogin-1 or MuRF1 mRNAs, suggesting an alternate mechanism for ATF4-mediated myofiber atrophy. Indeed, by comparing global mRNA expression profiles induced by fasting or ATF4 overexpression, we identified five mRNAs (CDKN1A, GADD45A, PEG3, CSRP3, and ANKRD1) that were highly induced by both conditions. These data suggest the hypothesis that ATF4 reduces myofiber size by increasing expression of CDKN1A, GADD45A, PEG3, CSRP3, and ANKRD1 mRNAs.

We do not yet know whether these five mRNAs are necessary for fasting-mediated myofiber atrophy or whether they are sufficient to induce myofiber atrophy in the absence of fasting. However, there is some evidence that they might be involved. CDKN1A and GADD45A mRNAs encode well-described inhibitors of cellular growth (p21Cip1/Waf1 and GADD45α, respectively) (27,28,29), and PEG3 mRNA encodes a protein required for cancer cachexia (PW1/Peg3) (30). CSRP3 and ANKRD1 mRNAs encode proteins that are thought to mediate titin-based stress signaling from the sarcomere to the nucleus (MLP and CARP, respectively) (31). Both MLP and CARP have the capacity to translocate to the nucleus and alter gene transcription (32,33) but are also found in myofibrils, where they may play structural roles at their specific binding sites within the sarcomere (the Z-line and the I-band for MLP and CARP, respectively) (34,35). Interestingly, a recent study found that MLP interacts with cofilin 2 at the Z-disc and facilitates cofilin 2-mediated actin filament depolymerization, suggesting that induction of MLP might a play direct role in myofibril disassembly (36). Determining the roles of p21Cip1/Waf1, GADD45α, PW1/Peg3, MLP, and CARP in fasting-mediated myofiber atrophy is an important area for future investigation.

Skeletal muscle atrophy is a nearly universal consequence of illness and aging; however, we still lack targeted medical therapies to prevent or reverse it. The current studies focused on the role of skeletal muscle ATF4 during starvation, but previous work associated increased skeletal muscle eIF2α phosphorylation with cancer cachexia (37) and increased skeletal muscle ATF4 mRNA levels with cancer cachexia, insulin deficiency, uremia, and muscle denervation (18). Large increases in skeletal muscle CSRP3, ANKRD1, CDKN1A, and GADD45A mRNA levels have been observed after muscle denervation (32,38,39), and skeletal muscle ANKRD1 and CSRP3 mRNA levels are increased by insulin deficiency (40). Furthermore, CDKN1A, GADD45A, and PEG3 mRNAs are increased in age-related sarcopenia (41,42,43). These considerations suggest that the induction of ATF4 may have a generalized role in the pathogenesis of skeletal muscle atrophy. Further investigations of this possibility may suggest new therapeutic approaches for this common and debilitating condition.

Materials and Methods

Plasmids

The coding region of mouse ATF4 (NM_009716) was amplified from mouse L cell cDNA and then cloned into p3XFLAG-CMV10 (Sigma Chemical Co., St. Louis, MO) to place three copies of the FLAG epitope tag at the NH3 terminus and generate pCMV-ATF4-FLAG. pCMV-ATF4 (ΔbZIP)-FLAG was generated by site-directed mutagenesis of pCMV-ATF4-FLAG, substituting 292GLYEAAA298 for 292RYRQKKR298. pU6-miR-ATF4 was generated by first using polymerase extension of overlapping DNA oligonucleotides to insert the antisense sequence 5′-TTAAACTAAAGGAATGCTCTG-3′ into a human miR-30-based micro-RNA shuttle (14) and then cloning this micro-RNA shuttle into the pU6 vector, which consists of a mouse U6 promoter, a multiple cloning site, and an RNA polymerase III terminator sequence in the TOPO-BluntII vector (Invitrogen, Carlsbad, CA). The antisense sequence encoded by miR-ATF4 is complementary to mouse ATF4 but is not complementary to other mRNAs described in these studies. Moreover, the seed region of miR-ATF4 does not show complementarity to other mRNAs described in these studies. Human FLAG epitope-tagged eIF2α-S51A was generously provided by Dr. Glen N. Barber (20) and subcloned into pcDNA3 to generate pCMV-eIF2α-S51A-FLAG. pCMV-GFP and pCMV-eGFP were pcDNA3 encoding GFP or eGFP, respectively.

Mouse muscle transfection

All studies described in this manuscript used male C57BL/6 mice that were obtained from the National Cancer Institute at ages 6–8 wk and used for experiments within 3 wk of their arrival. One advantage of the adult mouse TA muscle as an experimental system is that it can be readily transfected with plasmid DNA via electroporation. Electroporation transfects nuclei of terminally differentiated adult myofibers, but not nuclei of satellite cells or connective tissue cells (44), and has been used to study several proteins that regulate adult skeletal myofiber physiology and morphology (15,23,25,44,45,46,47,48,49,50,51,52,53,54). In our hands, plasmid DNA expression occurs in approximately 40% of TA myofibers and is maintained for more than 3 wk after transfection. Before transfection, mice were anesthetized by an ip injection of 91 mg/kg ketamine and 9.1 mg/kg xylazine, after which the hind limbs were shaved with a straight razor, and the TA muscles were injected with 30 μl hyaluronidase solution (which was prepared by resuspending bovine placental hyaluronidase (Sigma) in sterile injectable 0.9% NaCl at a concentration of 0.4 U/μl). Two hours later, the mice were reanesthetized, and TA muscles were injected with plasmid DNA (which was prepared using endotoxin-free plasmid purification kits (QIAGEN, Valencia, CA) and resuspended in sterile injectable 0.9% NaCl at a concentration of 1 μg/μl). All TA muscle injections used a 30-gauge needle inserted 2 mm into the distal end of the TA muscle (at a 10° angle to the long axis of the muscle). After injection of plasmid DNA, the hind limbs were placed between two-paddle electrodes and subjected to 10 pulses (20 msec) of 175 V/cm (with 480-msec intervals between pulses) using an ECM-830 electroporator (BTX Harvard Apparatus, Holliston, MA). All animal procedures were approved by the Institutional Animal Care and Use Committee of the University of Iowa.

Analysis of protein levels

After harvest, mouse TA muscles were snap frozen in liquid N2 and homogenized in 1 ml ice-cold lysis buffer [50 mm Tris (pH 7.4), 150 mm NaCl, 1% (vol/vol) Triton X-100, 1.5 mm MgCl2, 1 mm sodium EDTA, 1 mm sodium EGTA, 25 μg/ml N-acetyl-leucinal-leucinal-norleucinal (ALLN), 1 μg/ml pepstatin A, 2 μg/ml aprotonin, 10 μg/ml leupeptin, 200 μm phenylmethylsulfonyl fluoride, 640 μm benzamidine, and a 1:100 dilution of phosphatase inhibitor cocktail 1 (Sigma)] using a polytron (Tissue Master 240; Omni International, Kennesaw, GA) for 1 min on setting no. 8. The homogenate was rotated at 4 C for 1 h to generate a total muscle protein extract. In experiments involving eIF2α-S51A-FLAG, ATF4-FLAG, and ATF4ΔbZIP-FLAG, we were unable to detect the heterologously expressed proteins by SDS-PAGE of total muscle protein extract, and thus we performed immunoprecipitations as follows: the total muscle protein extract was centrifuged at 4 C and 16,000 × g for 20 min to generate a Triton X-100 soluble protein extract. The protein concentration of this Triton X-100 soluble extract was measured using the BCA kit (Pierce, Rockford, IL), after which the extract was diluted to a concentration of 3 mg/ml in ice-cold lysis buffer (final volume 500 μl). We then added 2 μg anti-FLAG mouse monoclonal antibody (Sigma) or 2 μg rabbit anti-ATF4 polyclonal antiserum (Santa Cruz Biotechnology, Santa Cruz, CA) plus 50 μl protein G plus Sepharose beads (Santa Cruz) and rotated the samples at 4 C for 16 h. Immunoprecipitates were washed three times for 20 min with 1 ml lysis buffer and then mixed with 100 μl 1× sample buffer [50 mm Tris-HCl (pH 6.8), 2% SDS, 5% glycerol, 0.04% (wt/vol) bromophenol blue] containing 5% (wt/vol) 2-mercaptoethanol and heated for 5 min at 95 C. Immunoprecipitates were subjected to 10% SDS-PAGE (for eIF2α) or 8% SDS-PAGE (for ATF4) and then transferred to Hybond-C extra nitrocellulose filters (Millipore, Billerica, MA). Immunoblots were performed at room temperature using a 1:1000 dilution of rabbit anti-eIF2α polyclonal antiserum (Cell Signaling, Danvers, MA) or a 1:1500 dilution of mouse anti-FLAG monoclonal antibody (Sigma). To assess p21 and MLP protein levels, the protein concentration of total muscle protein extract was measured with the BCA kit, after which an aliquot was mixed with 0.25 vol 5× sample buffer [250 mm Tris-HCl (pH 6.8), 10% SDS, 25% glycerol, 0.2% (wt/vol) bromophenol blue] containing 5% (wt/vol) 2-mercaptoethanol and heated for 5 min at 95 C. An equal amount of protein from each sample (100 μg) was subjected to 10% SDS-PAGE and then transferred to Hybond-C extra nitrocellulose filters. Immunoblots were performed at 4 C for 16 h using a 1:750 dilution of mouse anti-p21 monoclonal IgG (Sigma) or a 1:1000 dilution of chicken anti-MLP polyclonal antiserum (Abcam, Cambridge, MA).

Skeletal muscle mRNA analysis by qPCR

After harvest, muscle samples were immediately placed in RNAlater (Ambion, Austin, TX). Total RNA was extracted using TRIzol solution (Invitrogen) and then treated with deoxyribonuclease I using the Turbo DNA-free kit (Ambion). First-strand cDNA was synthesized in a 100-μl reaction that contained 2 μg RNA, random hexamer primers, and components of the TaqMan reverse transcription kit (Applied Biosystems, Foster City, CA). Levels of ATF4, MuRF1, and atrogin-1 mRNAs were measured using oligonucleotide primers (Supplemental Table 3) that were designed using Primer Express software (Applied Biosystems). For these studies, the real-time PCR contained, in a final volume of 20 μl, 20 ng reverse-transcribed RNA, 167 nm forward and reverse primers, and 10 μl Power SYBR Green PCR Master Mix (Applied Biosystems). Levels of CDKN1A, GADD45A, PEG3, CSRP3, and ANKRD1 mRNAs were measured using mouse TaqMan Gene Expression Assays (Applied Biosystems), and the real-time PCR contained, in a final volume of 20 μl, 20 ng reverse-transcribed RNA, 1 μl 20× TaqMan Gene Expression Assay, and 10 μl TaqMan Gene Expression Master Mix. qPCR was carried out using a 7500 Fast Real-Time PCR System (Applied Biosystems). All qPCR were performed in triplicate, and the cycle threshold (Ct) values were averaged to give the final results. To analyze the data, we used the ΔCt method, with level of 36B4 mRNA serving as the invariant control, and used t tests to determine statistical significance.

Histological analysis

After harvest, TA muscles were immediately placed in 4% (wt/vol) paraformaldehyde at 4 C for 16 h and then moved to 30% (wt/vol) sucrose solution at 4 C for at least 24 h. The middle of the muscle belly was then cut transversely with a straight razor and embedded in tissue freezing medium, after which 10-μm sections from the midbelly were prepared using a Microm HM 505 E cryostat equipped with a CryoJane sectioning system (Instrumedics, Richmond, IL). Sections were mounted with Vectashield mounting medium (Vector Laboratories, Burlingame, CA) and examined on an Olympus IX-71 microscope equipped with a DP-70 camera and a GFP-specific epifluorescence filter. Image analysis was performed using ImageJ software, and transfected fibers were defined as fibers with a mean fluorescence of 25 or more arbitrary units. In each experiment, we measured the diameter of at least 200 transfected myofibers per muscle, using the lesser diameter method described elsewhere (55), and used paired t tests to compare the results from bilateral TA muscles of each animal.

Microarray analysis of skeletal muscle mRNA levels

After harvest, muscles were immediately placed in RNAlater (Ambion) and stored at −80 C until further use. Total RNA was extracted using TRIzol solution (Invitrogen) according to the manufacturer’s instructions. Microarray hybridizations were performed at the University of Iowa DNA Facility. Briefly, 50 ng total RNA was converted to SPIA amplified cDNA using the WT-Ovation Pico RNA Amplification System, version 1 (NuGEN Technologies, San Carlos, CA; catalog no. 3300) according to the manufacturer’s recommended protocol. The amplified SPIA cDNA product was purified through a QIAGEN MinElute Reaction Cleanup column (QIAGEN; catalog no. 28204) according to modifications from NuGEN. Four micrograms of SPIA amplified DNA were used to generate ST-cDNA using the WT-Ovation Exon Module version 1 (NuGEN Technologies; catalog no. 2000) and again cleaned up with the QIAGEN column as above. Five micrograms of this product were fragmented (average fragment size = 85 bases) and biotin labeled using the NuGEN FL-Ovation cDNA Biotin Module, version 2 (NuGEN Technologies; catalog no. 4200) per the manufacturer’s recommended protocol. The resulting biotin-labeled cDNA was mixed with Affymetrix eukaryotic hybridization buffer (Affymetrix, Inc., Santa Clara, CA), placed onto Mouse Exon 1.0 ST arrays, and incubated at 45 C for 18 h with 60 rpm rotation in an Affymetrix Model 640 Genechip Hybridization Oven. After hybridization, the arrays were washed, stained with streptavidin-phycoerythrin (Molecular Probes, Inc., Eugene, OR), signal amplified with antistreptavidin antibody (Vector) using the Affymetrix Model 450 Fluidics Station. Arrays were scanned with the Affymetrix Model 3000 scanner with 7G upgrade, and data were collected using the GeneChip operating software (GCOS) version 1.4. In Mouse Exon 1.0 ST arrays, the log2 hybridization signal reflects the mean signal intensity of all exon probes specific for an individual mRNA and is proportional to the level of that mRNA. To determine which mRNAs were present at a significantly different level (P ≤ 0.01), we used paired t tests to compare log2 hybridization signals (between samples in the absence or presence of fasting or between samples transfected in the absence or presence of ATF4 transfection) and then applied a secondary requirement of a 1.4-fold change. To restrict our analysis to those mRNAs with reasonably high levels of expression, we excluded mRNAs that were not increased to a mean log2 hybridization signal of at least 5 or decreased from a mean log2 hybridization signal of at least 5. All raw microarray data have been deposited in Gene Expression Omnibus (GEO, accession number GSE 20104).

Supplementary Material

[Supplemental Data]
me.2009-0345_index.html (703B, html)

Acknowledgments

We thank Drs. Michael Welsh, Peter Snyder, Jennifer Stern, and Steven Kunkel for invaluable discussions and critical review of the manuscript. We also thank Tammy Lisi and Heather Lei for excellent technical assistance, Tom Moninger and Chantal Allamargot of the University of Iowa Central Microscopy Core Facility for assistance with microscopy, and Thomas Bair of the University of Iowa DNA Facility for assistance with the microarray analysis.

Footnotes

This work was supported by a Career Development Award from the Department of Veterans Affairs, Veterans Health Administration, Office of Research and Development, and a Junior Faculty Award from the American Diabetes Association (to C.M.A.), a Predoctoral Fellowship in Molecular and Cellular Biology (NIH T32 GM073610 to S.M.E.), the University of Iowa Medical Scientist Training Program (D.K.F. and K.S.B.), and the University of Iowa Summer Undergraduate Medical Scientist Training Program (B.E.S.).

Disclosure Summary: The authors have nothing to disclose.

First Published Online March 2, 2010

Abbreviations: ATF4, Activating transcription factor 4; bZIP, basic leucine zipper; CARP, cardiac ankyrin repeat protein; eGFP, enhanced GFP; eIF2α, eukaryotic translation initiation factor 2α; GFP, green fluorescent protein; MLP, muscle LIM protein; qPCR, quantitative real-time RT-PCR; TA, tibialis anterior.

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Supplementary Materials

[Supplemental Data]
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me.2009-0345_1.pdf (246KB, pdf)

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