A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle 2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach.The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, β2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and α-actinin; and membrane proteins, i.e. α1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats.The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs.The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6–8.

Two hybrid voltage-sensing systems based on fluorescence resonance energy transfer (FRET) were used to record membrane potential changes in the transverse tubular system (TTS) and surface membranes of adult mice skeletal muscle fibers. Farnesylated EGFP or ECFP (EGFP-F and ECFP-F) were used as immobile FRET donors, and either non-fluorescent (dipicrylamine [DPA]) or fluorescent (oxonol dye DiBAC4(5)) lipophilic anions were used as mobile energy acceptors. Flexor digitorum brevis (FDB) muscles were transfected by in vivo electroporation with pEGFP-F and pECFP-F. Farnesylated fluorescent proteins were efficiently expressed in the TTS and surface membranes. Voltage-dependent optical signals resulting from resonance energy transfer from fluorescent proteins to DPA were named QRET transients, to distinguish them from FRET transients recorded using DiBAC4(5). The peak ΔF/F of QRET transients elicited by action potential stimulation is twice larger in fibers expressing ECFP-F as those with EGFP-F (7.1% vs. 3.6%). These data provide a unique experimental demonstration of the importance of the spectral overlap in FRET. The voltage sensitivity of QRET and FRET signals was demonstrated to correspond to the voltage-dependent translocation of the charged acceptors, which manifest as nonlinear components in current records. For DPA, both electrical and QRET data were predicted by radial cable model simulations in which the maximal time constant of charge translocation was 0.6 ms. FRET signals recorded in response to action potentials in fibers stained with DiBAC4(5) exhibit ΔF/F amplitudes as large as 28%, but their rising phase was slower than those of QRET signals. Model simulations require a time constant for charge translocation of 1.6 ms in order to predict current and FRET data. Our results provide the basis for the potential use of lipophilic ions as tools to test for fast voltage-dependent conformational changes of membrane proteins in the TTS.

Duchenne muscular dystrophy is caused by mutations in DMD which disrupt the reading frame. Therapeutic strategies that restore DMD’s reading frame, such as exon skipping and CRISPR/Cas9, need to be tested in the context of the human DMD sequence in vivo. We have developed a novel dystrophic mouse model by using CRISPR/Cas9 to delete exon 45 in the human DMD gene in hDMD mice, which places DMD out-of-frame. We have utilized this model to demonstrate that our clinically-relevant CRISPR/Cas9 platform, which targets deletion of human DMD exons 45–55, can be directly applied in vivo to restore dystrophin.

Repetitive activation of skeletal muscle fibers leads to a reduced transmembrane K+ gradient. The resulting membrane depolarization has been proposed to play a major role in the onset of muscle fatigue. Nevertheless, raising the extracellular K+ (\documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$ {\text{K}}_{\text{o}}^{ + } $$\end{document}) concentration (\documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$ [ {\text{K}}^{ + } ]_{\text{o}} $$\end{document}) to 10 mM potentiates twitch force of rested amphibian and mammalian fibers. We used a double Vaseline gap method to simultaneously record action potentials (AP) and Ca2+ transients from rested frog fibers activated by single and tetanic stimulation (10 pulses, 100 Hz) at various \documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$ [ {\text{K}}^{ + } ]_{\text{o}} $$\end{document} and membrane potentials. Depolarization resulting from current injection or raised \documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$ [ {\text{K}}^{ + } ]_{\text{o}} $$\end{document} produced an increase in the resting [Ca2+]. Ca2+ transients elicited by single stimulation were potentiated by depolarization from −80 to −60 mV but markedly depressed by further depolarization. Potentiation was inversely correlated with a reduction in the amplitude, overshoot and duration of APs. Similar effects were found for the Ca2+ transients elicited by the first pulse of 100 Hz trains. Depression or block of Ca2+ transient in response to the 2nd to 10th pulses of 100 Hz trains was observed at smaller depolarizations as compared to that seen when using single stimulation. Changes in Ca2+ transients along the trains were associated with impaired or abortive APs. Raising \documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$ [ {\text{K}}^{ + } ]_{\text{o}} $$\end{document} to 10 mM potentiated Ca2+ transients elicited by single and tetanic stimulation, while raising \documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{...