Low-Frequency Electrical Stimulation Attenuates Muscle Atrophy in CKD—A Potential Treatment Strategy

Abstract

Effective therapeutic strategies to treat CKD-induced muscle atrophy are urgently needed. Low-frequency electrical stimulation (LFES) may be effective in preventing muscle atrophy, because LFES is an acupuncture technique that mimics resistance exercise by inducing muscle contraction. To test this hypothesis, we treated 5/6-nephrectomized mice (CKD mice) and control mice with LFES for 15 days. LFES prevented soleus and extensor digitorum longus muscle weight loss and loss of hind-limb muscle grip in CKD mice. LFES countered the CKD-induced decline in the IGF-1 signaling pathway and led to increases in markers of protein synthesis and myogenesis and improvement in muscle protein metabolism. In control mice, we observed an acute response phase immediately after LFES, during which the expression of inflammatory cytokines (IFN-γ and IL-6) increased. Expression of the M1 macrophage marker IL-1β also increased acutely, but expression of the M2 marker arginase-1 increased 2 days after initiation of LFES, paralleling the change in IGF-1. In muscle cross-sections of LFES-treated mice, arginase-1 colocalized with IGF-1. Additionally, expression of microRNA-1 and -206, which inhibits IGF-1 translation, decreased in the acute response phase after LFES and increased at a later phase. We conclude that LFES ameliorates CKD-induced skeletal muscle atrophy by upregulation of the IGF-1 signaling pathway, which improves protein metabolism and promotes myogenesis. The upregulation of IGF-1 may be mediated by decreased expression of microRNA-1 and -206 and/or activation of M2 macrophages.

Muscle atrophy is a major complication of CKD associated with excess morbidity and mortality.^1,2 Many investigators have studied muscle wasting to try to find the optimal therapy to cure muscle atrophy by improving muscle anabolism. However, we are still a long way from winning the battle against muscle wasting induced by CKD or other metabolic diseases.

Strategies to prevent or treat muscle wasting in CKD include using a bicarbonate regimen to correct metabolic acidosis,³ using anabolic androgenic steroids (derivatives of testosterone),⁴ using peroxisome proliferator-activated receptor inhibitors (i.e., rosiglitazone) to increase insulin sensitivity and reduce insulin resistance,⁵ and providing myostatin inhibitor (peptibody)⁶ or phosphorylated stat3 inhibitors⁷ among others. However, there are no simple and effective treatments for CKD-induced muscle wasting. It has been proved that exercise can prevent muscle wasting in CKD mice⁸ and patients with CKD,⁹ but patients with severe CKD are frequently unable to withstand routine daily physical activity, let alone exercise training.

Acupuncture as a therapeutic intervention is widely used in the United States and around the world.¹⁰ Traditional Chinese medicine considers acupuncture to be a safe, nonpharmacologic intervention.^11,12 Studies show that acupuncture treatment can ameliorate symptoms, such as fatigue, stress, hypertension, proteinuria, and pruritus, in patients with ESRD on dialysis.^13–15 Acupuncture has been shown to decrease skeletal muscle atrophy induced by hind-limb suspension in mice.¹⁶ Electrical acupuncture treatment has been shown to suppress myostatin expression, which leads to a satellite cell-related proliferative reaction and repair in skeletal muscle.¹⁷ Low-frequency electrical stimulation (LFES) is an acupuncture technique that replicates the benefits of resistance exercise through stimulation of muscle contraction.

In skeletal muscle, insulin or IGF-1 plays critical roles in maintaining protein metabolism. In general, upregulation of insulin and IGF-1 signaling will prevent against muscle atrophy induced by CKD.^5,8,18–20 IGF-1 is predominantly made by the liver. Recent studies strongly implicate macrophages as the major extrahepatic source of IGF-1 that contributes to the control of postnatal growth and organ maturation.²¹

Macrophages expressing either killer or repair phenotype are now mainly called M1 or M2 macrophages, respectively.²² M1 and M2 macrophages have distinct chemokine and chemokine receptor profiles. M1 killer macrophages are activated by proinflammatory cytokines, such as IFN-γ. M1 markers include IL-1β and inducible nitric oxide synthase. In contrast, the M2 repair macrophages function in constructive processes, like wound healing and tissue repair, by producing anti-inflammatory mediators, such as arginase-1 and IL-10, which are used as markers for M2 macrophages.²³ Macrophages are normally absent or very low in abundance in muscle and usually of the M2 phenotype.

To design rational therapeutic approaches for the CKD-induced muscle atrophy, in this study, we examined whether LFES affects the IGF-1 signaling pathway. We hypothesized that LFES would upregulate muscle protein synthesis and downregulate protein degradation in muscles of CKD mice. In addition, we report here an extension of the original work that examined whether LFES acts through influences on inflammation and macrophage accumulation, which in turn, upregulate IGF-1 signaling. Finally, we evaluated the effect of LFES on muscle-specific microRNAs (myomiRs) as a potential therapeutic option.

Results

LFES Prevents CKD-Induced Muscle Atrophy and Improves Muscle Grip Function

Mice (C57BL/6J; male; 8 weeks of age) were randomly assigned to four groups: sham, sham/LFES, CKD, and CKD/LFES (n=12/group). In CKD mice, BUN values were 3.4-fold higher than sham-operated pair-fed control mice (P<0.01). The weights of soleus and extensor digitorum longus muscles in the sham/LFES mice were not statistically different from those in sham mice without LFES (Table 1). However, the muscle weights in CKD/LFES mice were significantly higher versus CKD mice in both soleus (CKD: 8.3±0.6 mg; CKD/LFES: 9.5±0.9; P<0.05) and extensor digitorum longus (CKD: 8.2±1.0 mg; CKD/LFES: 9.7±0.4; P<0.05) muscles. Muscle function of the mice was measured using the grip strength meter (Table 2). CKD mice treated with LFES exhibited increased muscle grip capacity compared with CKD mice (CKD: 3.9±0.4 kg force/100; CKD/LFES: 5.9±0.4 kg force/100; P<0.05).

Table 1.

Muscle and body weights

	Sham	LFES	CKD	CKD+LFES
Body weight (g)	23.9±1.3	24.1±1.0	21.3±0.9	22.2±0.6
Soleus (mg)	10.6±1.0	11.9±0.7	8.3±0.6a	9.5±0.9b
EDL (mg)	10.8±0.3	11.0±0.6	8.2±1.0a	9.7±0.4b
Soleus/body (×100)	45.3±4.1	47.5±5.1	41.1±2.2a	44.3±2.9b
EDL/body (×100)	45.3±2.0	48.2±0.2	40.0±9.2a	45.3±6.7b
BUN (mg %)	30.3±2.1	29.2±0.3	103.0±9.2a	99.3±6.7a

Data are presented as mean±SEM (n=12/group). EDL, extensor digitorum longus.

^aP<0.05 is significant versus sham.

^bP<0.05 is significant versus CKD.

Table 2.

Muscle grip strength function was increased by LFES

Groups	Baseline (KGF−2)	Before LFES (KGF−2)	After LFES (KGF−2)
Sham	4.3±0.2	4.1±0.5	4.9±0.7
LFES	4.6±0.6	4.2±0.8	5.5±0.5
CKD	4.3±0.4	4.0±0.9	3.9±0.4a
CKD+LFES	4.3±0.4	4.0±0.7	4.8±0.4b

Data are presented as mean±SEM. KGF⁻², kilogram force/100.

^aP<0.05 is significant versus sham.

^bP<0.05 is significant versus CKD.

LFES Improves Muscle Protein Metabolism

p-Akt in the CKD group was 29% decreased compared with the sham. This lower level, however, was reversed by LFES (Figure 1A). LFES prevented CKD-induced increases in FoxO-1 (active form) and decreases in p-FoxO (p-FoxO-Thr32; inactive form). Foxo3a was unchanged in CKD mice (Figure 1A). To verify that LFES prevents muscle protein catabolism, actin cleavage was measured. CKD increased 14-kD actin fragments by 4.1-fold; LFES prevented CKD-induced actin breakdown (Figure 1B). We next looked at markers of protein synthesis. Phosphorylation of mammalian target of rapamycin (mTOR) was significantly decreased in the muscle of CKD mice versus sham mice. LFES reversed the suppression of p-mTOR, resulting in p-mTOR levels that were 2-fold higher in the CKD/LFES group versus the CKD group. LFES also prevented CKD-induced p-p70S6K depression (Figure 1C). These data indicated that LFES increases muscle protein synthesis and lessens CKD-induced muscle protein catabolism.

Figure 1.

LFES improves muscle protein metabolism markers. (A) Protein metabolism-related proteins (Akt, p-Akt, FoxO1, FoxO3, and p-FoxO), (B) actin cleavage, and (C) synthesis-related proteins (mTOR, p-mTOR, p70S6K, and p-p70S6K) were measured by Western blotting in combined gastrocnemius and extensor digitorum longus muscle lysates from sham, LFES, CKD, or CKD/LFES mice. The bar graphs compare the densities of protein bands in each group expressed as fold changes from levels in sham mice, which are represented by dotted lines at 1-fold. All band densities were normalized to the density of glyceraldehyde-3-phosphate dehydrogenase (GAPDH). Bars: mean±SEM (n=12). *P<0.05 versus CKD; ^#P<0.05 versus sham.

LFES Stimulates Myogenesis

In the muscle of CKD mice, the protein expressions of MyoD (proliferation marker), myogenin (differentiation marker), and eMyHC (differentiation and fusion marker) were significantly decreased versus in sham mice (Figure 2A). The CKD-induced suppression of the myogenesis-related proteins was reversed, and these protein levels were upregulated by LFES. Immunohistochemistry also provides evidence that LFES stimulates myogenesis (Figure 2B). Under normal control conditions, satellite cells, also known as muscle stem cells, are generally located at the periphery of the myofibers. On initiation of myogenesis by LFES (day 1), satellite cells migrate into myofibers (accompanied by increases in MyoD) (Figure 2A). Central nuclei are apparent at day 2, and by day 3, the decrease of central nuclei indicates that fusion has occurred. These results indicate that LFES prevents CKD-induced muscle atrophy, in part, by stimulation of myogenesis.

Figure 2.

LFES counteracts CKD-induced decrease of myogenesis. (A) Muscle protein lysates were prepared from combined gastrocnemius and extensor digitorum longus (EDL) muscles from sham, LFES, CKD, or CKD/LFES mice. Myogenesis-related proteins (myoD, myogenin, and eMyHC) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were measured by Western blotting. The bar graph compares the protein band densities in each treatment group expressed as a fold change from levels in sham mice (represented by a dotted line at 1-fold). All band densities were normalized to the density of GAPDH. Bars: mean±SEM (n=12). *P<0.05 versus CKD; ^#P<0.05 versus sham. (B) Representative cross-sections from EDL muscles after staining for laminin (red) and counterstaining with 4′,6-diamidino-2-phenylindole (blue) are shown. The white arrows point to nuclei inside of myofibers, which indicate migration at control (before LFES) and days 1–3 after LFES. The bar graph shows the nuclei number inside (central nuclei) per 500 muscle fibers. Bars: mean±SEM (n=6). ^#P<0.05 versus control.

LFES Upregulates IGF-1 and Mechanogrowth Factor in Skeletal Muscle of Sham and CKD Mice

To study how LFES prevents CKD-induced muscle atrophy, we identified mRNA expression of growth factors using quantitative real-time PCR (qPCR), which included IGF-1, mechanogrowth factor (MGF; the muscle isoform of IGF-1), basic fibroblast growth factor, and nerve growth factor. LFES increased IGF-1 and MGF expressions 2.5- and 4.7-fold, respectively, in sham mice. IGF-1 expression in CKD muscle was 20% reduced relative to sham mice; however, LFES reversed this trend and resulted in 1.9-fold increased IGF-1 expression and 2.9-fold increased MGF expression in the CKD/LFES group versus CKD (Figure 3, A and B). Nerve growth factor and basic fibroblast growth factor were not changed by LFES in either sham or CKD muscles (data not shown). Protein levels of IGF-1 were measured by ELISA in serum and muscle lysate. In serum, IGF-1 was lower in CKD versus sham mice, and LFES did not have any significant effect on serum IGF-1 levels in either group (Figure 3C). In muscle lysate, CKD showed 50% lower levels of IGF versus sham mice. LFES increased IGF-1 protein 2-fold in the muscles of both groups (Figure 3D), indicating that IGF-1 is locally increased in the LFES-treated mice.

Figure 3.

LFES upregulates IGF-1 and MGF in skeletal muscle of sham and CKD mice. Total RNA isolated from combined gastrocnemius and extensor digitorum longus (EDL) muscles of sham, LFES, CKD, and CKD/LFES mice were assayed for (A) IGF-1 and (B) MGF expression by qPCR. The bar graphs show mRNA from the muscles of each group of mice expressed as fold changes from the sham mice (represented by dotted lines at 1-fold). Results are normalized to 18S RNA. Bars: mean±SEM (n=12). *P<0.05 versus CKD. ^#P<0.05 versus sham. IGF-1 protein levels were measured by ELISA in (C) serum and (D) combined gastrocnemius and EDL lysates from sham, LFES, CKD, or CKD/LFES mice. The bar graphs show the IGF-1 protein levels in each group, and the sham mice are represented by dotted lines. IGF-1 levels in the muscle lysates were normalized to the total protein concentration. Bars: mean±SEM (n=9). *P<0.05 versus CKD. ^#P<0.05 versus sham.

LFES Upregulation of IGF-1 Results from Stimulation of Macrophage Infiltration of Muscle

To determine how LFES upregulated IGF-1, we did separate experiments in mice without CKD to examine inflammation cytokines and macrophage markers. We examined the muscle of LFES mice at days 0 (immediately), 1, 2, 3, and 5 after LFES (Figure 4A). We found that IL-6 was increased 23-fold in the muscle of LFES mice immediately after LFES (day 0; P<0.05 versus control) but did not return to unstimulated levels at day 2. IFN-γ showed a 6.2-fold increase at day 0 (P<0.05 versus control), and TNF-α mRNA showed a trend toward increased expression; however, these changes never reached statistical significance (Figure 4A).

Figure 4.

LFES increases M1 macrophage expression in an early response phase and M2 macrophage expression in a later phase in the muscle of normal mice. (A) Total RNA was isolated from combined gastrocnemius and extensor digitorum longus (EDL) muscles of control and LFES mice. Inflammation cytokines IL-6 (blue), IFN-γ (green), and TNF-α (red) were assayed by qPCR. The graph shows mRNA levels of the cytokines in each group of muscles expressed as a fold change from the controls (without LFES; set up for 1-fold). The x axis presents the time after LFES, with control (Ctrl) being no LFES and day 0 being immediately after LFES. Results are normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA (trend lines on the basis of mean values; n=9). ^#P<0.05 versus control. (B) Muscle cross-sections from control (no LFES) and day 3 after LFES mice. Immunostaining was assessed for F4/80 (green) and 4′,6-diamidino-2-phenylindole (DAPI; blue). Light blue color in the merged picture indicates F4/80-positive cells. The bar graph shows that the percentage of F4/80-positive numbers in total cell numbers in control and days 1, 2, and 3 after LFES. Bars: mean±SEM (n=6). ^#P<0.05 versus control. (C) Total RNA was isolated from combined gastrocnemius and EDL muscles of control and LFES mice. The general macrophage marker F4/80 (red), M1 macrophage marker IL-1β (green), M2 macrophage marker arginase-1 (blue), and IGF (purple) were assayed by qPCR. The graph shows mRNA of the markers in each group of muscles expressed as a fold change of the controls (without LFES; set up for 1-fold). The x axis presents the time after LFES, and day 0 indicates immediately after LFES. Results are normalized to GAPDH mRNA (trend lines on the basis of mean values; n=9). ^#P<0.05 versus control. (D) Muscle cross-sections from control mice (no LFES; upper panel) and mice 3 days after LFES (lower panel). Immunostaining was assessed for IGF-1 (green) and arginase-1 (red). Yellow color in the merged picture indicates that arginase-1–positive cells expressed IGF-1.

We found that the general macrophage marker F4/80 was increased after LFES using immunofluorescence (Figure 4B). We assayed mRNA levels of F4/80 as well as IL-1β (an M1 macrophage marker), arginase-1 (an M2 macrophage marker), and IGF-1 by qPCR (Figure 4C). F4/80 mRNA levels were elevated in the muscle of LFES mice beginning at day 0 through day 5 (P<0.05 at days 0, 1, 2, and 3 versus control). IL-1β was increased at day 0 but began to decrease by day 2 (P<0.05 at day 0 versus control) and was back to control levels at day 3. Arginase-1 was increased at day 1 and consistently increased through day 3 (P<0.05 at days 1, 2, and 3 versus control), after which it decreased to control levels by day 4. IGF-1 showed a similar time course of increase with the M2 macrophage marker arginase-1. These data indicate that LFES upregulates the M1 macrophage markers in an early response phase and the M2 macrophage markers at later time points. In addition, the increase in IGF-1 expression is more temporally aligned with M2 macrophage activation. To identify the relationship between macrophage and IGF-1, muscle cross-sections from control and LFES mice were stained with IL-1β, arginase-1, and IGF-1 antibodies. Macrophage markers were barely found in control muscle but identifiable in LFES muscle. IGF-1 staining is colocalized with the M2 macrophage marker arginase-1 (Figure 4D) but not for the M1 marker IL-1β (data not shown). These results indicate that the LFES-increased IGF-1 could be secreted by M2 macrophages in muscle.

The Effect of LFES on myomiRs

To identify whether LFES stimulates the expression of myomiRs, we measured microRNA by qPCR at various times after LFES. Immediately after the first round of LFES (day 0), the expression of miR-1 is decreased by 26%. Over the next days, the level reached the initial control level, and at later times, it progressed to its highest level at days 3 and 4 (Figure 5). miR-1 returned to its unstimulated control level by day 5. miR-206 showed similar changes (i.e., decreased 19% at day 0 and then increased 2-fold on day 2; P<0.05 versus control). There was no significant change in the levels of miR-133a and miR-133b. These results indicate that the LFES response has two phases. Select myomiRs were initially decreased by LFES and gradually but transiently increased by LFES in a later response.

Figure 5.

LFES alters myomiRs expression. Total RNA was isolated from combined gastrocnemius and extensor digitorum longus muscles of control and LFES mice and then assayed for myomiRs expression. miR-1 (□), miR-206 (X), and miR-133a (Δ) expressions were measured before (Ctrl) and daily after LFES by qPCR with LNA-enhanced oligonucleotide primers. The graph shows myomiRs expression in LFES-treated mice expressed as fold changes of the controls (without LFES; set up for 1-fold). The x axis presents the time after LFES, and day 0 indicates immediately after LFES. Results are normalized to U6 RNA (n=9 pairs). LNA, locked nucleic acid. ^#P<0.05 versus no LFES.

Discussion

This study shows that LFES ameliorates CKD-induced skeletal muscle atrophy by increasing muscle mass and function. Here, we present novel evidence that transient acute inflammation provides benefits that improve muscle protein metabolism and myogenesis to counteract CKD-related muscle wasting.

Our previous studies proved that IGF-1 plays a central role in controlling the muscle wasting of CKD.^5,24–26 In this study, we found that LFES upregulates local IGF-1 and thus, provides a means of increasing muscle mass and function. There are four lines of evidence supporting this conclusion. First, IGF-1 and MGF are substantially increased in the muscle of LFES-treated mice (Figure 3). Second, LFES prevents the increase in protein catabolism induced by CKD (Figure 1, A and B). Third, LFES prevents CKD-induced decrease of muscle protein synthesis markers (Figure 1C). Fourth, LFES stimulates production of muscle regeneration biomarkers and increases satellite cell migration (Figure 2). However, the actual protein synthesis and total protein catabolism were not measured in this study.

There are two mechanisms by which LFES upregulates IGF-1. LFES causes a temporary inflammatory response, indicated by high levels of IL-6 and IFN-γ, and it causes a transient decrease in myomiRs, which result in increased IGF-1.

We found that LFES causes a temporary acute inflammation phenomenon, which is showed by a sharply increased IL-6 and IFN-γ mRNA immediately after LFES. These levels go down over the first 24 hours and are back to normal levels within 48 hours. Inflammation is a common physiologic response to exercise and leads to increased muscle mass.²⁷ IL-6 knockout mice were shown to have an impaired hypertrophic response to overloading.²⁸ We previously reported that myogenesis response to muscle injury was blunted in IL-6 knockout mice,²⁹ which proved the positive effect of IL-6 on myogenesis. Other groups have also reported that electrical stimulation increases IL-6 in both cultured C2C12 cells³⁰ and human primary muscle cells.^31,32 In patients with CKD, Caglar et al.³³ found that the increase in IL-6 concentration was modest during hemodialysis (14%) but that the levels increased dramatically at the end of the 2-hour posthemodialysis period (68% higher compared with baseline).

LFES mimics resistance exercise. Exercise increases muscle mass, in part, through activation of macrophages.^34,35 The type of macrophage involved in the macrophage response depends on their function. M1 killer macrophages are involved in phagocytosis or meeting a microbe challenge, which leads to a sustained inflammation. The other M2 repair macrophages are more closely aligned with repair functions and tend to limit or reverse inflammation.²³ We found that the M1 marker, IL-1β mRNA, is increased in 24 hours and declined at day 3 after LFES. The M2 marker arginase-1 is increased at day 2 and remains at a high level at day 3 after LFES. The time course of the high level of arginase-1 paralleled the time course for high IGF-1 levels, prompting us to hypothesize that accumulation of M2 macrophages leads to upregulation of IGF-1.

IGF-1 is predominantly made by the liver. Several studies strongly suggest that macrophages are a major extrahepatic source of IGF-1.^21,36–38 In an animal injury model, Lu et al.³⁶ found that macrophages of the Ly-6C(-) subtype accumulated in damaged muscle and produced a high level of IGF-1 to promote muscle regeneration. In a human study, IGF-1 has been implicated in the pathogenesis of idiopathic pulmonary fibrosis, and interstitial macrophages were identified as a source of IGF-1.³⁷ In our study, we found that IGF-1 and MGF were increased in muscle by LFES but not in the circulation (serum). We also found that the macrophages numbers were increased in LFES-treated muscle and that M2 macrophages colocalized with IGF-1, indicating that IGF-1 is locally produced by M2 macrophages.

The effect of IL-6 on IGF-1 is controversial. Many studies, including those by our group, found that IL-6 negatively regulates muscle protein metabolism by downregulating IGF-1.³⁹ Raj et al.⁴⁰ showed that IL-6 levels were increased during muscle protein catabolism in patients on hemodialysis. In this study, we found that IL-6 was temporarily increased after LFES. This increase in IL-6 could be facilitating the local infiltration of M2 macrophages that produce anti-inflammation cytokines, which in turn, upregulate locally produced IGF-1. A sustained increase in IL-6 over the long term would consistently induce M1 macrophages to produce inflammation cytokines, leading to downregulation of IGF-1 signaling and increased muscle protein degradation. However, the increase in IL-6 was transient, resulting only in the beneficial recruitment of the M2 macrophages.

The discovery of miRs has increased our knowledge about controlling skeletal muscle metabolism and function. myomiRs, such as miR-1 and miR-206, generally cause a decrease in IGF-1 levels. Both miR-1 and miR-206 target the 3′ untranslated region of IGF-1 and inhibit its translation, leading to a decreased IGF-1 protein amount.^41,42 This study shows that these two myomiRs were decreased in the muscle of CKD mice in the early period after LFES, which could, along with macrophage-mediated stimulation, contribute to the upregulation of IGF-1.

We also found that miR-1 and miR-206 are increased at the later phase after LFES. myomiRs have been reported to play multiple roles in the control of muscle growth and differentiation in the skeletal muscle. Both miR-1 and miR-206 directly target Pax3 and Pax7 and ignite the myogenic program.⁴³ Our study showed that miR-1 and miR-206 were increased by LFES and remained elevated in the muscle of CKD mice over a sustained period, which could be related with increased myogenesis. However, the details of the effect of LFES on myomiRs need additional study. Whether the change of myomiRs is regulated by inflammatory cytokines is another interesting topic for future research.

Our study suggests that acupuncture with LFES could provide a viable therapeutic intervention for treatment of CKD muscle wasting. Actually, acupuncture has been used clinically to treat muscle atrophy in patients with CKD and other patients in the United States and other countries.^13–15 The muscle contractions elicited by LFES provide similar benefits to the patients as exercise would provide. For patients with severe CKD, this treatment may be an alternative way to achieve exercise benefits. The timing and intensity of LFES seem to be very important in the degree of inflammatory response and the accumulation of proinflammatory cytokines. We tried to choose the LFES intensity that (1) showed an effect against muscle wasting and (2) the animal tolerated without apparent distress that might require additional drug treatment for pain. The mice were kept in specially designed restraint, in which the legs that contained the acupuncture needles were immobilized, but the rest of the mouse’s body was free to move. However, the mice barely moved during this 15-minute treatment, which indicated that the mice were comfortable. A stronger intensity of muscle contraction might have potential benefit for protein anabolic response, but because it would cause the animals discomfort, we chose not to include these measurements in this study. There was also the possibility that excessive stimulation could lead to muscle damage, but we did not investigate these conditions in this study. The long-term sustainability of the benefit of LFES has not been established in this study. In addition, this study does not distinguish between the effect of acupuncture and electrical stimulation, which is another limitation of the study. LFES causes an acute inflammatory response, and long-term treatment may cause chronic inflammation, leading to protein catabolism. The intensity and duration conditions for LFES in patient treatment will need to be carefully controlled if it is to be used therapeutically.

In conclusion, LFES ameliorated CKD-induced skeletal muscle atrophy by improvements in muscle protein metabolic status and muscle regeneration capacity, which lead to increased muscle mass and function. Both increasing protein metabolism and myogenesis are the results of improving the insulin/IGF-1 signaling pathway. In this study, we provided evidence suggesting that LFES improves muscle protein metabolism and myogenesis by increasing IGF-1 through two potential mechanisms: (1) a decrease in myomiRs in the early response phase and (2) accumulation of M2 macrophages in the later response phase.

Concise Methods

Animals and CKD Model

The CKD animal model and LFES experiments were approved by the Institutional Animal Care and Use Committee of Emory University. CKD was induced by 5/6 nephrectomy.⁴⁴ Initially, mice were pair fed with a weight-matched sham or a CKD mouse with 14% protein chow for 1 week and then, a high-protein diet (40% protein) for an additional 1 week. BUN is measured the rate of conversion of NADH to NAD monitored at 340 nm using the BUN Kinetic Procedure Kit (Thermo Electron, Louisville, CO).

Muscle function was measured using a mouse grip strength meter with dual computerized sensors to detect and record the grip force (Columbus Instruments, Columbus, OH). Mice were allowed to grip a grid connected to a force transducer and gently pulled by the tail for 5 seconds. The computerized sensors determine what force was needed to counterbalance the grip of the mice. Mice were tested before the CKD surgery (baseline) and before and after LFES. The grip strength of each mouse was tested five times on each testing occasion, with 10 minutes rest between each test. The average of the five determinations was reported.

LFES Treatment

The mice were kept in specially designed restraint without anesthesia so that they would remain in a recumbent position during LFES treatment. Acupuncture points selected were according to the World Health Organization Standard Acupuncture Nomenclature.^45,46 The positive point (GB34, Yang Ling Quan) is under the front head of the fibula about 6 mm (20-g mouse) from the superficial fibular nerve and deep fibular nerve. The negative point (ST36, Zu San Li) is outside of the knee joint under the head of the fibula about 7 mm from the fibular nerve. The needles were connected into an SDZ-II Electronic Acupuncture Instrument using a consistent pulse, an electric frequency of 20 Hz, and electric current of 1 mA. The LFES was administered for 15 minutes every day for 15 days after the second nephrectomy. Disposable sterile needles with a diameter of 0.25 mm (Shen Li Medical & Health Material Co., Ltd., Wujiang, China) were used.

Western Blot and Antibodies

Hind-limb muscles were homogenized in Gentle Lysis Buffer.⁴⁷ Proteins were subjected to Western blot analysis using previously published methods.¹⁹ Primary antibodies (1:1000 dilution except where indicated) that we used included Akt/p-Akt (Ser473), FoxO1/p-FoxO1 (Thr32), FoxO3/p-FoxO3 (Thr32), mTOR/p-mTOR (Ser2448), and 70S6K/p-p70S6K (Thr389) and were from Cell Signaling Technology. MyoD, Myogenin, and eMyHC were from DSHB Product (University of Iowa, Lows, IA). pTEN(FL-403) was from Santa Cruz Biotechnology (Santa Cruz, CA); glyceraldehyde-3-phosphate dehydrogenase was from EMD Millipore (Burlington, MA). Protein bands were scanned and quantified using the Li-Cor Odyssey Infrared Scanning System (Li-COR Biosciences, Lincoln, NE).

Muscle Immunohistology

Muscles were embedded under TBS Tissue Freezing Media (Thermo Fisher Scientific, Pittsburgh, PA) in isopentane cooled in dry ice. The method for immunohistology of muscle cross-sections has been previously described.⁴⁸ The positive cell numbers of at least 500 individual myofibers per muscle were measured using the Micro-Suite Five Biologic Software (Olympus, Melville, NY).

Antibody Usage

Anti-F4/80 was from Abcam, Inc. (Cambridge, MA); anti–Mac-2 and anti–arginase-1 were from Santa Cruz Biotechnology. Polyclonal anti-Laminin antibody (1:50 dilution; L9393; Sigma-Aldrich) and goat anti-mice IGF-1 were from Lifespan Bioscience (Seattle, WA).

Quantitative Measurement of Mouse IGF-1 in Serum and Muscle Lysis

The IGF-1 Mouse ELISA Kit (ab100695) was purposed from Abcam, Inc. and used according to the manufacturer’s instructions.

Reverse Transcription and qPCR

Total RNA was extracted using Tri-Reagent (Molecular Research Inc., Cincinnati, OH). RNA was subjected to reverse transcription and qPCR using previously published methods.⁴⁷ Primers were designed to cross intron–exon boundaries as described in Table 3. For myomiRs, the miRCURY LNA Universal cDNA Synthesis Kit (Exiqon Inc., Woburn, MA) was used for reverse transcription of RNA. The primers were custom designed by Exiqon Inc. The miRCUTY LNA microRNA PCR SYBR Green Master Mix (Exiqon Inc.) was used for qPCR with the following cycle parameters: 95°C for 10 minutes and 45 cycles at 95°C for 10 seconds and 60°C for 60 seconds. Expression of individual myomiRs was standardized to the mouse U6 gene and calculated as the difference between the threshold values of the two genes (ΔΔcq).

Table 3.

Primer sequence

Name	Sequence	Amplicon	Code
IL-6
Forward	TTC CAT CCA GTT GCC TTC TTG	101	NM_031168
Reverse	TTG GGA GTG GTA TCC TCT GTGA
IFN-γ
Forward	TGCTGATGGGAGGAGATGTCT	101	NM_008337
Reverse	TTTCTTTCAGGGACAGCCTGTT
TNF-α
Forward	GACGTGGAACTGGCAGAAGAG	101	U68415.1
Reverse	GCCACAAGCAGGAATGAGAAG
F4/80
Forward	CTTTGGCTATGGGCTTCCAGTC	165	X93328
Reverse	GCAAGGAGGACAGAGTTTATCGTG
IL-1β
Forward	CTTCCCCAGGGCATGTTAAG	101	NM_008361
Reverse	ACCCTGAGCGACCTGTCTTG
IGF-1
Forward	GACCGCACCTGCAATAAAG	91	NM_010512
Reverse	TGTGGTGGAGCTGGTGAAG
MGF
Forward	GACATGCCCAAGACTCAG	80	NM_184052
Reverse	GGCTCACCTTTCCTTCTC
Arginase-1
Forward	AACACGGCAGTGGCTTTAAC	168	NM_007482
Reverse	GAGGAGAAGGCGTTTGCTTA
Nerve growth factor
Forward	GACAGTGTCAGCGTGTGGGTT	74	NM_001112698
Reverse	CCAACACCATCACCTCCTT
bFGF
Forward	GAGAAGAGCGACCCACACG	78	NM_008006
Reverse	GGCACACACTCCCTTGATAGA
18S
Forward	CCA GAG CGA AAG CAT TTG CCA AGA	101	X00686
Reverse	TCG GCA TCG TTT ATG GTC GGA ACT

bFGF, basic fibroblast growth factor.

Statistical Analyses

Data were presented as mean±SEM. To identify significant differences between two groups, comparisons were made using the t test. When multiple treatments were compared, ANOVA was performed. Differences with P values<0.05 were considered significant.

Disclosures

None.