Aberrant cell cycle reentry in human and experimental inclusion body myositis and polymyositis

Abstract

Inclusion body myositis (IBM), a degenerative and inflammatory disorder of skeletal muscle, and Alzheimer's disease share protein derangements and attrition of postmitotic cells. Overexpression of cyclins and proliferating cell nuclear antigen (PCNA) and evidence for DNA replication is reported in Alzheimer's disease brain, possibly contributing to neuronal death. It is unknown whether aberrant cell cycle reentry also occurs in IBM. We examined cell cycle markers in IBM compared with normal control, polymyositis (PM) and non-inflammatory dystrophy sample sets. Next, we tested for evidence of reentry and DNA synthesis in C2C12 myotubes induced to express β-amyloid (Aβ42). We observed increased levels of Ki-67, PCNA and cyclins E/D1 in IBM compared with normals and non-inflammatory conditions. Interestingly, PM samples displayed similar increases. Satellite cell markers did not correlate with Ki-67-affected myofiber nuclei. DNA synthesis and cell cycle markers were induced in Aβ-bearing myotubes. Cell cycle marker and cyclin protein expressions were also induced in an experimental allergic myositis-like model of PM in mice. Levels of p21 (Cip1/WAF1), a cyclin-dependent kinase inhibitor, were decreased in affected myotubes. However, overexpression of p21 did not rescue cells from Aβ-induced toxicity. This is the first report of cell cycle reentry in human myositis. The absence of rescue and evidence for reentry in separate models of myodegeneration and inflammation suggest that new DNA synthesis may be a reactive response to either or both stressors.

INTRODUCTION

Inclusion body myositis (IBM) is the most common acquired muscular disease of the aged. IBM has pathological features in common with Alzheimer's disease (AD) including β-amyloid and ubiquitin deposits, neurofilament and tau hyperphosphorylation and cell death (1). Aberrant cell cycle reentry involving neurons is an increasingly recognized phenomenon in some neurodegenerative diseases including AD, but not reported in IBM. Previous studies have shown aberrant cell cycle reentry in the brains of AD patients (2–4) and transgenic AD animal models (5–8). The evidence suggests that at an early time point and throughout the course of AD, neuronal death is correlated with ectopic reentrance into the cell cycle. It is hypothesized that aberrant entry at premitotic checkpoints, precedes and contributes to neuronal death. Accordingly, robust expression of cell cycle phase markers; cyclin D1 (the regulator of cyclin-dependent kinase (Cdk) 4/6 in the G0/G1 transition), proliferating cell nuclear antigen (PCNA; S phase) and cyclin B1 (the regulator of cell division cycle (Cdc) 2 in G2) are found in mild cognitive impairment (6,9,10). Signs of coordinate activation of cell cycle machinery leading to DNA replication were also reported in AD brain. Moreover, one source of tau-phosphorylation may involve the dysregulation of a cell cycle kinase, Cdc2k that is biochemically similar to Cdk5. The latter is a well-characterized tau kinase and is pathologically activated in AD (5,11). Cdc2k may also initiate apoptosis in some circumstances (4,12). Interestingly, Cdc2k and Cdk4/6 are ectopically expressed in another proteinopathy of skeletal muscle, myofibrillar myopathy (13). Furthermore, β-amyloid exposure is a known activator of cell cycle entry and apoptosis in neurons (14,15).

The biochemistry behind the decision of postmitotic cells to die instead of divide following abnormal cell cycle reactivation is unclear. The triggers for cell cycle reentry under these conditions are also not known. One hypothesis suggests that it is a physiological response to DNA damage and linked to the repair DNA strand breaks. Since oxidative stress is a common theme in neurodegeneration and DNA is a target for oxyradical attack, accumulation of oxidative byproducts in neurons can theoretically trigger a coordinate cell cycle entry response. The outcome is either to repair the damage or initiate apoptosis (16,17). An alternative to the DNA oxidative damage hypothesis proposes that dysregulation of the molecular chaperone–ubiquitin–proteasome system (UPS) triggers cell cycle reentry. The UPS can regulate the cell cycle in two opposing ways, by degrading either cyclin D1 or the Cdk inhibitors p21 (Cip1/WAF1) and p27 (18,19). Inhibition of the proteasome arrests neuronal cells at the G1/S boundary (9). There is also evidence for the regulation of various cell cycle stages by heat shock protein (HSP) chaperones 70, HSP90 and HSP27 (20,21). For instance, a specific inhibitor of HSP90, geldanamycin, has an anti-proliferative effect by halting the G0 to G1 transition, blocking the action of HSP90-specific client proteins Cdc37 and FKBP52 (22,23). In addition to changes in HSP levels possibly triggering cell cycle reentry, several mitogenic substances such as growth factors and reactive oxygen/nitrogen species can trigger cell proliferation. Cyclins E and D1 act as a switch between these mitogenic signals and Cdks during the G1 phase to promote cell cycle progression (24,25).

Some of the same oxidative stress and UPS abnormalities affect IBM as in AD. In both conditions, electrically specialized, postmitotic cell populations are vulnerable to these insults. In the present study, we examined cell cycle reentry in human IBM muscle for the first time and make comparisons with another muscle inflammatory disorder, polymyositis (PM) as well as age-matched normal controls and a set of non-inflammatory myopathic (NIM) conditions. Consistent with findings in AD, we found increased expression of cell cycle markers including cyclins E and D1, as well as induction of caspases and HSPs in IBM compared with normal control and NIM sets. Experimentally, we observed that new DNA synthesis was increased in β-amyloid (Aβ42)-bearing myotubes correlated with an increase in cell cycle markers. Furthermore, we found similar changes in PM cases, consistent with the notion that IBM and PM share inflammatory mechanisms. Based upon in vivo experimental evidence that inflammation (independent of amyloid) can also drive cycle reentry, we conclude that in IBM both degenerative (amyloid) and inflammatory (cellular) features are responsible for stimulating the myofiber through the G1/S boundry.

RESULTS

Cell cycle reentry in human IBM and PM

In preliminary tests for cell cycle deregulation in muscle samples from IBM and PM patients, immunocytochemical studies were performed using a general cell cycle maker. Biopsy specimens of IBM (n = 6), PM (n = 6) and control (n = 6) (see Supplementary Material, Tables S1 and S2 for patient demographics) patients were immunostained with anti-Ki-67 (Fig. 1A). Sections were counterstained with Hoechst dye (DAPI, blue) to identify nuclei. Sarcolemma were defined by differential interference contrast (DIC) and β-sarcoglycan immuno-decoration. Intra-myofiber nuclei were thus identified by DAPI/DIC/β-sarcoglycan overlay. The number of anti-Ki-67 decorated myofiber nuclei were counted in ten 40× fields from each section. Ki-67-positive myofiber nuclei were more numerous in IBM and PM samples compared with normal control group (Fig. 1B). While these represented a small fraction of total nuclei, they were statistically more frequent among intra-myofiber compared with extramysial [inflammatory and satellite cells (SCs)] nuclei. We confirmed the mature myoskeletal phenotype of Ki-67-positive cells by immunocytochemistry with markers: Pax7 (SCs), CD8 (cytotoxic T cells) and β-sarcoglycan (sarcolemma). Intra-myofiber Ki-67-positive nuclei were negative for co-staining with antibodies to Pax7 or CD8 in IBM and PM samples (Supplementary Material, Fig. S1A and B) and were located in the subsacolemma compartment (Supplementary Material, Fig. S1C). Various methods of SC counts in control human muscle yield differing but generally low values of ∼5% of all myonuclei (SC/N) (27,28), a frequency of ∼0.07 per myofiber (SC/F) (29,30) and 20–34 per mm² (31,32). By double Pax7 and Ki-67 evaluation, the proportion of these SCs that are activated is extremely low, up to a maximum of 1.3%, in control muscle (29). We counted such rare activated SC nuclei as were positive for both Pax7 and Ki-67 and found they were not increased in number per mm² of cross-sectioned IBM and PM muscle compared with control (1.58 ± 1.1, 3.19 ± 1.8, 2.45 ± 1.1 respectively, P > 0.2 versus control, n = 7 cases). Although total satellite number in IBM and PM is debated (28,30,32), most indicators suggest a significant reduction in their proliferation and activation state in inflammatory myositis (28,32,33). These results indicate that the increase in Ki-67 myofiber counts in IBM and PM is not accounted for by satellite activation.

Figure 1.

Induction of Cdc and characterization of inflammation and myodegeneration in human IBM and PM. Photomicrographs of immunofluorescently labeled Ki-67 (cellular proliferation marker, red) in representative muscle sections of IBM, PM and age-matched normal control (A). Patient demographics are given in (26). The mean number of Ki-67-positive myofiber nuclei in 6 random 40× fields was significantly increased in IBM and PM compared with normal control. Small but significant Ki-67 induction in both intra-myofiber and to lesser extent epimysial nuclei is noted (B). IBM and PM showed comparable significant increases in inflammatory infiltration and decreases in fiber diameter compared with normal control (C). The distribution frequency of fiber diameters in IBM, PM and normal control is given in (D). Colocalization of Ki-67 nuclei and Aβ deposits in human IBM (E). Serial quadriceps muscle sections from IBM were labeled with anti-Ki-67 and -Aβ (R1282). H&E and modified Gomori trichrome (G-T) histochemistry is shown. Ki-67 nuclei (arrows) were present with Aβ deposits (arrow head) in the same abnormal myofiber bearing rimmed vacuoles. All data presented are mean ± SEM. *P < 0.05 versus control (ANOVA). Scale bar: 50 μm. DIC, differential interference contrast.

To further characterize these specimens, the degree of inflammatory infiltration was measured in the same ten 40× fields by quantifying the % section area occupied by leukocyte clusters. WBC's were confirmed by anti-CLA uptake (not shown). As expected, both IBM and PM showed significantly increased inflammatory infiltrates compared with the control group (Fig. 1C). There was no positive correlation between the number of Ki-67-positive cells and the percentage of inflammatory infiltration in IBM and PM samples (R² = 0.30, P = 0.35). Moreover, the Ki-67 signal was myofiber nuclei-based, and not apparently from CD8 typed T lymphocytes (Supplementary Material, Fig. S1B). CD45 (leukocyte common antigen, LCA) immunofluorescence and H&E stain confirmed the cellular character of infiltrates in IBM and PM (Supplementary Material, Fig. S1D). Ki-67-positive myofiber nuclei did not correlate with CD45 lymphocytes (Supplementary Material, Fig. S1D). To further confirm diagnosis and comparable disease severity, a total of 97 ± 11 muscle fibers in each sample were measured for fiber diameter. As expected, the mean fiber diameter was similarly decreased in IBM and PM compared with the control group (Fig. 1C). The distribution frequency of fiber diameters in IBM (n = 754) or PM (n = 445) was left-shifted relative to control (n = 260) (Fig. 1D). In serial muscle sections from IBM patients, the Ki-67 reactivity could be assigned to β-amyloid and inclusion-bearing myofibers (Fig. 1E).

Myotubes expressing Aβ trigger the expression of cell cycle markers

In order to understand the mechanism of cell cycle reentry pertaining to IBM muscle, we performed in vitro cell culture experiments on cultured myotubes coinfected with Adv-TRE-Aβ42 and Adv Tet-On, an inducible expression system that produces Aβ42. Transgene expression was induced with doxycycline for 24 h followed by fixation and staining with a panel of cell cycle markers. The non-induced control myotubes (Fig. 2A, left three panel columns) showed no nuclear staining corresponding to the different cell cycle proteins. However, the doxycycline-induced myotubes (Fig. 2A, right three panel columns) showed nuclear staining with PCNA (S-phase marker; red) and Ki-67 (cellular proliferation marker; green), cyclin D1 (G1 to S phase marker; red) and cyclin E (G1 to S phase marker; red). We confirmed Aβ expression in the induced myotubes using antibody 6E10. In order to demonstrate that the cell cycle markers colocalize with Aβ expression, myotubes were double labeled with cell cycle markers and Aβ antibodies (6E10 or R1282; Supplementary Material, Fig. S2A). The non-induced myotubes showed no staining with Ki-67 or Aβ antibodies as expected (top row), whereas doxycycline-induced myotubes showed positive staining with both cell cycle markers and Aβ antibodies in same cells (rows 2–3, Supplementary Material, Fig. S2A). We counted all the myotubes with positive nuclei stained with the various cell cycle markers and all those decorated with Aβ antibodies in a total of 15 separate 40× fields. The correlation of Aβ expression with cell marker induction in the same cell is highest with PCNA, followed by Ki-67. Cyclin induction was less correlated to Aβ expression but still highly significant (Table 1). Control experiments with Dox alone or Adv-GFP +Dox showed no evidence of cell cycle protein expression (Supplementary Material, Fig. S2B).

Figure 2.

Cultured myotubes bearing intracellular β-amyloid evidence cell cycle initiation. Immunofluorescence photomicrographs of Adv Aβ-infected myotubes. Following a 24 h treatments with Adv-TRE-Aβ and Adv Tet-On, Aβ expression was induced by treatment of doxycycline (Dox, 2 µg/ml) for 24 h. The Dox-untreated myotubes showed no positive staining with the cell cycle markers (left 3 panels), whereas nuclei in Dox-treated myotubes were positive for antibodies to PCNA (S-phase marker; red) and Ki-67 (cellular proliferation marker; green), cyclin D1 (G1 to S phase marker; red) and cyclin E (G1 to S phase marker; red) (right 3 panel columns). Arrows indicate myonuclei in atrophic cells labeled with cell cycle markers indicated along figure left. Absence of IgG uptake serves as negative control. Scale bar: 50 μm.

Table 1.

Statistical quantification of Supplementary Material, Fig. S2

	PCNA	Ki-67	Cyclin D	Cyclin E	Aβ-6E10	Aβ-R1282	IgG
Mean # of myotubes: 1 positive nucleus	40 (1)	34 (0)	31 (0)	42 (0)	$41 (1)	$89	0
Mean # of myotubes: ≥2 positive nuclei	14 (0)	18 (0)	11 (0)	26 (0)			0
Mean # of myotubes: ≥4 fragmented or pyknotic nuclei	18 (0)	18 (0)	25 (1)	16 (0)	17 (3)	22	0
Density of myotubes: myotubes/mm2	20 (29)	21 (30)	22 (26)	23 (25)	22 (33)	21
Total # of myotubes	1269 (4615)	1107 (3161)	3445 (1612)	3510 (1560)	2320 (5200)	2233
Ratio of myotubes positive for Aβ and cell cycle marker to myotubes positive for either Aβ or cell cycle marker only	5.00	1.93	0.46	1.05

C2C12 myotubes were cultured on chamber slides coinfected with Adv-TRE-Aβ and Adv Tet-On and expression induced with 24 h doxycycline. No added doxycycline served as control. Immunofluorescence co-staining with anti-Ki-67 and Aβ (6E10 or R1282), anti-PCNA and Aβ, anti-cyclin D1 and Aβ and anti-cyclin E and Aβ was carried out and detected using antibodies of monoclonal goat anti-mouse cy2 or cy3 or polyclonal goat anti-rabbit cy2 or cy3. Myonuclei were identified by DAPI and the number of multinucleated myotubes containing 1 and ≥2 nuclei bearing Ki-67, PCNA, cyclin D1 or E decoration were counted over five random, 40× fields in each of the four development conditions. Means and SEM (not shown) were calculated over triplicate (n = 3) experiments. In each 8 × 8 mm well, mean counts were also made of Aβ bearing myotubes and of those expressing both Aβ and cyclin marker. The correlation of Aβ expression with cell cycle entry markers in the same myotube was tested using 2 × 2 tables and χ² method. The ratio of Aβ and cycle marker co-occurrence to the sum number of their dissociation is given at bottom. In each of the four marker and Aβ expression tests listed at the top, the association was significant to P < 0.0001. χ²-values were highest for PCNA/Aβ and in descending order; Ki-67/Aβ, cyclin E/Aβ and cyclin D1/Aβ (960.4, 834.9, 675.8, 298.2, respectively, df = 1). $: mean # of myotubes positive for cytosolic Aβ expression by antibody as listed; ( ): mean # of myotubes, density or total # in the absence of doxycycline (control).

New DNA synthesis in myotubes overexpressing Aβ

We further investigated the involvement of Aβ in new DNA synthesis with BrdU incorporation assays. BrdU staining as well as ELISA-based incorporation experiments were performed in cultured myotubes coinfected with Adv-TRE-Aβ and Adv Tet-On. Immunofluorescence data revealed more nuclei decorated with BrdU antibody in Aβ-overexpressed myotubes than in the control (Fig. 3A). These results were confirmed and quantified using a BrdU chemiluminescence assay (400–650 nm) in a multi-well format (n = 6, each group). New DNA synthesis over a 2 h labeling period was doubled in Aβ-overexpressed myotubes compared with control samples (either doxycycline alone without virus or non-induced myotubes; Fig. 3B). New DNA synthesis was also measured at various time points after Aβ induction (n = 3 experiments, each in triplicate). BrdU application commenced at 24 h following initial doxycycline treatment, and continued until harvest at time points from 0 to 36 h. BrdU incorporation was significantly increased in Aβ-overexpressed myotubes compared with non-induced myotubes at both 24 and 36 h time points after BrdU addition (Fig. 3C, left graph). Control experiments included treatment with cytosine arabinoside (AraC, 50 µm), resulting cell cycle arrest in normal dividing myoblasts (Fig. 3C, right graph), which also blocked Aβ-induced new DNA synthesis up to and beyond 24 h after BrdU treatment (Fig. 3C, middle graph).

Figure 3.

BrdU incorporation and time course in Aβ42-expressing myotubes. New DNA synthesis in myotubes shown by immunofluorescence in (A) and quantified by ELISA in (B). Cell cultures were treated as in Figure 2. BrdU (10 µM) added to the medium for 2 h before fixation and ELISA (n = 6). Scale bar: 30 μm. (C) New DNA synthesis rate is significantly increased in Aβ-induced cultured myotubes at 24 and 36 h after Aβ induction (left). Treatment with AraC (50 µm) blocked Aβ-induced new DNA synthesis beginning at 4 h after Aβ induction (meddle, note different scale from left). Control elimination of cell division by AraC in dividing myoblasts shown to the right. All data presented are mean ± SEM. *P < 0.05 versus Adv Aβ-Dox (ANOVA).

Increased expression of cyclin E and PCNA in EAM-like mice

The number of nuclei engaged in the cell cycle (Ki-67 staining) was increased in human PM as well as in IBM cases (Fig. 1A and B), indicating that amyloid and/or cell inflammation can stimulate reentry. Therefore, we examined the levels of other cell cycle markers in a previously characterized (see Materials and Methods) murine C-protein-induced myositis model of PM similar to experimental allergic myositis (EAM). Muscle proteins were extracted from EAM (n = 7) and control (n = 6) mice and analyzed by western blots. The levels of cyclin E and PCNA were significantly increased in the EAM samples compared with the littermate control group (Fig. 4A and B). There were no significant differences in the levels of cyclin D1 between the groups (Fig. 4A and B). The level of CD4, another T lymphocyte marker, was also significantly increased in the EAM samples compared with control, indicating a cellular inflammatory response (Fig. 4A and B). As similar to human cases (inflammatory infiltration % versus Ki-67), there was no positive correlation between cell cycle markers and CD4 expression in EAM samples (PCNA versus CD4, R² = −0.005, P = 0.99; cyclin E versus CD4, R² = 0.61, P = 0.14; cyclin D1 versus CD4, R² = −0.25, P = 0.59). The trend in cyclin E induction with increasing inflammation is nevertheless interesting (see below). We then characterized the cyclin E-positive cells immunocytochemically using antibodies to CD8, β-sarcoglycan and Pax7. The majority were mature myofibers. Thus, >90% of cyclin E-positive nuclei showed no co-staining with anti-CD8 in EAM samples (Supplementary Material, Fig. S3A), whereas a small number of cyclin E-positive nuclei double-stained with CD8 antibody in areas of inflammatory infiltration (Supplementary Material, Fig. S3A). Similar results were observed with Pax7 antibody (Supplementary Material, Fig. S3A), excluding a satellite origin for the cyclin E signals. Cyclin E-positive nuclei were accordingly located in the subsarcolemma (Supplementary Material, Fig. S3A). CD4/8 immunofluorescence and H&E stain confirmed the induction of cellular inflammation in EAM mice (Supplementary Material, Fig. S3B). Next, treatment of myotubes in culture with a combination of proinflammatory cytokines, tumor necrosis factor (TNF)-α and interleukin (IL)-1β (100 and 50 ng/ml, respectively), for 72 h increased the expression level of cyclin D1 in myotubes to a moderate degree (1.0 ± 0.14 control (n = 3) versus 1.86 ± 0.16 TNF-α/IL-1β (n = 4), P = 0.01; Figure 4C). The levels of cyclin E and PCNA were unchanged (Fig. 4C). As control, we show that TNF-α and IL-1β increased the phosphorylation of nuclear factor (NF)-κB, confirming the induction of an appropriate intracellular inflammatory response to the treatment (Fig. 4C). The combination of TNF-α and interferon (IFN)-γ had no effect on expression levels of either cyclins E and D1 or PCNA in myotubes (not shown). Thus, other T cellular immune responses, cytokines contributing, are primary associated with stimulation of cell cycle in muscle.

Figure 4.

An in vivo model of myositis (EAM) induces cell cycle entry markers independent of β-amyloid. Normal mice immunized with C muscle protein. (A) Muscle proteins were extracted and analyzed by western blot. Representative immunoblot shows induction of cyclin E and PCNA. CD4, a helper T cell marker of cellular inflammation, was also induced. (B) Densitometric quantification of western blots. (C) An in vitro model of passive inflammation in myotubes employing exogenous cytokines evidenced modest cyclin D1induction. TNF-α and IL-1β (100 and 50 ng/ml, respectively) were co-added to myotubes for 72 h. Cyclin E and PCNA were unchanged. Co-additions of TNF-α and IFN-γ had no effect (not shown). All data presented are mean ± SEM. *P < 0.01 versus control (t-test).

Increased expression of cell cycle markers, caspases and HSPs in human IBM and PM inflammatory myopathies

To further quantify the phenomenon of postmitotic cell cycle entry in human muscle biopsy samples from IBM (n = 8), PM (n = 8) and control (n = 8) patients (for clinical details, see Supplementary Material, Tables S1 and S2), the levels of cyclins E and D1 and PCNA were examined by western blots. Since HSPs (34,35) and caspases (36) are important regulators of cell cycle control, the levels of chaperones (HSPs 27, 60, 70 and 90) and apoptotic modulators (caspases −3 and −9) were examined in the same samples. Levels of cyclins E and D1 and PCNA were significantly increased in both IBM and PM samples compared with the control, consistent with Figure 1 results (Fig. 5A and B). The level of CD4 was also significantly increased in both IBM and PM samples compared with control (Fig. 5A and B). As with the EAM model, there was no positive correlation between cell cycle induction (PCNA or cyclin D1) and inflammation (CD4 expression) in IBM and PM samples (PCNA versus CD4, R² = 0.20, P = 0.58; cyclin D1 versus CD4, R² = −0.15, P = 0.68). However, we observed a positive correlation between levels of cyclin E and CD4 expression (cyclin E versus CD4, R² = 0.71, P = 0.03). CD8 and markers of cycle reentry did not colocalize (Supplementary Material, Fig. S1B), similarly, nor did CD4 (not shown). Thus, there remains a causal possibility that the level of cellular inflammation is somehow linked with the degree to which at least this cell cycle marker is stimulated.

Figure 5.

Coordinate expression of cell cycle markers with caspase activation and HSP induction in human IBM and PM. Muscle proteins were extracted from human IBM, PM and normal control muscle (n = 8 each group) and analyzed by western blot. (A) Representative western blots of human muscle proteins and (B) densitometric quantification. The levels of cyclins E and D1 and PCNA are increased in inflammatory myositis, consistent with Figure 1A result. CD4 showed induction in IBM and PM. In the same samples, levels of HSP60, 70 and 90 are also increased. HSP27 is unchanged. Evidence for procaspase-3 induction and caspases-3 and -9 cleavage is shown (B, middle and low charts). Changes in procaspase-9 levels were not observed. For comparison, proteins extracted from human AD brains were analyzed by western. The levels of cyclins E and D1 and HSP70 were similarly increased in AD compared with the control (25 µg total protein). All data presented are mean ± SEM. *P < 0.05 versus control (ANOVA). ^†P < 0.05 versus control (t-test). Twenty micrograms of protein loaded per lane.

As negative control, we examined cyclin E and PCNA protein levels in a diverse set of age-matched non-inflammatory myopathy and dystrophy cases (NIM). In 10 such samples, no increase in cycle markers was observed above the same normal control set (result not shown, cases are given in Supplementary Material, Table S2).

The levels of procaspase-3 and cleaved caspases-3 and -9 were significantly increased in IBM and PM samples compared with the control (Fig. 5A and B). The level of HSP70 was significantly increased in both IBM and PM samples, whereas elevated levels of HSPs 60 and 90 reached significance only in the PM samples compared with the control (Fig. 5A and B; t-test). There were no significant differences in the levels of HSP27 and procaspase-9 among the groups (Fig. 5A and B). Straight westerns from these IBM lysates were characteristically negative for anti-Aβ development, due likely to an insensitivity to detect low levels of intracellular amyloid affecting a relatively small number of myofibers. For a positive control, we examined the levels of cyclins and HSP70 in brains of Braak Stages V–VI AD patients by western blots. As expected, levels of cyclins E and D1 and HSP70 were increased in AD compared with the control (Fig. 5C), consistent with published data.

Interaction of cyclins E and D1 with HSP70 in human muscle

In order to investigate the possibility that the increased cyclin levels are stabilized by HSPs, the interaction between the cyclins and HSPs or caspases was examined. Human muscle protein extracts from IBM (n = 8), PM (n = 8) and control (n = 8) patients were immunoprecipitated with cyclins E or D1 antibody. The immune complexes were denatured and the levels of co-precipitating HSPs 27, 60, 70 and 90 were measured by western blot. We found that HSP70 protein physically interacted with cyclins E or D1 across all the three subject groups (Fig. 6A) and in proportion to their individual levels. The levels of cyclins E and D1 and complexed HSP70 were higher in IBM and PM than in the control (Fig. 6A) as expected from straight westerns in Figure 5. However, the binding ratio of HSP70 to the induced cyclins E or D1 was not different among the three groups (Fig. 6B). Reverse IP showed the expected pull-down of cyclins with HSP (not shown). HSPs 27, 60 and 90 were not detected in immune precipitates with cyclins in any group (not shown). As control, AD brain samples were immunoprecipitated with cyclins E or D1 antibody and the level of HSP70 was measured by western blots. Similar to the muscle studies, HSP70 is shown to interact with cyclins E or D1 in both AD and control brains (Fig. 6C) without major changes in ratio (quantitation not shown). Caspases −3 and −9 were not detected in complex with any of the cyclin proteins.

Figure 6.

Cyclin induction in IBM and PM is proportional to physical interaction with HSP chaperones. (A) Interaction of cyclins E and D1 with HSP70 in human muscle. Human muscle proteins extracted from IBM, PM and normal control patients (n = 8 in total each group) were immunoprecipitated with cyclins E or D1 rabbit polyclonal antibody and detected by western blot with mouse monoclonal antibodies as indicated. (B) Densitometric quantification of western blot (WB). HSP70 coprecipitated with cyclins E or D1 in all the three groups. Consistent with Figure 5, the binding ratio of HSP70 to cyclins E or D1 did not differ. (C) Proteins extracted from human AD brains (250 µg total brain lysate) were like-wise analyzed by immunoprecipitation (IP)-WB to show binding of cyclins E or D1 to HSP70. A reverse IP shows like pull-down of cyclins E or D1 with anti-HSP70 (results and binding ratio not shown). All data presented are mean ± SEM.

Aβ expression decreased cdk inhibitor p21 levels and viability in myotubes

The Cdk inhibitor, p21 Cip1/WAF1 and retinoblastoma tumor suppressor protein (Rb) are known to play an important role in cell cycle regulation, through association with cyclins D and E and their CDK families cdk 4/6 and cdk2 (37). Moreover, p21 is crucial in maintaining differentiated myotube in a state of exit from the cell cycle (38,39). We tested if levels of p21 and phosphorylated Rb change during Aβ-induced cycle reentry in amyloid-bearing myotubes. Myotubes were infected with Adv-TRE-Aβ and Adv Tet-On for 24 h and then Aβ expression was induced by treatment with doxycycline for another 24 h. Levels of total p21 decreased in Aβ-expressing myotubes (Dox) compared with the control (No Dox) (P < 0.05; Fig. 7A and B). Control westerns with Dox alone or Adv-GFP +Dox show no changes in p21 or phosphorylated Rbs (not shown). Consistent with our models, p21 levels were decreased in IBM and PM without difference compared with normal control (n = 5 each group, P < 0.05; result not shown). There were no differences in the levels of phosphorylated and total Rb between the same two conditions (±Dox) in myotubes (Fig. 7A and B). Phosphorylated Rb levels were too variable between control, IBM and PM patients to further quantify. In order to examine if p21 can rescue Aβ toxicity in myotubes (by inhibiting reentry), p21 was overexpressed by coviral infection with Herpes simplex virus (HSV)-p21 and compared with control vector (encoding GFP). Whereas Aβ expression over 48 h (Dox) decreased myotube viability (WST-1 assay), overexpression of p21 did not rescue the toxic effect (Fig. 7C). Viral expressions of Aβ and p21 were confirmed (Fig. 7C, lower panel). As control, we confirmed that overexpression of p21 appropriately inhibited DNA replication in dividing myoblasts. HSV-p21 infected myoblasts showed far fewer BrdU-positive nuclei than HSV-control infected myoblasts (Fig. 7D). Moreover, overexpression of p21 in myotubes resulted in a moderate reduction in phosphorylated Rb (S780 and S807/811), as expected (not shown). Expression was confirmed by immunocytochemistry (Fig. 7D, lower panel). Additionally, we tested if flavopiridol, a general cdk inhibitor (40), rescues Aβ toxicity in myotubes. Similar to the p21result, flavopiridol (75 nm) did not rescue cells from the toxic effect of Aβ (data not shown).

Figure 7.

Aβ expression and cell cycle regulators Rb and p21. (A) Myotubes were infected with Adv TRE-Aβ and Tet-On for 24 h and induced to express Aβ after adding doxycycline (Dox, 2 µg/ml) for another 24 h. Western of whole cell extracts (A) and densitometric quantification (B) demonstrate a decrease in p21 corresponding to Aβ overexpression. There were no differences in the levels of phosphorylated or total Rb between the groups. (C) p21 overexpression (HSV-p21) did not rescue the decrease in myotube viability attributable to Aβ induction (lane 4 versus lane 3). Doxycycline alone control does not affect WST reduction (not shown). Western blot below confirms expression of Aβ and/or p21. (D) Control experiments for p21 activity. The stimulation of DNA synthesis by Aβ (lane 3) is reversed by co-expression of p21 (lane 4) in myotubes. Quantification is of BrdU-positive nuclei from 6 random 40× fields. Representative immunofluorescence photomicrographs below demonstrate p21 expression in myoblasts. All data presented are mean ± SEM. ***P < 0.001 versus no Dox (t-test). ^†P < 0.05, ^†††P < 0.001 versus no Dox and no HSV-p21or HSV-GFP control (ANOVA). Scale bar: 30 μm.

DISCUSSION

Under various pathological conditions, including AD, neurons can be induced to initiate DNA replication. While not resulting in mitosis (41), cell cycle reentry is likely to initiate apoptosis (3,42,43). However, cell cycle activation is also involved in a reparative response to oxidative DNA damage (18). Until the current study, no documentation or experimentation has tested whether cell cycle reentry occurs in human myositis or in cell or animal models PM and IBM.

In this report, we demonstrate aberrant cell cycle reentry in IBM and obtained confirmatory evidence that AD brain is similarly affected. We observed increased expression of cell cycle markers Ki-67, PCNA and cyclins E and D1, and provide the first documentation of cyclin interaction with chaperone HSP70 in both human IBM muscle and AD brain. These findings suggest that the two conditions may share biological triggers for the misregulation of cell cycle proteins. Experimental evidence is presented that new DNA synthesis occurs in myotubes expressing Aβ accompanied by the induction of the same cell cycle markers. Mechanistically, intracellular Aβ expression in myotubes decreased levels of p21 (Cip1/WAF1), a well-known cdk inhibitor. This may partially account for the Aβ-dependent contribution to cycle reentry, as demonstrated through the experimental restoration of p21 levels. Interestingly, increased expression of cell cycle regulatory proteins was also observed by us in a related inflammatory disorder of muscle, PM. We further show, using an EAM model of PM, that autoimmune cellular activation can also drive cycle progression, independent of Aβ expression. Our finding of cycle reentry in IBM and PM involved mature muscle and did not reflect satellite cell activation or division. The same conclusion that SCs are not activated in IBM and PM was noted by another group (30). Hematopoietic/endothelial progenitor cells were reported upregulated in inflammatory myopathies, but not correlated with Ki-67 (44).

The exact mechanism through which Aβ accumulation is toxic to skeletal muscle is unknown. In this work, we looked for evidence of cell cycle reentry and finding such, analyzed several possibilities to explain the induction and how they might lead to muscle cell attrition. As described in greater detail below, these mechanisms included: (i) chaperone mediated stabilization of cyclins, (ii) loss of Cdk inhibitory control, (iii) involvement of caspases and (iv) interaction with mediators of inflammation.

Molecular chaperones in general maintain intracellular protein homeostasis in cooperation with the ubiquitin–proteasome degradation system. HSPs are also induced when the cell cycle is activated in neurons and in turn, various HSPs regulate the G1 phase (45,46). For instance, the chaperone HSP70 interacts with cyclins E and D1 during cell cycle progression (47–49). We show that induction of molecular chaperones is coordinated with the upregulation of cyclins in IBM and PM. The interaction of cyclins E or D1 with HSP70 is proportionally increased in IBM and PM muscle and in AD brain. Similar to our results, De Paepe et al. (80) showed elevated levels of HSPs 70 and 90 in human IBM and PM biopsy specimens, presumably in a stress-protective role. Taken together, the interaction between HSP70 and cyclins E and D1 may have significance during cell cycle progression in the inflammatory myopathies, possibly regulating cyclin maturation or degradation.

Another potential mechanism for Aβ-stimulated DNA synthesis investigated here is the dysregulation of the Rb and/or Cdk inhibitor p21 proteins. Either an inhibitory phosphorylation of the former or decrease in levels of the latter can be expected to stimulate the cell cycle. p21-mediated cell cycle attenuation is significantly associated with the maintenance of Rb in its hypophosphorylated or active form (37,50–52). p21 directly inhibits the activity of cyclin E-Cdk2 and cyclin D-Cdk4/6 complexes, thereby negatively regulating cell cycle progression in G1, S and G2 phases (53–55). p21 also binds to PCNA, inhibits its activity and blocks DNA replication (56). In skeletal and vascular smooth muscle, Rb is a downstream effector of p21 (57–59), where both maintain terminally differentiated muscle in a state of arrested DNA synthesis (38,60,61). Overexpression of p21 inhibits proliferation of vascular smooth muscle cells in vitro (62,63) while suppression of p21 by siRNA initiates DNA synthesis and cyclin D-Cdk4/6-dependent cell cycle reentry in myotubes (64). In similar studies, inactivation or deletion of p21 and Rb led to renewed DNA replication and restoration of cyclin E-Cdk2 activity in terminally differentiated C2 myotubes (57,65).

Alongside the aberrant expression of cell cycle regulatory proteins, we detected decreased p21 levels in Aβ-expressing myotubes. Unexpectedly, lowered levels of p21 in Aβ-expressing myotubes were not associated with changes in the levels of phosphorylated Rb. The decrease in p21 levels may have several causes not tested here, including diminished transcription (involving p53, myoD), proteasome degradation (66) and/or caspase cleavage. For instance, conversion from G1 or G2/M arrest to apoptosis in the course of genotoxic stress is associated with p21 cleavage by caspase-3 (67,68). In our hands, p21 co-expression was sufficient to reverse induced BrdU incorporation following Aβ expression, attesting to its effectiveness in cell cycle control under these conditions. Regulation by p21 may be mediated by its action to inhibit PCNA (69) rather than through Rb. Importantly, our experiments found that p21 overexpression did not overcome the Aβ-mediated decrease in myotube viability. p21 and phosphorylated Rb levels did not distinguish between IBM and PM patients. These results suggest that cell cycle reentry in the context of amyloid toxicity is not pathogenic and possibly epiphenomenal. Thus, other derangements may predominate in mediating amyloid toxicity in muscle including perturbations in insulin signaling (26,70), Ca2+ homeostasis (71,72), mitochondrial function (73) or proteasome/ER stress (74,75).

The partial activation of initiator/effector apoptotic pathways is suggested by the association of caspase changes with aberrant G1/S transition. In the IBM and PM samples, procaspase-3 and cleaved caspases-3 and -9 were elevated (Fig. 5). Interestingly, one study has shown increased expression of caspase-3 and -9 in hereditary inclusion body myopathy, a condition with amyloid deposits and downregulation of Akt signaling but relatively lacking inflammation (76). Independent of proteinopathy, inflammatory muscle death from IFN-γ application involves the ER-resident caspase-12 and Bax/Bcl2 activations (77,78). While caspase activation may regulate the cell cycle (e.g. p21 cleavage) or result from cyclin induction, the non-existence or rarity of apoptosis in IBM does not suggest a major pathological role.

Finally, we examined the contributory role of inflammation in muscle cell cycle reentry. The two major inflammatory myositis disorders, IBM and PM, share T-cell-mediated cytotoxicity and the increased production of proinflammatory cytokines: TNF-α, IFN-γ and IL-1 (79–82). Their respective mouse models also provide evidence for inflammation (83–85). Both PM and IBM sample sets used here had comparable levels of moderate inflammation and reduction in muscle fiber size. It is therefore not surprising that in both conditions, expression of nuclear Ki-67, PCNA and cyclins E and D1 proteins were upregulated. Moreover, there were no differences between IBM and PM in caspase activation or HSP induction. Importantly, we also found no changes in cyclin or cell cycle marker levels in a diverse set of non-inflammatory, non-amyloid-bearing myopathy and dystrophy conditions, similar in comparison to a separate normal control muscle tissue set. For this reason, we separately investigated models of amyloid accumulation and cell-mediated inflammation for evidence of cyclin induction. Using an EAM model of PM (85,86), we found that autoimmune T-cell inflammation was also sufficient to induce cell cycle regulatory proteins. However, cycle reentry expression did not correlate with the extent of inflammation, nor did Ki-67 colocalize with the inflammatory cells. These findings argue against a simple direct relationship. Moreover, like the expression of β-amyloid, the combined application of cytokines TNF-α and IL-1β also induced cell cycle protein expression in myotubes, albeit only partially (cyclin D1). We found the combined cytokines TNF-α and IFN-γ (87) had no effect on cell cycle protein expression in myotubes. The importance of IL-1β is consistent with in vitro and in vivo results (83,84). In addition, it is likely that other T-cell-mediated immune factors indirectly drive the skeletal muscle cell into DNA replication, as noted in vascular smooth muscle cells (88,89). Therefore, we hypothesize that β-amyloid-mediated injury and cell-mediated inflammation are independently driving cell cycle reentry in IBM.

In the present work, we focused on G1/S events. Future work will establish if a block occurs specifically at G2/M (90). Another area of future research is to explore the role of Cdk5 and cycle control in IBM. Furthermore, the available cell cycle studies in AD still leave for question whether cell cycle reentry is pathogenic or epiphenomenal. The restoration of cell cycle withdrawal in AD animal and cellular models has not to our knowledge been attempted. Here, we overexpressed p21 and applied the cdk inhibitor flavopiridol in a skeletal muscle cell model of IBM, which did not rescue Aβ-induced cell death. In contrast, flavopiridol is found to be cytoprotective in stroke and Parkinson's disease models (91–93) and similarly, p21 in a trauma model (94). Interestingly, in one AD-like mouse, non-steroidal anti-inflammatory drug treatments prevented new neuronal cell cycle entry events but do not reverse existing ones or amyloid pathology (95). In our preliminary testing, aberrant cell cycle reentry in skeletal muscle cells, at least from amyloid deposition, does not present a strong therapeutic target. However, the critical experiments in other myositis and AD models should address this question before the role of cycle reentry in neurodegenerative diseases can be fully defined.

MATERIALS AND METHODS

Materials

The following antibodies were purchased from the cited manufacturers; anti- cyclins E and D1, PCNA, BrdU, Ki-67, Pax7, p21, HSPs 27 and 60 (Santa Cruz Biotechnology, Santa Cruz, CA), anti-caspases-3 and -9, phospho-NF-κB p65 (pNF-κB p65, Ser536), phosphorylated (Ser780, Ser795 and Ser870/811) and total Rb (Cell Signaling Technology, Danvers, MA, USA), anti-HSPs 70 and 90 (Stressgen, San Diego, CA, USA), anti-β-amyloid (6E10; Signet Pathology Systems, Dedham, MA, USA), anti-CD4 and -CD8 (Abcam, Cambridge, MA, USA), anti-CD45 (Cell Marque, Rocklin, CA, USA), anti-β-sarcoglycan (Vector Labs, Burlingame, CA, USA) and anti-actin (Sigma, St. Louis, MO, USA). Recombinant human TNF-α, IFN-γ and IL-1β (Invitrogen, Carlsbad, CA, USA) was dissolved in dH₂O and stored at −20°C until used. IBM, PM and normal (no diagnostic abnormality) control skeletal muscle biopsy specimens were obtained from the Pathology Departments of Tufts Medical Center and CSEMC (Boston, MA, USA; total n = 8, 8, 11, respectively). A second control set consisted of cases of NIM or dystrophy (total n = 10). All biopsy specimens were obtained before any steroid or immunomodulating treatment was initiated. Diagnostic (96) and clinicopathologically (188th ENMC 2011 workshop) defined IBM criteria, patient characteristics (age, gender, degree of vacuolization/inflammation, storage time) and sample handling are identical as recently published (see Supplementary Material, Tables S1 and S2) (26). Human control and AD brain samples (Frontal cortex, Braak Stages V–VI) were kindly provided by the Harvard Brain Tissue Resource Center (McLean Hospital; Harvard Medical School; Belmont, MA, USA). Diagnostic and clinicopathologic criteria for PM were followed as defined in (97,98).

Viral-mediated expression of β-amyloid1–42 in C2C12 myotubes

Mouse C2C12 myoblasts were grown to 70–80% confluence in Dulbecco's modified Eagle's medium (Invitrogen) containing 20% fetal bovine serum (Invitrogen), penicillin/streptomycin, 1 mm sodium pyruvate and 2 mm l-glutamine and maintained at 37°C in 5.5% CO₂. After 24 h, cells were changed to differentiation media containing 2% horse serum for another 24 h. At this time point, the mixed population of myotubes and myoblasts were co-infected with Adv Tet-On and Adv TRE-Aβ42 1:5 ratio (100 multiplicity of infection) for 24–36 h in differentiation media. Aβ expression was induced by the addition of doxycycline at 2 µg/ml. HSV encoding-p21 and GFP (control) and FLAG tagged-flt were designed and used exactly as reported in Querfurth et al. (99). FLAG-flt is a 44 amino acid fusion in peptide corresponding to the VEGF-R transmembrane domain serving as another control. Synthesis and preparation of stocks were performed in the MIT viral core laboratory (Dr Rachael Neve Ph.D., director).

Experimental allergic myositis, a model of PM

To test the regulation of cell cycle proteins under inflammatory myositis conditions independent of amyloid, we employed an EAM-like model of PM (85,86). C57BL/6 mice with subacute myositis were generated by skeletal muscle fast-type C protein immunization and described in our previous study (26). Animals used in this study were 8–10 weeks age, 2–3 weeks after injection. This time point is associated with a robust inflammatory response. The endomysial infiltrates of inflammatory lymphocytes in EAM mouse muscle were demonstrated in our previous study (26).

Immunofluorescence microscopy of myotubes, myoblasts and human and mouse skeletal muscle

Myotubes coinfected with Adv-TRE-Aβ and Adv Tet-On (with or without added doxycycline) grown on chamber slides or 10 µm cryostat sections of frozen human and mouse skeletal muscle biopsy samples mounted on glass slides were fixed with methanol at −20°C or 4% paraformaldehyde at room temperature for 10 min followed by a PBS wash and then blocked for 1 h with PBS/BSA containing 10% goat serum. Slides were incubated overnight at 4°C in primary antibodies against cell proliferation markers Ki-67 (1:150), PCNA (1:150), cyclin E (1:150), cyclin D1 (1:150), Pax7 (1:50) and CD8 (1:200) and against β-amyloid (1:150, 6E10). To test the effect of p21 overexpression on DNA replication, myoblasts were infected with HSV expressing p21 or control HSV for 24 h. The myoblasts were treated with bromodeoxyuridine (BrdU; 10 µm) for the last 2 h of the 24 h HSV exposure period and then fixed in 4% paraformaldehyde at room temperature for 10 min. In order to denature DNA, the myoblasts were incubated in 2 m HCl for 20 min at room temperature. After blocking, primary antibodies (p21, 1:200; BrdU, 1:200) were incubated overnight at 4°C. Thereafter, sections were washed and incubated in fluorescent-labeled secondary antibody (1:200; Jackson ImmunoResearch, West Grove, PA, USA) for 1 h. The fluorescent signals were visualized under a Nikon TE200 epifluorescence microscope. Immunofluorescent nuclei were confirmed by DAPI stain. Ten random fields (two in each quadrant and two in the center) at 40× were counted and the mean per field was calculated for each patient, then averaged again. Satellite cells were operationally defined as small myogenic cells with a single large (≥8 µm), anti-Pax7 stained nucleus alongside muscle fibers, between the fiber's sarcolemma and the basal lamina (32). Nuclei and sarcolemma were confirmed with bisbenzamide and anti-β-sarcoglycan, respectively. Total SCs were counted per section from n = 2–3 each control, IBM and PM cases and normalized to section area determined by ImageJ software (NIH).

DNA replication

New DNA synthesis was assayed by standard BrdU incorporation. Quiescent myotubes were coinfected with Adv Tet-On and Adv TRE-Aβ42, as described above, for 24–36 h in differentiation media followed by the addition of 2 µg/ml doxycycline for another 24 h to induce Aβ production. At 0–36 h time points after doxycycline addition, control and Aβ-bearing myotubes were incubated with BrdU (10 µm) for 2 h followed by fixation, acid denaturation and incubation with the BrdU antibody (1:200). Incorporated BrdU was detected by ELISA (400–650 nm) according to the manufacturer's instructions (Roche, Mannheim, Germany) and a Perkin-Elmer plate reader.

Protein analysis

Western blots

Frozen human and mouse muscle tissue was crushed using pestle and mortar. The muscle fragments were lysed in Nonidet P (NP)-40 lysis buffer (20 mm Tris–HCl, pH 7.4, 150 mm NaCl, 10% glycerol, 2 mm EDTA and 1% NP-40) containing protease inhibitor cocktail (Roche, Mannheim, Germany) at 4°C for 30 min, centrifuged at 12 000×g for 15 min, and the supernatant transferred into new tubes. The protein concentration was determined using a Bio-Rad Protein Assay kit (Bio-Rad, Richmond, CA, USA). Each protein sample (40 µg) was heated at 95°C for 10 min in Laemmli sample buffer, separated on 10% SDS–polyacrylamide gels, and then electrotransferred onto polyvinylidene-difluoride (PVDF) membranes. The membranes were blocked in 5% non-fat dry milk in Tris-buffered saline (20 mm Tris, pH 7.6, 0.8% NaCl) containing 0.1% Tween 20 (TBST) for 1 h and then hybridized with primary antibodies in block at 4°C overnight. After incubation with primary antibodies, membranes were washed with TBST, incubated with HRP-conjugated secondary antibodies (1:4000 dilution; Cell Signaling Technology) in TBST at room temperature for 1 h, and then washed with TBST again. The signal was detected using enhanced chemiluminescence reagents and film (GE Healthcare, Piscataway, NJ, USA). Band density was measured from the film using ImageJ software (NIH). Background signal changes were subtracted and each value was normalized to its control within any given experiment.

Immunoprecipitations

Homogenates of muscle (500 µg protein per sample) were incubated at 4°C overnight with 3 µg of primary antibodies (1:50, cyclin E or cyclin D1). Protein A/G-Agarose (Roche, Mannheim, Germany) was added and an additional incubation was performed at 4°C for 3 h. Immunoprecipitates were harvested by centrifugation at 12 000×g for 1 min and washed several times at 4°C in NP-40 lysis buffer containing protease inhibitor cocktail (Roche Biochemical). After the final wash, immunoprecipitates were denatured in Laemmli sample buffer by heating at 95°C for 5 min and then centrifuged at 12 000×g for 5 min. The supernatant was used for western blot analysis as described above.

Cell viability

Myotubes were infected with Adv Tet-On and Adv TRE-Aβ42 in differentiation medium in a 24-well plate. Twenty-four hours later, Aβ expression was induced by the addition of doxycycline at 2 µg/ml for a further 48 h. To overexpress p21, the myotubes were infected with HSV-p21 (or HSV-flt-control) for the last 24 h of the Aβ induction period. 4-[3-(4-Iodophenyl)-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate (WST-1) solution (Roche, Mannheim, Germany) was added to the myotubes in each well for the last 4 h of the experiment period. Reduced WST was detected using absorbance at 490 nm by a Vmax microplate reader (Molecular Device, Sunnyvale, CA, USA).

Statistical analysis

Statistical significance across treatment groups was detected by t-test and/or one-way ANOVA with Neuman–Keuls post hoc tests (Prism, GraphPad Software). The χ²-test was used to compare the number of immunofluorescent-labeled myonuclei with Aβ and (or) cell cycle markers in Table 1. All data are presented as the mean ± standard error of the mean. Measurements were conducted in triplicate over n-independent experiments, unless otherwise stated. Inflammation was quantified as % section area covered by extramysial nuclei and fiber diameters were measured using ImageJ software (NIH) and distribution plotted using Excell.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at HMG online.

FUNDING

This work was supported by NIH 14373 to H.W.Q., the Joseph and Eileen Blake Fund, The Bennett Foundation, NIH AGO34972 to K.W., and Institutional Resources.