Acetylcholine receptor epsilon-subunit deletion causes muscle weakness and atrophy in juvenile and adult mice

Abstract

In mammalian muscle a postnatal switch in functional properties of neuromuscular transmission occurs when miniature end plate currents become shorter and the conductance and Ca2+ permeability of end plate channels increases. These changes are due to replacement during early neonatal development of the gamma-subunit of the fetal acetylcholine receptor (AChR) by the epsilon-subunit. The long-term functional consequences of this switch for neuromuscular transmission and motor behavior of the animal remained elusive. We report that deletion of the epsilon-subunit gene caused in homozygous mutant mice the persistence of gamma-subunit gene expression in juvenile and adult animals. Neuromuscular transmission in these animals is based on fetal type AChRs present in the end plate at reduced density. Impaired neuromuscular transmission, progressive muscle weakness, and atrophy caused premature death 2 to 3 months after birth. The results demonstrate that postnatal incorporation into the end plate of epsilon-subunit containing AChRs is essential for normal development of skeletal muscle.

Figure 1

Targeted disruption of the gene encoding the ɛ-subunit of muscle AChR. (A) Schematic representation of AChR ɛ-subunit cDNA with exons E7, E8, E9, and E12 (solid bars). The regions encoding M1, M2, M3, and M4 segments (open boxes), which are essential for the assembly of functional AChR complexes, are indicated. The wild-type ɛ-subunit allele carries the putative membrane regions in exons 7, 8, 9, and 12. The targeting vector consists of 3.3 kb of 5′ untranslated sequences and 1.9 kb containing exons 1 to 7. A PGK-neo cassette (1.9 kb) was inserted to replace sequences from the AatII restriction site of exon 7 to the Nsil site located in intron 8. Deletion of this sequence region in the mutated product was expected to completely prevent the assembly of functional ɛ-AChR channels in adult muscle. The final construct contains a 5.2-kb 5′ homologous region and a 1.4-kb 3′ homologous region. The 3′ end extended from the NsiI site to the XhoI site of exon 10. Hybridization probe used for screening of ES cells and mutant mice is indicated as thin bar and represents a 1.1-kb genomic XhoI/Sau3AI DNA fragment excised from cloned genomic DNA. Thick black bars represent the different BstEII fragments expected from the wild-type and mutated allele corresponding to 3 kb and 5.3 kb, respectively. (B) Southern blot analysis performed with DNAs isolated from tails of AChRɛ^+/− × AChRɛ^+/− crosses. DNAs from AChRɛ^+/+, AChRɛ^+/−, and AChRɛ^−/− mice were digested with BstEII and hybridized with XhoI/Sau3AI probe as shown in A. The wild-type allele generates a 3-kb fragment and the mutant allele a 5.3-kb fragment.

Figure 2

AChR γ- and ɛ-subunit transcripts and AChR expression at end plates. (A) Reverse transcription–PCR. Total RNA from hind leg muscles was extracted at P5 and P34. Transcript levels were low, and direct visualization of ethidium stained PCR products gave no clear results for γ-subunit transcripts at P34. PCR products were therefore transferred to Biodyne A membranes and were stained using an Enhanced Chemiluminescence detection kit. (B and C) AChRs visualized with rhodamine-labeled α-bungarotoxin. Diaphragm of 3-month-old wild-type (B) and AChR ɛ^−/− mutant mouse (C) littermates of line 399. (Bars = 75 μm.)

Figure 2

AChR γ- and ɛ-subunit transcripts and AChR expression at end plates. (A) Reverse transcription–PCR. Total RNA from hind leg muscles was extracted at P5 and P34. Transcript levels were low, and direct visualization of ethidium stained PCR products gave no clear results for γ-subunit transcripts at P34. PCR products were therefore transferred to Biodyne A membranes and were stained using an Enhanced Chemiluminescence detection kit. (B and C) AChRs visualized with rhodamine-labeled α-bungarotoxin. Diaphragm of 3-month-old wild-type (B) and AChR ɛ^−/− mutant mouse (C) littermates of line 399. (Bars = 75 μm.)

Figure 3

Low conductance end plate channels persist in AChRɛ^−/− mice. (A) Single channel currents in wild-type (wt) (Left) and AChRɛ^−/− muscle (Right) recorded from P5 (upper traces) and P30 (lower traces) end plate. Membrane potential was approximately −100 mV in each experiment. Membrane potential was estimated from current reversal potential measurement in the same patch. Temperature 20°C, inward current is downward. (B) Amplitude distribution of single channel currents recorded from the end-plate of wild-type (wt) and AChRɛ^−/− muscle at P30 as shown in A. Each histogram was fitted to a single Gaussian and show that current amplitudes in both patches represent homogeneous conductance class channels. Mean amplitudes are 3.3 ± 0.1 pA (AChRɛ^−/−) and 5.4 ± 0.2 pA (wt). (C) Current-voltage (I–V) relations of single end plate channels in P30 wild-type (open symbols) and AChRɛ^−/− muscle (solid symbols). Voltage (V) represents shift of patch-membrane potential from extrapolated current reversal potential. Straight lines (not shown) were fitted to data points in the voltage range of −20 mV to −100 mV. Slope conductance is 32 pS (AChRɛ^−/−) and 52 pS (wt), respectively. Different experiment from that shown in A and B.

Figure 4

Amplitude and decay time course of miniature end plate currents. Records of representative mEPCs from wild-type (Left) and AChRɛ^−/− mutant mouse muscle (Right). All records at −70 mV, 20–21°C. Inward current is downward. Sampling at 15 kHz. Scale bars in C refer to all traces. (A) P5 muscles. Peak mEPC amplitude is 3.2 nA and decay time constant is 4.6 ms for wild-type and 3.2 nA and 4.7 ms for AChRɛ^−/− end plate. (B) P30 muscles: 3.3 nA and 1.0 ms for wild-type and 2.3 nA and 4.2 ms for AChRɛ^−/− end plate. (C) P70 muscles: 3.3 nA and 1.0 ms for wild-type and 1.0 nA and 4.2 ms for AChRɛ^−/− end plate.

Figure 5

Reduction of isometric tetanic muscle contraction force. (A) Isometric tension measurements on soleus muscle in vitro from a heterozygous AChRɛ^+/− animal at P32. Single twitches and tetanic contractions were evoked by direct muscle stimulation or by nerve stimulation in normal Tyrode solution. The records of contractions evoked by nerve or muscle stimulation superimposed perfectly indicating no difference in contraction force upon nerve or muscle stimulation. (B) Isometric tension measurements on soleus muscle from an AChRɛ^−/− animal at P32. Single twitches and tetanic contractions evoked by direct muscle stimulation (m; solid lines) and nerve stimulation (n; dashed lines) are superimposed for direct comparison. Note different scales in A and B. Tetanic stimulation frequencies (T) were 20 or 100 Hz as indicated. Vertical scale bars indicate muscle contraction force in mN; horizontal ones indicate duration of stimulation in seconds (s). Body weight of the three animals studied in each group was on average 22 ± 4.1 g for the AChRɛ^+/− mice and 11 ± 1.1 g for AChRɛ^−/− mice; wet weight of soleus muscles was 11.5 mg ± 2.2 and 7.5 mg ± 1.1, respectively (n = 6 each) (P < 0.005 for each comparison, t test).

Figure 6

Differential changes in muscle strength and body weight during development. (A) Forelimb grip strength (measured in N) of male wild-type or heterozygous AChRɛ^+/− mice (○), and homozygous AChRɛ^−/− mutant mice (•). (B) Body weight (measured in g) of male wild-type or heterozygous AChRɛ^+/− mice (○) and of male AChRɛ^−/− mice (•) during postnatal development. Mean values ± 1 SD (three to five animals) are shown in A and B. Symbols without error bars represent individual animals.