Our favored testing approach is exome based NextGen sequencing with CNV analysis. This will allow cost effective reflexing to PGxome or other exome based tests. However, if full gene Sanger sequencing is desired for STAT turnaround time, insurance, or other reasons, please see link below for Test Code, pricing, and turnaround time information. If the Sanger option is selected, CNV detection may be ordered through Test #600.

The great majority of tests are completed within 26 days.

Congenital myopathy refers to a group of neuromuscular disorders that most often present in childhood and have characteristic histopathological features in skeletal muscle. Based on histopathology, the three main types of congenital myopathy are core myopathy, centronuclear myopathy, and nemaline rod myopathy. Pathological RYR1 gene variants are causative for core myopathy, centronuclear myopathy, and core/rod myopathy. Cores are areas of the muscle that abnormally lack oxidative and glycolytic enzymatic activity. These areas of clearing reflect an absence of mitochondria in the muscle. Cores are further classified as central cores, which run the length of the muscle cell, or minicores, which are wider than they are long. Nemaline rods are rod-shaped structures in the sarcoplasm that contain material from the degenerated Z-disk.

Central core disease (CCD) is a static or slowly progressive myopathy that demonstrates a wide range of clinical severity and age of onset, but uniform histopathology. In the newborn period, CCD results in a floppy infant with feeding difficulties, respiratory insufficiency, myopathic facies, and extraocular muscle weakness. Skeletal abnormalities include talipes equinovarus and congenital dislocation of the hips. Childhood-onset CCD presents with mild proximal weakness involving hip and axial muscles and delayed motor development. Skeletal findings such as scoliosis and equinovarus are other complications (Jungbluth et al. 2007). Cases of adult-onset CCD with generally milder clinical symptoms have been described (Duarte et al. 2011). One representative patient had normal arm and leg strength and normal muscle bulk, but was easily fatigued and had difficulty maintaining an upright posture (Jungbluth et al. 2009). Intrafamilial variability of disease severity and pathology is also known (Sewry et al. 2002). Imaging of muscle groups in CCD patients reveals a distinct pattern of involvement (Fischer et al. 2006). The most severely affected muscles are gluteus maximus, medial and anterior compartments of the thigh muscles, and soleus and lateral gastrocnemius muscles of the lower leg.

The other congenital myopathies associated with the RYR1 gene demonstrate a much broader spectrum of histopathology and diagnostic imaging patterns than does CCD (Wilmshurst et al. 2010; Klein et al. 2011; Mercuri, E. et al. 2010). Associated histopathological findings include multiminicores, centrally placed nuclei, and congenital fiber type disproportion.

RYR1-related minicore myopathy is also known as multiminicore myopathy with external ophthalmoplegia because involvement of extraocular muscles is a nearly universal finding (Jungbluth et al. 2000; Monnier et al. 2009). Clinical signs include severe neonatal hypotonia, respiratory problems, feeding difficulties, frequent respiratory infections, delayed motor development, scoliosis, kyphoscoliosis, contractures, and myopathic facies (Wilmshurst et al. 2010).

RYR1-related centronuclear myopathy is associated with infantile-onset proximal weakness, external ophthalmoplegia, and bulbar involvement, followed by progressive improvement of symptoms. Histopathological findings in muscle biopsied from young patients include internalized nuclei and type 1 fiber predominance. However, when biopsied later in life, two-thirds of patients in one study had central cores or minicores in their muscle (Wilmshurst et al. 2010).

Congenital fiber type disproportion is a genetically heterogeneous disorder defined as hypotrophy of type 1 fibers compared to type 2 fibers. Patients with RYR1-related CFTD exhibit generalized hypotonia, motor delays, recurrent respiratory infections, a weak cry, ptosis, ophthalmoplegia, facial weakness, scoliosis, and contractures (Clarke et al. 2010).

The clinically most severe cases of RYR1-related myopathy present with decreased fetal movement, hypotonia, poor feeding, respiratory involvement, arthrogryposis, and ophthalmoplegia (Bharucha-Goebel et al. 2013). Additional features include fractures or hip dislocation at birth. In the published cohort of patients, the severe neonatal onset cases with dominant acting RYR1 mutations had classic central cores on muscle biopsy, while those with recessive acting RYR1 mutations had notable histologic variability, including fibrosis, variation in fiber size, and nuclear internalization.RYR1 is also involved in exercise-induced rhabdomyolysis and myalgia (Dlamini et al. 2013). Many of the RYR1 mutations identified in these patients are documented to be associated with malignant hyperthermia risk. Patients that span a broad range of ages presenting with symptoms of exercise or heat-induced rhabdomyolysis and sometimes myalgia have been reported (Dlamini et al. 2013).

Malignant hyperthermia susceptibility is another well documented RYR1 disorder.

The RYR1 gene encodes the skeletal muscle isoform of the ryanodine receptor. The RYR1 protein functions to regulate calcium release at the sarcoplasmic reticulum by serving as the calcium-induced, voltage-gated receptor, as well as the channel through which calcium flows. RYR1-related congenital myopathies are inherited as autosomal dominant and recessive disorders. Central core disease is typically dominant, with de novo and inherited mutations being well documented (Klein et al. 2012). The vast majority of CCD-causing mutations result in amino acid substitutions that lie in the C-terminus of the RYR1 protein (Zhou et al. 2007). The other RYR1-related congenital myopathies are most often inherited as autosomal recessive disorders, although exceptions exist. Recessive RYR1 myopathy cases are typically found to have one null allele and a second variant that results in an amino acid substitution or, as in the most severe neonatal forms, a second null allele (Bevilacqua et al. 2011; Monnier et al. 2008).

For this Next Generation Sequencing (NGS) test, sequencing is accomplished by capturing specific regions with an optimized solution-based hybridization kit, followed by massively parallel sequencing of the captured DNA fragments. Additional Sanger sequencing is performed for regions not captured or with insufficient number of sequence reads.

For Sanger sequencing, polymerase chain reaction (PCR) is used to amplify targeted regions. After purification of the PCR products, cycle sequencing is carried out using the ABI Big Dye Terminator v.3.0 kit. PCR products are resolved by electrophoresis on an ABI 3730xl capillary sequencer. In nearly all cases, cycle sequencing is performed separately in both the forward and reverse directions.

Copy number variants (CNVs) are also detected from NGS data. We utilize a CNV calling algorithm that compares mean read depth and distribution for each target in the test sample against multiple matched controls. Neighboring target read depth and distribution and zygosity of any variants within each target region are used to reinforce CNV calls. All CNVs are confirmed using another technology such as aCGH, MLPA, or PCR before they are reported.

This test provides full coverage of all coding exons of the RYR1 gene plus 10 bases of flanking noncoding DNA in all available transcripts along with other non-coding regions in which pathogenic variants have been identified at PreventionGenetics or reported elsewhere. We define full coverage as >20X NGS reads or Sanger sequencing.

Since this test is performed using exome capture probes, a reflex to any of our exome based tests is available (PGxome, PGxome Custom Panels).

Patients that display clinical and histopathological features of the RYR1-related myopathies.

Diseases

For PGxome® we use Next Generation Sequencing (NGS) technologies to cover the coding regions of targeted genes plus 10 bases of flanking non-coding DNA in all available transcripts along with other non-coding regions in which pathogenic variants have been identified at PreventionGenetics or reported elsewhere. As required, genomic DNA is extracted from patient specimens. Patient DNA corresponding to these regions is captured using Agilent Clinical Research Exome hybridization probes. Captured DNA is sequenced on the NovaSeq 6000 using 2x150 bp paired-end reads (Illumina, San Diego, CA, USA). The following quality control metrics are generally achieved: >97% of target bases are covered at >20x, and mean coverage of target bases >120x. Data analysis and interpretation is performed by the internally developed software Titanium-Exome. In brief, the output data from the NovaSeq 6000 is converted to fastqs by Illumina Bcl2Fastq, and mapped by BWA. Variant calls are made by the GATK Haplotype caller and annotated using in house software and SnpEff. Variants are filtered and annotated using VarSeq (www.goldenhelix.com).

For Sanger sequencing, polymerase chain reaction (PCR) is used to amplify targeted regions. After purification of the PCR products, cycle sequencing is carried out using the ABI Big Dye Terminator v.3.0 kit. PCR products are resolved by electrophoresis on an ABI 3730xl capillary sequencer. In nearly all cases, cycle sequencing is performed separately in both the forward and reverse directions.

Copy number variants (CNVs) are also detected from NGS data. We utilize a CNV calling algorithm that compares mean read depth and distribution for each target in the test sample against multiple matched controls. Neighboring target read depth and distribution and zygosity of any variants within each target region are used to reinforce CNV calls. All CNVs are confirmed using another technology such as aCGH, MLPA, or PCR before they are reported.

NextGen Sequencing: As of March 2016, 6.36 Mb of sequence (83 genes, 1557 exons) generated in our lab was compared between Sanger and NextGen methodologies. We detected no differences between the two methods. The comparison involved 6400 total sequence variants (differences from the reference sequences). Of these, 6144 were nucleotide substitutions and 256 were insertions or deletions. About 65% of the variants were heterozygous and 35% homozygous. The insertions and deletions ranged in length from 1 to over 100 nucleotides.

In silico validation of insertions and deletions in 20 replicates of 5 genes was also performed. The validation included insertions and deletions of lengths between 1 and 100 nucleotides. Insertions tested in silico: 2200 between 1 and 5 nucleotides, 625 between 6 and 10 nucleotides, 29 between 11 and 20 nucleotides, 25 between 21 and 49 nucleotides, and 23 at or greater than 50 nucleotides, with the largest at 98 nucleotides. All insertions were detected. Deletions tested in silico: 1813 between 1 and 5 nucleotides, 97 between 6 and 10 nucleotides, 32 between 11 and 20 nucleotides, 20 between 21 and 49 nucleotides, and 39 at or greater than 50 nucleotides, with the largest at 96 nucleotides. All deletions less than 50 nucleotides in length were detected, 13 greater than 50 nucleotides in length were missed. Our standard NextGen sequence variant calling algorithms are generally not capable of detecting insertions (duplications) or heterozygous deletions greater than 100 nucleotides. Large homozygous deletions appear to be detectable.

Copy Number Variant Analysis: The PGxome test detects most larger deletions and duplications including intragenic CNVs and large cytogenetic events; however aberrations in a small percentage of regions may not be accurately detected due to sequence paralogy (e.g., pseudogenes, segmental duplications), sequence properties, deletion/duplication size (e.g., 1-3 exons vs. 4 or more exons), and inadequate coverage. In general, sensitivity for single, double, or triple exon CNVs is ~70% and for CNVs of four exon size or larger is >95%, but may vary from gene-to-gene based on exon size, depth of coverage, and characteristics of the region.

Interpretation of the test results is limited by the information that is currently available. Better interpretation should be possible in the future as more data and knowledge about human genetics and this specific disorder are accumulated.

When sequencing does not reveal any heterozygous differences from the reference sequence, we cannot be certain that we were able to detect both patient alleles.

For technical reasons, the PGxome test is not 100% sensitive. Some exons cannot be efficiently captured, and some genes cannot be accurately sequenced because of the presence of multiple copies in the genome. Therefore, a small fraction of sequence variants will not be detected.

We sequence coding exons for all available transcripts plus 10 bp of flanking non-coding DNA for each exon. We also sequence other regions within or near genes in which pathogenic variants have been identified at PreventionGenetics or reported elsewhere.  Unless specifically indicated, test reports contain no information about other portions of genes.

In most cases, we are unable to determine the phase of sequence variants. In particular, when we find two likely causative mutations for recessive disorders, we cannot be certain that the mutations are on different alleles.

Our ability to detect minor sequence variants due to somatic mosaicism is limited. Sequence variants that are present in less than 50% of the patient's nucleated cells may not be detected.

Runs of mononucleotide repeats (eg (A)n or (T)n) with n >8 in the reference sequence are generally not analyzed because of strand slippage during amplification.

Unless otherwise indicated, DNA sequence data is obtained from a specific cell-type (usually leukocytes if taken from whole blood). Test reports contain no information about the DNA sequence in other cell-types.

We cannot be certain that the reference sequences are correct.

Balanced translocations or inversions are only rarely detected.

Certain types of sex chromosome aneuploidy may not be detected.  

Our ability to detect CNVs due to somatic mosaicism is limited.

We have confidence in our ability to track a specimen once it has been received by PreventionGenetics. However, we take no responsibility for any specimen labeling errors that occur before the sample arrives at PreventionGenetics.

A negative finding does not rule out a genetic diagnosis.

Genetic counseling to help to explain test results to the patients and to discuss reproductive options is recommended.