The Angiotensin Converting Enzyme Inhibitor Lisinopril Improves Muscle Histopathology but not Contractile Function in a Mouse Model of Duchenne Muscular Dystrophy

Mice and treatment

All protocols were approved by the Institutional Animal Care and Use Committee of The Ohio State University and are in compliance with the laws of The United States of America and conform to the Guide for the Care and Use of Laboratory Animals published by the United States National Institutes of Health.

For this study we used a dystrophin-deficient, utrophin haplo-insufficient (utrn  ^+/- ; mdx) “het” mouse model, which has a more severe phenotype than mdx mice [9, 10]. C57BL/10 (Harwell) wild-type control mice were also used in this study. Dystrophic and wild-type control mice were bred and dystrophic mice were genotyped, as described previously [11]. Mice were weaned at 4 weeks-of-age and males and females from each litter were housed 2 per cage and used for treatment or as untreated controls. Mice were treated for 6 weeks and then analyzed by personnel not involved in genotyping or treating the animals and blinded to the groups.

Equal numbers of female and male het mice were given water bottles containing 66, 133, or 330 mg/L of the ACEi lisinopril (SBH Medical) in reverse-osmosis water or no drug (reverse-osmosis water only). Medicated water bottles were replaced 3 times per week; mice were weighed and the volume of water consumed was recorded to ensure mice were receiving the estimated dosage (10, 20 or 50 mg/kg × day). The 2 lowest dosages were based on allometric scaling of clinical dosages to use in mice (based on the FDA: Guidance for Industry; Estimating the Maximum Safe Starting Dose in Initial Clinical Trials for Therapeutics in Adult Healthy Volunteers) and are commonly used for experiments in cardiomyopathic mice and rats. The lowest dosage (10 mg/kg × day) was that used in our previous combinatorial ACEi plus MR antagonist treatments shown to be efficacious in dystrophic mice [4, 12]. 50 mg/kg × day was included as a 5X normal dosage to capture any increased dose-response. In total, 5 groups of mice were included in this study: C57BL/10 wild-type control mice (C, n = 10), untreated het mice (U, n = 10), and lisinopril treated het mice at 3 different dosages (L10, L20, L50, n = 6 respectively). The combination of the three groups of het mice receiving lisinopril is referred to as total lisinopril treated mice (L) for some dataanalyses.

In vivo cardiac and forelimb grip strength measurements

All in vivo measurements were performed by individuals blinded to the genotype and treatment of the mice. Each in vivo measurement was performed bythe same individual to limit variability. Within 4 days of the animals reaching 10 weeks-of-age, het untreated and C57BL/10 wild-type control mice were anesthetized (isoflurane) and magnetic resonance imaging (MRI) was performed on a 9.4 or 11.7 Tesla 30 mm bore system (Bruker Biospin) with electrocardiographic (ECG) leads while under body temperature control (37°C), as described previously [4]. Myocardial strain and strain rate were computed using vector-based tracking software (Vector Velocity Imaging, Siemens).

Blood pressure was measured non-invasively in conscious mice by the tail cuff method using a 6-channel CODA High Throughput Acquisition System (Kent Scientific Corporation). Measurement recording sessions included 10 acclimation cycles followed by 10 measurement cycles. The average of accepted measurement cycles for each mouse was used for systolic, diastolic, and mean arterial blood pressure [13]. Measurements were done between 10:00 AM and 12:00 PM in untrained mice at least one day before ECG, grip strength, and in vitro muscle force experiments.

On the day of sacrifice, the body weight of each mouse was recorded and resting, non-anesthetized, and non-invasive electrocardiographic recordings were taken using the ECGenie system (Mouse Specifics Inc.) as described previously [4]. Briefly, mice were placed on the ECG platform in a quiet room and allowed sufficient time (typically 5– 10 minutes) for heart rate to level after handling. ECG tracings were continuously recorded for 30 minutes using LabChart7 (AD Instruments), and later analyzed using ECG e-Mouse 9 software (Mouse Specifics Inc). Analysis was done using time intervals when the paws were in contact with the electrodes (typically 10– 15 seconds) and heart rate (HR) remained consistent. ECG signals were used to assess: QT-interval, QT-interval:heart rate, and heart rate variability.

Forelimb grip strength was then assessed in conscious, non-anesthetized mice. The grip strength meter (Columbus Instruments) consists of a wire mesh grid attached to a force transducer atop a sturdy aluminum plate. Mice were suspended so that their front paws were able to grasp the center edge of the wire mesh. Once the mouse had established a grip without body movement, it was swiftly and uniformly pulled backwards in plane with the transducer to break the clasp [3] and as described in Treat-NMD SOP DMD_M.2.2.001. This procedure was repeated three times with one-minute rests between measurements. The highest value (N) of the three was reported. All grip strength measurements were conducted by the same investigator to limit variability.

In vitro diaphragm and extensor digitorum longus (EDL) contraction force measurements

All force measurements were performed by individuals blinded to the genotype and treatment of the mice. Measurement of each muscle type was performed by the same individual to limit variability. Mice were sacrificed by cervical dislocation. The thorax was opened and the diaphragm and ribcage were removed and placed in a modified Krebs-Henseleit solution [95% O₂/5% CO₂ (pH 7.4), 5 mM KCl, 137 mM NaCl, 1.2 mM NaH₂PO₄, 1.2 mM MgSO₄, 20 mM NaHCO₃, 10 mM D-glucose, and 0.25 mM CaCl₂ ] with 20 mM 2,3-butanedione monoxime (BDM) added to prevent muscle damage during dissection. All regents were purchased from Sigma Aldrich.

Two linear strips of muscle approximately 2-3 mm in width were carefully dissected from the center of each diaphragm so that one end of the muscle had rib attached. This allowed the diaphragm strip to be held in place, inside the muscle bath, by inserting the muscle through a stainless steel basket connected to a KG2 force transducer (World Precision Instruments). The central tendon on the interior end of the diaphragm strip was then pierced over a hook connected to a linear micro-manipulator. The muscle bath contained oxygenated Krebs-Henseleit solution as described above (without BDM and CaCl₂ raised to 2.0 mM) at 37°C, since this is the temperature at which the in vivo diaphragm operates [14]. Single electrical stimulation pulses (180 Hz for 4 ms) were delivered via two parallel platinum-iridium electrodes surrounding the muscle to determine the “optimal length” for measuring maximum twitch force. The muscles were subsequently allowed to rest for 10 min, and then subjected to a series of tetanic contractions. Six tetanic contractions, one at each 20, 50, 80, 120, 150, and 180 Hz (250 ms duration each) were performed with 2 minutes of rest between stimulations. 5 minutes after the final tetanic measurement, a fatigue protocol (stimuli at a frequency of 100 Hz and 250 ms duration each second for a total of 66 seconds) was performed. After a 15 minute rest the protocol was repeated.

For comparative purposes, all force measurements are expressed per unit of cross-sectional area (CSA) (normalized isometric force or tension: mN/mm²). CSA was calculated using the following equation: muscle mass in mg/[(optimal length in mm) × (muscle density in mg/mm³)], where muscle density is assumed to be 1.056 mg/mm³. Mass was determined after all force measurements by blotting the muscle between two Kimwipes beneath a 10 g weight for 10 seconds [12].

The EDL muscle from each leg was carefully excised under a dissection microscope. Before removal, two pieces of suture were used to tie a knot on each tendon. The muscle was then removed and placed in the modified Krebs-Henseleit solution (with BDM) while 5– 8 additional knots were tied onto each tendon using the original suture ends described above. Sutures were tied around the tendons on each end of the muscle so that knots could be placed over two hooks (one attached to a force transducer, the other a linear micromanipulator) in a muscle bath through which oxygenated Krebs-Henseleit (without BDM and 2.0 mM CaCl₂ added) at 30°C was continuously cycled. This multi-knot attachment method removes any attachment compliance and ensures that all length changes are directly reflecting changes in the muscle length. Muscles were stretched to optimal length using twitch contractions (evolked by a single 4 ms pulse) as previously described [12, 14]. After 10 minutes, a tetanus contraction was performed (150 Hz for 250 ms). After another 5 min rest period, 10 eccentric contractions (150 Hz for 450 ms, subjected to a 3% stretch for the final 200 ms of contraction) were done with two minutes of rest between stimulations. Following force measurements, the sutures were removed, and muscles were weighed as described above. Forces are expressed per unit of cross-sectional area (CSA).

Force recordings and analysis were done using custom-made LabView (National Instruments) programs. If more than one diaphragm strip or both EDL muscles were assessed from the same mouse, values were averaged for each muscle type and treated as n = 1 for statistical analyses [12].

Histopathology and quantitation

All histological assessments were performed by individuals blinded to genotypes and treatments of the mice. Each measurement was performed by the same individual to limit variability. From each mouse used for all of the above measurements, additional striated muscle tissues were embedded in optimal-cutting temperature (OCT) medium and frozen on liquid-nitrogen cooled isopentane for histological analyses. These samples included heart, quadriceps, rectus abdominis, and the remaining portions of the diaphragm muscle. 8μm sections were cut using Cryostat model OTF 5000 (Bright). One section from each tissue block was stained with Hematoxylin and Eosin (H&E), using standard methods to assess overall histopathology. An additional section from each tissue block was used for immunofluorescence experiments for analysis of ongoing muscle damage, with an antibody against mouse IgG, as previously described (Alexa 488 goat anti-mouse IgG, 1:200; Life Technologies) [4].

Wide-field images were taken on a Nikon Eclipse 800 microscope through a 10X objective using a SPOT RT slider digital camera and SPOT software. The percentage of damage in cardiac, quadriceps and abdominal muscle tissue was assessed on IgG stained sections using Photoshop CS6 (Adobe) similar to that described previously [12]. The photographs were quickly composited using the automation function “Photomerge.” For all composite images, the background and any holes within the sections were painted white using the paint-bucket tool. For IgG stained sections, damaged areas were painted blue using the paint-bucket tool. Normal IgG localization in the capillaries and endomysium was left unpainted. Painted sections were analyzed using the magic-wand tool and the histogram window. The magic wand tool was used to select all blue portions of the section and the number of selected pixels was shown in the histogram. The magic wand tool was then used to select all white portions of the section. The inverse function was used to switch the selection to the entire section including damage (everything that is not white), the total number of selected pixels was shown in the histogram. Damage is reported as a percentage of the entire section. Longitudinal sections were excluded from analysis.

Data and statistical analysis

All data was included for statistical calculations and analyzed using one-way ANOVA. If the overall ANOVA indicated statistical significance, the non-parametric Dunnett post-hoc test was used to test for significant differences to all other groups compared with the untreated het mice. The untreated het mice allow for the most unambiguous interpretation as they are the only group that differ in only 1 degree of freedom (genotype or dose) from all other groups. Summary values are presented as mean ± SE. Two-tailed P values <  0.05 were accepted as significant. The percentage of damage in quadriceps muscle tissue and the percentage of EDL specific force ratio data were also compared with 20 week-old male C57BL/10 wild-type control mice and untreated het mice.

Lisinopril treatment does not affect mouse body weight or grip strength

Body weight was not different between treatment groups (P = 0.1572; Fig. 1A). Forelimb total grip strength (P = 0.0001) and normalized forelimb grip strength (P <  0.0001) was significantly (Dunnett) higher only in C57BL/10 compared to untreated het mice (Fig. 1B and not shown). Treatment of het mice with lisinopril did not significantly improve total or normalized forelimb grip strength at any of the three dosages tested (10, 20, or 50 mg/kg × day [L10, L20, L50]) (Fig. 1B and not shown).

Diaphragm and extensor digitorum longus (EDL) muscle forces are not improved with lisinopril treatment

Only C57BL/10 mice showed a significant difference (P = 0.0009) in maximal in vitro specific contractile force in diaphragm muscle compared with the untreated het mice, suggesting lisinopril treatment has no beneficial effect on diaphragm function (Fig. 2A). Diaphragm specific forces were 196 ± 21 mN/mm² for C57BL/10 compared with 118 ± 8 mN/mm² for untreated hets and 122 ± 9 mN/mm² for all of the lisinopril-treated mice combined. The highest lisinopril dosage only resulted in a diaphragm specific force of 128 ± 13 mN/mm². EDL specific force (Fig. 2B) was statistically similar between all groups, including C57/BL10 control mice. C57BL/10 and untreated het mice had an average EDL specific force of 343 ± 23 and 301 ± 26 mN/mm², respectively, compared to 343 ± 46 mN/mm² for the het mice treated with the highest lisinopril dosage (L50).

Lisinopril treatment reduces muscle damage

Histopathological analysis was performed on quadriceps, diaphragm and abdominal muscles from all treatment groups. Hematoxylin and eosin (H&E) staining was used to examine the overall pathology of the muscle. Immunofluorescence staining for accumulation of serum proteins (IgG) both intracellularly and in fibrotic areas replacing muscle fibers provided a quantifiable measurement of damage. Representative H&E staining and IgG immunofluorescence for C57BL/10, untreated het, and het mice treated with the highest dosage of lisinopril (50 mg/kg × day) are shown (Figs. 3, 4A). Het mice treated with 50 mg/kg × day of lisinopril (L50) showed reduced overall pathology in quadriceps, diaphragm, and abdominal muscles, compared with untreated het mice (Fig. 3).

A similar pattern was also observed with IgG staining. The highest dosage lisinopril treatment group showed some abnormal IgG staining in all three muscle types analyzed, but was not as severe as observed in untreated het mice (Fig. 4A). To measure the effect of lisinopril on muscle damage in het mice, IgG localization within myofibers, replacing myofibers, and abnormal staining in endomysium, was quantified as a percentage of the total muscle section area for all three muscle types. As expected, C57BL/10 wild-type controls had very little or no muscle damage in any muscle type (Figs. 3, 4). ANOVA revealed a significant difference between groups in the amount of muscle damage in quadriceps (P = 0.00071) and diaphragm (P <  0.0001) muscles. The percentage of damage in quadriceps muscle was significantly lower in C57/BL10 mice (Dunnett, P <  0.0001), all lisinopril treated het mice (L10 + L20 + L50; P = 0.013) and mice treated with both 20 (L20; P = 0.031), or 50 mg/kg × day of lisinopril (L50; P = 0.0239) compared to untreated het mice (Fig. 4B). The highest lisinopril-treatment group had 9.1 ± 1.9% IgG staining in quadriceps muscles, compared to 20.3 ± 3.8% in the untreated het group and 0.7 ± 0.1% in the C57BL/10 control group. A similar reduction was seen in diaphragm muscle, for C57/BL10 control mice (P <  0.0001) and mice treated with 50 mg/kg × day of lisinopril (L50 mg) (P = 0.0494) compared to untreated mice (Fig. 4C). L50 lisinopril treated mice had 14.9 ± 0.8% IgG staining in diaphragm muscle compared to 19.3 ± 1.5% in the untreated het and 1.9 ± 0.4% in the C57BL/10 control group.

There was also a significant difference between groups for abdominal muscle damage (P = 0.00314). However, no significant reduction was observed between lisinopril treated mice (L20, P = 0.8862; L50, P = 0.1395) and untreated het mice although there was a trend towards improvement with lisinopril treatment with a reduction from 16.4 ± 3.1% in untreated to 9.3 ± 1.9% in the L50 group. Only C57BL/10 wild type and untreated het were significantly different for abdominal muscles (P = 0.0012). These results suggest lisinopril treatment leads to a limited reduction in abdominal muscle damage (Fig. 4D).

Heart rate, and blood pressure are unaffected by lisinopril treatment, but QT-interval is lower in lisinopril treated mice

Blood pressure recordings measured by tail cuff and heart rate measurements from electrocardiogram recordings, in conscious mice, showed no significant differences between any of the groups (P = 0.17 and 0.46 respectively, Table 1). However, QT-interval was significantly lower in total lisinopril treated het mice (P = 0.001), the two higher dosage lisinopril treatment groups (L20, P = 0.0146; and L50, P = 0.0025) and C57BL/10 (P = 0.027) mice compared to the untreated het mice (Table 1). The ratio between QT – interval:heart rate (HR) was also different (P = 0.019) between total lisinopril (P = 0.0049), L20 mg (P = 0.04) and L50 mg (P = 0.0235) treated mice compared to untreated het mice (Table 1). A summary of heart rate, mean arterial pressure, QT-interval and QT-interval:heart rate ratio is shown in Table 1.

Analysis at 20 weeks-of-age provides more informative outcome measures than 10 weeks-of-age for all compared parameters except ongoing quadriceps muscle damage

To determine whether 10 or 20 weeks-of-age is a better time point for analyzing treatment efficacy infuture preclinical studies, several parameters were compared between C57BL/10 and untreated het mice. Differences in EDL specific force measurements between untreated het and C57BL/10 mice at 20 weeks-of-age were more informative than at 10 weeks-of-age (Fig. 5A). In the current study, 20 week-old untreated het mice showed 59% of the specific EDL force observed in 20 week-old C57BL/10 controls, compared to 88% of wild-type force levels observed in 10 week-old untreated het mice compared to 10 week-old C57BL/10 mice. However, ongoing damage in quadriceps muscles measured by IgG staining is more informative at 10 weeks-of-age (20.3% of total muscle section area) than 20 weeks-of-age (4.4% of total muscle section area) in het untreated mice(Fig. 5B).

Importantly, analysis of cardiomyopathy is uninformative at 10 weeks-of-age in het mice. H&E and IgG staining of the entire left ventricles of hearts from 10 week-old untreated het mice did not show quantifiable damage compared to that previously observed and quantified in 20 week-old het mice (Fig. 5C, and not shown). Furthermore, cardiovascular MRI performed on untreated het mice and C57BL/10 wild-type control mice at 10 weeks-of-age showed no significant differences in peak systolic strain rate at the base of left ventricle between the 2 groups (P = 0.097, Fig. 5D), compared to previous differences observed at 20 weeks-of-age. These data suggest that cardiomyopathic changes are not yet detectable in 10 week-old het mice and preclinical studies to detect therapeutic effects on the heart will need to be carried out in 20 week or older het mice.