Advances in circulating biomarker research have led to the use of blood samples to characterize cancer patients’ tumour DNA where a lack of tumour tissue prevents molecular testing. This is critical for non-small cell lung cancer (NSCLC) patients who require tumour molecular characterization in order to access life-extending treatments that would be denied without biopsy. Here we describe a new liquid biopsy diagnostic service for NSCLC patients at the All Wales Medical Genetics Service, Cardiff, UK.
by Dr Angharad Williams, Dr Daniel Nelmes, Helen Roberts and Dr Rachel Butler
Liquid biopsies and cell-free circulating tumour DNA (ctDNA) in clinical practice
The term ‘liquid biopsy’ comes from the sampling of a cancer patient’s tumour DNA from a simple, non-invasive blood test rather than an invasive surgical biopsy. This circulating tumour DNA (ctDNA) is a small fraction of the total cell-free circulating DNA (cfDNA) and consists of short strands of DNA shed by degrading tumour cells directly into a patient’s bloodstream. The levels of ctDNA present will vary greatly based on clinical factors such as proximity of sampling to chemotherapy or radiotherapy, as well as the burden and activity of the tumour [1].
Genetic mutations within the patient’s tumour are detectable at extremely low levels in the ctDNA in the blood [2]. The detection of such mutations provides many potential uses for ctDNA as a biomarker in disease diagnosis and screening, monitoring of therapy response and resistance and detection of minimal residual disease and relapse [3–6].
The many advantages for using ctDNA as a biomarker rest on the fact that ctDNA can be simply extracted from blood; therefore, invasive biopsy procedures can be avoided. Such simple blood sampling is beneficial if the patient is too ill for invasive surgery and is also useful if biopsy-based tumour analysis has failed; thus, unnecessary re-biopsies can be averted. Another benefit of the use of ctDNA over biopsies is that serial blood samples can be taken to replace the need for a re-biopsy to monitor a patient’s response to therapy in ‘real-time’ in the clinic. Practically, blood samples can be arranged, taken and sent for processing at a much faster pace than surgery, gaining valuable time for patients who are in need of urgent cancer-related treatments.
There are, however, potential pitfalls in using ctDNA as a diagnostic biomarker that should be considered prior to setting up a ctDNA-based diagnostic service as well as when interpreting genetic results from ctDNA (summarized in Figure 1). The greatest concern is the fragile nature of cfDNA molecules [7], which means that cfDNA will degrade in a blood sample to undetectable levels the longer that the blood is left unprocessed. Efficient centrifugation and separation of the blood to plasma and storage at −80 °C can be used to halt degradation of cfDNA. In cases where analysis of the cfDNA sample identifies no genetic mutations, this raises the important question of whether the patient was actually shedding ctDNA at the time of the blood sampling or did the ctDNA degrade prior to sample processing? This indicates the unfortunate possibility of false negative results when using ctDNA in the diagnostic setting. Another important factor to consider is that the level of a mutation in the ctDNA, which can quite often be as low as ≤1% mutated ctDNA to wild-type patient cfDNA [8]. Thus, only highly sensitive molecular analysis options should be considered for diagnostic testing strategies using ctDNA.
Molecular analysis of the epidermal growth factor receptor gene in non-small cell lung cancer patients
The epidermal growth factor receptor gene (EGFR) encodes the EGRF protein, a signalling protein that is part of the cellular pathways that control normal cell growth, differentiation and angiogenesis [9]. Approximately 10–20% of ethnically Caucasian non-small cell lung cancer (NSCLC) patients with the adenocarcinoma histological subtype will have a DNA mutation in the EGFR gene, which will activate abnormal constitutive signalling and tumorigenesis [10].
The most common sensitizing EGFR mutations, which represent 85% of known activating EGFR mutations in NSCLC, are the exon 21 point mutation c.2573T>G (p.Leu858Arg) and in-frame deletions in exon 19 [9]. These activating mutations provide a convenient target for first and second generation tyrosine kinase inhibitor (TKI) treatments such as gefitinib (Iressa®, AstraZeneca) [11–13] and act as positive predictive biomarkers for response to these drugs. Traditionally, for patients to access these TKI treatments, tumour biopsy in the form of a formalin-fixed sample is tested for evidence of these activating EGFR mutations at clinical genetic testing centres, such as the All Wales Medical Genetics Service (AWMGS) in Cardiff. However, preservation of the tumour biopsy as formalin-fixed paraffin-embedded (FFPE) tissue leads to a number of issues with genetic analysis including poor quality and yields of DNA (noted in Figure 1). Additionally, a large proportion of NSCLC patients are not well enough to have a biopsy taken and so genetic analysis of tumour DNA and subsequent access to TKI treatments is not possible. This inequity in service provision indicated a clinical need to expand current testing options for NSCLC patients to reach those patients who cannot access TKI-based stratified medicine treatment options. To address this clinical need, a ctDNA-based diagnostic NHS service was developed within AWMGS to detect activating EGFR mutations from patient blood samples in order to alleviate the need for biopsy.
In addition to the availability of first and second generation TKIs, a new third generation TKI, osimertinib (Tagrisso®, AstraZeneca), has recently been made available to a specific group of NSCLC patients. Approximately 50% of patients on first and second generation TKIs will develop an EGFR resistance mutation, c.2369C>T (p.Thr790Met) (commonly known as T790M), leading to disease progression [6]. Since October 2016, osimertinib (Tagrisso®, AstraZeneca), has been available to UK patients shown to harbour the T790M mutant in their tumour via either biopsy or ctDNA analysis through the NHS Cancer Drugs Fund [14]. CtDNA testing has become a popular method of testing for resistance mutations as it mitigates the need for a second invasive biopsy for the patient and, also, serial blood samples can be used to track the patient’s response over a period of time [15].
Establishing the ctDNA-based NSCLC stratified medicine service in the All Wales Medical Genetics Service
Since 2009, the AWMGS has been providing stratified medicine services for NSCLC patients, as well as metastatic colorectal cancer patients, melanoma and gastrointestinal stromal tumour patients in Wales. Though all of these services are based on FFPE tumour analysis, we have developed a wealth of experience in using ctDNA from blood in the field of clinical trials. By 2015, following a number of successful ctDNA-based feasibility studies by laboratory staff and research students, we were confident that we had the knowledge and expertise to bring ctDNA into service, and were one of the first laboratories in the UK to do so.
Owing to the inherent shortcomings of using ctDNA as a biomarker, discussed previously, the following questions were deliberated during validation to find the most appropriate testing methods for the diagnostic service:
- How do we best protect ctDNA in blood during sampling and shipping to the lab, to ensure that the sample that reaches us is faithful to the patient’s real mutation status?
- As important mutations in ctDNA can be at very low levels, how do we ensure we can detect a low enough range of mutations to be clinically relevant to interpretation?
To guarantee sample quality and maintain sufficient levels of ctDNA, we have imposed stringent quality measures on the blood collection and dispatch. The main requirement is that blood samples should be taken in a specialist preservative tube such as CellSave Preservative Tubes (Janssen Diagnostics) or Cell-Free DNA BCT® (Streck) and must reach the laboratory for processing within a strict 96-hour window. This was decided on after discussion with other research groups and internal investigations on the stability of ctDNA in blood and the use of preservative tubes [7].
The sensitivity of the molecular ctDNA assay was paramount in our decision to use the recently developed technology droplet digital polymerase chain reaction (ddPCR) by Bio-Rad (Bio-Rad Laboratories, Inc, California, USA). ddPCR is a highly sensitive fluorescence-based PCR method with an extreme lower limit of detection of 0.0001% of mutant DNA in a wild-type background, which makes it the superior choice over other technologies such as next-generation sequencing and quantitative PCR (qPCR). Practically, in the service, we have detected EGFR mutations in patient ctDNA at an abundance as low as 0.7%.
The AWMGS now provides ctDNA-based testing of the EGFR gene in NSCLC patients at both first-line testing for sensitizing mutations and for resistance mutation testing on patient progression on TKIs (Figure 2). The service launched across Wales in April 2016 and has since been expanded to provide testing for certain centres in the South West of England with funding from AstraZeneca. A year on, over 100 patients have been tested, the majority (approximately 60%) of patient referrals have been for T790M progression testing to avoid repeat biopsies for patients. Six patients, for whom TKIs were previously inaccessible due to failed FFPE-based testing or inability to biopsy, were successfully tested through the ctDNA service and are now receiving first-line EGFR TKI therapy following the detection of activating EGFR mutations in ctDNA.
Ongoing and future developments
The field of liquid biopsies is steadily gaining pace in the UK and abroad with a number of centres now providing EGFR ctDNA testing. New circulating biomarkers, such as exosomes and circulating tumour cells, are coming through from translation research and have vast potential in the field of stratified medicine. At the AWMGS, we aim to expand our current liquid biopsy testing in the near future with targets for both metastatic colorectal cancer and metastatic melanoma.
References
1. Schwarzenbach H, Hoon DS, Pantel K. Cell-free nucleic acids as biomarkers in cancer patients. Nat Rev Cancer 2011; 11: 426–437.
2. Bettegowda C, Sausen M, Leary RJ, Kinde I, Wang Y, Agrawal N, Bartlett BR, Wang H, Luber B, et al. Detection of circulating tumor DNA in early- and late-stage human malignancies. Sci Transl Med 2014; 6: 224ra24.
3. Chen KZ, Lou F, Yang F, Zhang JB, Ye H, Chen W, Guan T, Zhao MY, Su XX, et al. Circulating tumor DNA detection in early-stage non-small cell lung cancer patients by targeted sequencing. Sci Rep 2016; 6: 31985.
4. Dawson S-J, Tsui DWY, Murtaza M, Biggs H, Rueda OM, Chin SF, Dunning MJ, Gale D, Forshew T, et al. Analysis of circulating tumor DNA to monitor metastatic breast cancer. N Engl J Med 2013; 368: 1199–1209.
5. Forshew T, Murtaza M, Parkinson C, Gale D, Tsui DW, Kaper F, Dawson SJ, Piskorz AM, Jimenez-Linan M, et al. Noninvasive identification and monitoring of cancer mutations by targeted deep sequencing of plasma DNA. Sci Transl Med 2012; 4: 136ra68.
6. Murtaza M, Dawson SJ, Tsui DW, Gale D, Forshew T, Piskorz AM, Parkinson C, Chin SF, Kingsbury Z, et al. Non-invasive analysis of acquired resistance to cancer therapy by sequencing of plasma DNA. Nature 2013; 497: 108–112.
7. Rothwell DG, Smith N, Morris D, Leong HS, Li Y, Hollebecque A, Ayub M, Carter L, Antonello J, et al. Genetic profiling of tumours using both circulating free DNA and circulating tumour cells isolated from the same preserved whole blood sample. Mol Oncol 2016; 10: 566–574.
8. Heitzer E, Ulz P, Geigl JB. Circulating tumor DNA as a liquid biopsy for cancer. Clin Chem 2015; 61: 112–123.
9. Jänne PA, Engelman JA, and Johnson BE. Epidermal growth factor receptor mutations in non–small-cell lung cancer: implications for treatment and tumor biology. J Clin Onc 2005; 23: 3227–3234.
10. Li T, Kung H-J, Mack PC, Gandara DR. Genotyping and genomic profiling of non-small-cell lung cancer: implications for current and future therapies. J Clin Onc 2013; 31: 1039–1049.
11. Douillard JY, Ostoros G, Cobo M, Ciuleanu T, Cole R, McWalter G, Walker J, Dearden S, Webster A, et al. Gefitinib treatment in EGFR mutated Caucasian NSCLC: circulating-free tumor DNA as a surrogate for determination of EGFR status. J Thorac Oncol 2014; 9: 1345–1353.
12. Goto K, Ichinose Y, Ohe Y, Yamamoto N, Negoro S, Nishio K, Itoh Y, Jiang H, Duffield E, et al. Epidermal growth factor receptor mutation status in circulating free DNA in serum: from IPASS, a phase III study of gefitinib or carboplatin/paclitaxel in non-small cell lung cancer. J Thorac Oncol 2012; 7: 115–121.
13. National Institute for Health and Care Excellence (NICE). Gefitinib for the first-line treatment of locally advanced or metastatic non-small-cell lung cancer. Technology appraisal guidance [TA192] 2010. [https: //www.nice.org.uk/guidance/ta192]
14. NICE. Osimertinib for treating locally advanced or metastatic EGFR T790M mutation-positive non-small-cell lung cancer. Final appraisal determination [TA10022] 2016. [https: //www.nice.org.uk/guidance/GID-TA10022/documents/final-appraisal-determination-document]
15. Sundaresan TK, Sequist LV, Heymach JV, Riely GJ, Jänne PA, Koch WH, Sullivan JP, Fox DB, Maher R, et al. Detection of T790M, the acquired resistance EGFR mutation, by tumor biopsy versus noninvasive blood-based analyses. Clin Cancer Res 2016; 22: 1103–1110.
The authors
Angharad Williams1 PhD, Daniel Nelmes2,3 PhD, Helen Roberts1 BSc and Rachel Butler*1 FRCPath
1All Wales Medical Genetics Service,
NHS Wales, The Institute of Medical
Genetics, Cardiff and Vale University LHB,
University Hospital of Wales, Cardiff
CF14 4XW, Wales, UK
2School of Medicine, Cardiff University, Cardiff CF14 4XN, Wales, UK
3Velindre Cancer Centre, Cardiff
CF14 2TL, Wales, UK
*Corresponding author
E-mail: Rachel.Butler@wales.nhs.uk
ReaScan CXCL13 – the world’s fastest tool
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, /in Featured Articles /by 3wmediaTowards meeting the global requirement for safe blood
, /in Featured Articles /by 3wmediaAccording to the WHO, an estimated 2 % of the world’s population needs to regularly donate blood to ensure that supply meets demand. Currently approximately 85 million units of red blood cells, the most frequently transfused blood product, are provided per annum globally. Over half the recipients, predominantly in the less developed countries, are children with severe anemia and women suffering from peri-partum hemorrhage. The major problem here is the serious shortage of suitable blood donors: WHO data reveal that in 75 such countries the supply of safe blood is inadequate, leading to medically avoidable maternal and child mortality. In high income countries, however, around 70 % of blood transfusions are given for surgical reasons, particularly to support cardiac, cancer and transplantation patients. Whilst in these countries the blood supply is currently maintained at an adequate level (though the ageing population will inevitably affect this), there is still a small, but crucially not zero, risk associated with blood transfusion.
Donors in the West, however, are carefully screened, and blood is comprehensively tested for transfusion-transmitted infections. Leucocytes, known to harbour infectious agents and to have potentially adverse effects on recipients’ immune systems, are depleted, which can remove 99.995% of the approximately two billion white cells present in a 500 mL unit of blood. Why then is there still a risk? The problem is that stored blood, usually kept for up to five weeks at around 4 °C, deteriorates over time. The residual white cells cause components such as histamine, eosinophil cationic protein and eosinophil protein X to be released into the supernatant fluid, which inhibit neutrophil function and thus impair the immune system of the recipient. Older red cells are also less able to deform and unload oxygen; capillaries can become obstructed leading to tissue ischemia.
As the development of a robust infrastructure for the collection and storage of safe blood in the less developed countries remains an ongoing project, and in the West lowering the storage time for blood is unworkable, is there a solution for the global shortage of safe blood for transfusion? A joint project involving research workers in the UK, Thailand and Japan has demonstrated a feasible approach via the generation of immortalized adult erythroid progenitor cell lines. These allow an unlimited supply of red cells to be produced with minimal culture requirements. In future such technology could not only make transfusion in the West risk-free but might provide a solution for areas of the world with inadequate supplies of safe blood.
Liquid biopsy for diagnostic epidermal growth factor receptor gene testing in non-small cell lung cancer
, /in Featured Articles /by 3wmediaAdvances in circulating biomarker research have led to the use of blood samples to characterize cancer patients’ tumour DNA where a lack of tumour tissue prevents molecular testing. This is critical for non-small cell lung cancer (NSCLC) patients who require tumour molecular characterization in order to access life-extending treatments that would be denied without biopsy. Here we describe a new liquid biopsy diagnostic service for NSCLC patients at the All Wales Medical Genetics Service, Cardiff, UK.
by Dr Angharad Williams, Dr Daniel Nelmes, Helen Roberts and Dr Rachel Butler
Liquid biopsies and cell-free circulating tumour DNA (ctDNA) in clinical practice
The term ‘liquid biopsy’ comes from the sampling of a cancer patient’s tumour DNA from a simple, non-invasive blood test rather than an invasive surgical biopsy. This circulating tumour DNA (ctDNA) is a small fraction of the total cell-free circulating DNA (cfDNA) and consists of short strands of DNA shed by degrading tumour cells directly into a patient’s bloodstream. The levels of ctDNA present will vary greatly based on clinical factors such as proximity of sampling to chemotherapy or radiotherapy, as well as the burden and activity of the tumour [1].
Genetic mutations within the patient’s tumour are detectable at extremely low levels in the ctDNA in the blood [2]. The detection of such mutations provides many potential uses for ctDNA as a biomarker in disease diagnosis and screening, monitoring of therapy response and resistance and detection of minimal residual disease and relapse [3–6].
The many advantages for using ctDNA as a biomarker rest on the fact that ctDNA can be simply extracted from blood; therefore, invasive biopsy procedures can be avoided. Such simple blood sampling is beneficial if the patient is too ill for invasive surgery and is also useful if biopsy-based tumour analysis has failed; thus, unnecessary re-biopsies can be averted. Another benefit of the use of ctDNA over biopsies is that serial blood samples can be taken to replace the need for a re-biopsy to monitor a patient’s response to therapy in ‘real-time’ in the clinic. Practically, blood samples can be arranged, taken and sent for processing at a much faster pace than surgery, gaining valuable time for patients who are in need of urgent cancer-related treatments.
There are, however, potential pitfalls in using ctDNA as a diagnostic biomarker that should be considered prior to setting up a ctDNA-based diagnostic service as well as when interpreting genetic results from ctDNA (summarized in Figure 1). The greatest concern is the fragile nature of cfDNA molecules [7], which means that cfDNA will degrade in a blood sample to undetectable levels the longer that the blood is left unprocessed. Efficient centrifugation and separation of the blood to plasma and storage at −80 °C can be used to halt degradation of cfDNA. In cases where analysis of the cfDNA sample identifies no genetic mutations, this raises the important question of whether the patient was actually shedding ctDNA at the time of the blood sampling or did the ctDNA degrade prior to sample processing? This indicates the unfortunate possibility of false negative results when using ctDNA in the diagnostic setting. Another important factor to consider is that the level of a mutation in the ctDNA, which can quite often be as low as ≤1% mutated ctDNA to wild-type patient cfDNA [8]. Thus, only highly sensitive molecular analysis options should be considered for diagnostic testing strategies using ctDNA.
Molecular analysis of the epidermal growth factor receptor gene in non-small cell lung cancer patients
The epidermal growth factor receptor gene (EGFR) encodes the EGRF protein, a signalling protein that is part of the cellular pathways that control normal cell growth, differentiation and angiogenesis [9]. Approximately 10–20% of ethnically Caucasian non-small cell lung cancer (NSCLC) patients with the adenocarcinoma histological subtype will have a DNA mutation in the EGFR gene, which will activate abnormal constitutive signalling and tumorigenesis [10].
The most common sensitizing EGFR mutations, which represent 85% of known activating EGFR mutations in NSCLC, are the exon 21 point mutation c.2573T>G (p.Leu858Arg) and in-frame deletions in exon 19 [9]. These activating mutations provide a convenient target for first and second generation tyrosine kinase inhibitor (TKI) treatments such as gefitinib (Iressa®, AstraZeneca) [11–13] and act as positive predictive biomarkers for response to these drugs. Traditionally, for patients to access these TKI treatments, tumour biopsy in the form of a formalin-fixed sample is tested for evidence of these activating EGFR mutations at clinical genetic testing centres, such as the All Wales Medical Genetics Service (AWMGS) in Cardiff. However, preservation of the tumour biopsy as formalin-fixed paraffin-embedded (FFPE) tissue leads to a number of issues with genetic analysis including poor quality and yields of DNA (noted in Figure 1). Additionally, a large proportion of NSCLC patients are not well enough to have a biopsy taken and so genetic analysis of tumour DNA and subsequent access to TKI treatments is not possible. This inequity in service provision indicated a clinical need to expand current testing options for NSCLC patients to reach those patients who cannot access TKI-based stratified medicine treatment options. To address this clinical need, a ctDNA-based diagnostic NHS service was developed within AWMGS to detect activating EGFR mutations from patient blood samples in order to alleviate the need for biopsy.
In addition to the availability of first and second generation TKIs, a new third generation TKI, osimertinib (Tagrisso®, AstraZeneca), has recently been made available to a specific group of NSCLC patients. Approximately 50% of patients on first and second generation TKIs will develop an EGFR resistance mutation, c.2369C>T (p.Thr790Met) (commonly known as T790M), leading to disease progression [6]. Since October 2016, osimertinib (Tagrisso®, AstraZeneca), has been available to UK patients shown to harbour the T790M mutant in their tumour via either biopsy or ctDNA analysis through the NHS Cancer Drugs Fund [14]. CtDNA testing has become a popular method of testing for resistance mutations as it mitigates the need for a second invasive biopsy for the patient and, also, serial blood samples can be used to track the patient’s response over a period of time [15].
Establishing the ctDNA-based NSCLC stratified medicine service in the All Wales Medical Genetics Service
Since 2009, the AWMGS has been providing stratified medicine services for NSCLC patients, as well as metastatic colorectal cancer patients, melanoma and gastrointestinal stromal tumour patients in Wales. Though all of these services are based on FFPE tumour analysis, we have developed a wealth of experience in using ctDNA from blood in the field of clinical trials. By 2015, following a number of successful ctDNA-based feasibility studies by laboratory staff and research students, we were confident that we had the knowledge and expertise to bring ctDNA into service, and were one of the first laboratories in the UK to do so.
Owing to the inherent shortcomings of using ctDNA as a biomarker, discussed previously, the following questions were deliberated during validation to find the most appropriate testing methods for the diagnostic service:
To guarantee sample quality and maintain sufficient levels of ctDNA, we have imposed stringent quality measures on the blood collection and dispatch. The main requirement is that blood samples should be taken in a specialist preservative tube such as CellSave Preservative Tubes (Janssen Diagnostics) or Cell-Free DNA BCT® (Streck) and must reach the laboratory for processing within a strict 96-hour window. This was decided on after discussion with other research groups and internal investigations on the stability of ctDNA in blood and the use of preservative tubes [7].
The sensitivity of the molecular ctDNA assay was paramount in our decision to use the recently developed technology droplet digital polymerase chain reaction (ddPCR) by Bio-Rad (Bio-Rad Laboratories, Inc, California, USA). ddPCR is a highly sensitive fluorescence-based PCR method with an extreme lower limit of detection of 0.0001% of mutant DNA in a wild-type background, which makes it the superior choice over other technologies such as next-generation sequencing and quantitative PCR (qPCR). Practically, in the service, we have detected EGFR mutations in patient ctDNA at an abundance as low as 0.7%.
The AWMGS now provides ctDNA-based testing of the EGFR gene in NSCLC patients at both first-line testing for sensitizing mutations and for resistance mutation testing on patient progression on TKIs (Figure 2). The service launched across Wales in April 2016 and has since been expanded to provide testing for certain centres in the South West of England with funding from AstraZeneca. A year on, over 100 patients have been tested, the majority (approximately 60%) of patient referrals have been for T790M progression testing to avoid repeat biopsies for patients. Six patients, for whom TKIs were previously inaccessible due to failed FFPE-based testing or inability to biopsy, were successfully tested through the ctDNA service and are now receiving first-line EGFR TKI therapy following the detection of activating EGFR mutations in ctDNA.
Ongoing and future developments
The field of liquid biopsies is steadily gaining pace in the UK and abroad with a number of centres now providing EGFR ctDNA testing. New circulating biomarkers, such as exosomes and circulating tumour cells, are coming through from translation research and have vast potential in the field of stratified medicine. At the AWMGS, we aim to expand our current liquid biopsy testing in the near future with targets for both metastatic colorectal cancer and metastatic melanoma.
References
1. Schwarzenbach H, Hoon DS, Pantel K. Cell-free nucleic acids as biomarkers in cancer patients. Nat Rev Cancer 2011; 11: 426–437.
2. Bettegowda C, Sausen M, Leary RJ, Kinde I, Wang Y, Agrawal N, Bartlett BR, Wang H, Luber B, et al. Detection of circulating tumor DNA in early- and late-stage human malignancies. Sci Transl Med 2014; 6: 224ra24.
3. Chen KZ, Lou F, Yang F, Zhang JB, Ye H, Chen W, Guan T, Zhao MY, Su XX, et al. Circulating tumor DNA detection in early-stage non-small cell lung cancer patients by targeted sequencing. Sci Rep 2016; 6: 31985.
4. Dawson S-J, Tsui DWY, Murtaza M, Biggs H, Rueda OM, Chin SF, Dunning MJ, Gale D, Forshew T, et al. Analysis of circulating tumor DNA to monitor metastatic breast cancer. N Engl J Med 2013; 368: 1199–1209.
5. Forshew T, Murtaza M, Parkinson C, Gale D, Tsui DW, Kaper F, Dawson SJ, Piskorz AM, Jimenez-Linan M, et al. Noninvasive identification and monitoring of cancer mutations by targeted deep sequencing of plasma DNA. Sci Transl Med 2012; 4: 136ra68.
6. Murtaza M, Dawson SJ, Tsui DW, Gale D, Forshew T, Piskorz AM, Parkinson C, Chin SF, Kingsbury Z, et al. Non-invasive analysis of acquired resistance to cancer therapy by sequencing of plasma DNA. Nature 2013; 497: 108–112.
7. Rothwell DG, Smith N, Morris D, Leong HS, Li Y, Hollebecque A, Ayub M, Carter L, Antonello J, et al. Genetic profiling of tumours using both circulating free DNA and circulating tumour cells isolated from the same preserved whole blood sample. Mol Oncol 2016; 10: 566–574.
8. Heitzer E, Ulz P, Geigl JB. Circulating tumor DNA as a liquid biopsy for cancer. Clin Chem 2015; 61: 112–123.
9. Jänne PA, Engelman JA, and Johnson BE. Epidermal growth factor receptor mutations in non–small-cell lung cancer: implications for treatment and tumor biology. J Clin Onc 2005; 23: 3227–3234.
10. Li T, Kung H-J, Mack PC, Gandara DR. Genotyping and genomic profiling of non-small-cell lung cancer: implications for current and future therapies. J Clin Onc 2013; 31: 1039–1049.
11. Douillard JY, Ostoros G, Cobo M, Ciuleanu T, Cole R, McWalter G, Walker J, Dearden S, Webster A, et al. Gefitinib treatment in EGFR mutated Caucasian NSCLC: circulating-free tumor DNA as a surrogate for determination of EGFR status. J Thorac Oncol 2014; 9: 1345–1353.
12. Goto K, Ichinose Y, Ohe Y, Yamamoto N, Negoro S, Nishio K, Itoh Y, Jiang H, Duffield E, et al. Epidermal growth factor receptor mutation status in circulating free DNA in serum: from IPASS, a phase III study of gefitinib or carboplatin/paclitaxel in non-small cell lung cancer. J Thorac Oncol 2012; 7: 115–121.
13. National Institute for Health and Care Excellence (NICE). Gefitinib for the first-line treatment of locally advanced or metastatic non-small-cell lung cancer. Technology appraisal guidance [TA192] 2010. [https: //www.nice.org.uk/guidance/ta192]
14. NICE. Osimertinib for treating locally advanced or metastatic EGFR T790M mutation-positive non-small-cell lung cancer. Final appraisal determination [TA10022] 2016. [https: //www.nice.org.uk/guidance/GID-TA10022/documents/final-appraisal-determination-document]
15. Sundaresan TK, Sequist LV, Heymach JV, Riely GJ, Jänne PA, Koch WH, Sullivan JP, Fox DB, Maher R, et al. Detection of T790M, the acquired resistance EGFR mutation, by tumor biopsy versus noninvasive blood-based analyses. Clin Cancer Res 2016; 22: 1103–1110.
The authors
Angharad Williams1 PhD, Daniel Nelmes2,3 PhD, Helen Roberts1 BSc and Rachel Butler*1 FRCPath
1All Wales Medical Genetics Service,
NHS Wales, The Institute of Medical
Genetics, Cardiff and Vale University LHB,
University Hospital of Wales, Cardiff
CF14 4XW, Wales, UK
2School of Medicine, Cardiff University, Cardiff CF14 4XN, Wales, UK
3Velindre Cancer Centre, Cardiff
CF14 2TL, Wales, UK
*Corresponding author
E-mail: Rachel.Butler@wales.nhs.uk
Diagnosis of the intestinal parasite Strongyloides stercoralis by detection of cell-free parasite DNA fragments in urine
, /in Featured Articles /by 3wmediaDiagnosis of infection with the parasitic roundworm Strongyloides stercoralis is currently done by stool sample culture to detect active larvae. However, the sensitivity of this method can be as low as 28%. This article describes how cell-free parasite DNA can be detected in urine when the results of stool-sample testing are negative.
by Dr Clive Shiff and Dr Alejandro Krolewiecki
Introduction
Of the neglected tropical diseases, Strongyloides stercoralis infection has emerged as a global problem because it is difficult to diagnose and is often silent but long-lived [1]. Currently the definitive test is performed by examining or making a culture of fresh stool to detect active larvae. Serological analysis for specific antibodies is also used but this is far from definitive [2]. This intestinal parasite has an unusual life cycle which is still somewhat enigmatic and can have dire implications if the patient becomes immunosuppressed [3]. This can happen as a patient ages but also if, for some reason, is placed on immunosuppression therapy. To put this in perspective it is important to appreciate the complexity of this infection and the importance of a simple effective diagnostic test.
The infection is caused by parthenogenetic females that live in the upper reaches of the intestinal tract. Unlike hookworms, which also have adults in the gut, Str. stercoralis does not lay eggs which exit in the stool, embryonate and hatch outside the body, Str. stercoralis females incubate the eggs and deposit the eggs in the intestinal mucosa from which the first-stage rhabditiform larvae emerge in large numbers. Some larvae pass in the stool moult, and commence a free-living sexual phase with male and female adults living in the human fecal material. Free-living stages are not parasitic but after several cycles the larvae change from producing the benign, rhabditiform stage to a parasitic filariform stage. These are infective and will penetrate the skin of anyone approaching or contacting the fecal mass. The parasitic stage occurs after the second moult producing the stage called ‘L3’. These larvae secrete proteolytic enzymes and are tissue invasive. However, not all the larvae produced by the parthenogenetic gut parasites are voided in the stool. A proportion of these larvae moult internally and commence the autoinfection stage. They reinvade the host mucosa and reinfect the host and are distributed round the body in the blood and other fluids. In immunocompetent persons these larvae are killed off and their matter is finally excreted through the urine [3]. In people who become immunosuppressed, these larvae continue to survive and accumulate in large numbers and constitute an urgent, life-threatening condition.
Detecting species-specific DNA from urine
Cell-free DNA of parasite origin has been detected in urine of patients with a number of blood-borne and tissue-dwelling parasites. This has been shown with malaria [4], urogenital schistosomes [5], Schistosoma mansoni [6] and others. In all these publications detection of DNA from urine was the most sensitive of serology, parasitological examination of excreta or antigen capture test and the specificity was equal to detection of eggs in excreta [7]. There is also an advantage in using urine specimens. It is simple and can be collected almost on demand. For this work the specimen is filtered through a standard filter paper cone. Approximately 40 ml of urine is filtered, and then the paper is removed from the beaker, opened and allowed to dry in a fly-proof, clean area [8]. When dry each filter is placed in a sealable zip lock plastic bag with a small desiccant capsule. Papers can be stored at 4 °C for months without deterioration of the parasite DNA. In the field when survey work is carried out, urine collection can be carried out simply and in a single day, but filtration and drying of the filters needs to be done within 3 to 4 hours of collection as DNA is degraded by long storage in the urine specimen.
Methods
Ethical clearances
The specimens were collected as part of an ongoing programme to find and cure infections of soil-transmitted helminths by the Ministry of Health and approved by Commité de Ética Colegio Médico de Salta, Salta, Argentina and Johns Hopkins University (IRB number 6199).
Extraction of parasite DNA from filter paper
Filter papers (Whatman No. 3, 12.5 cm diameter) clearly labelled with pencil received in the laboratory are processed as follows. Using a metal punch fifteen 1.00 mm discs are removed from the apex of the quadrant sampled. These are placed in a sterile 1.5 mL Eppendorf tube and 600 µL of nuclease free water added, then incubated at 95 °C for 10 min, and subject to gentle agitation overnight at room temperature. Tubes were then centrifuged at 4000 r.p.m. for 5 min and the supernatant was removed and processed for DNA extraction. We used QIAmpDNA Blood Mini Kit (Qiagen) according to manufacturer’s protocol. The amount of recovered DNA was measured by NanoDrop, ND-1000 spectrophotometer (Thermo Scientific) and stored at −20 °C [9].
Identification of specific Str. stercoralis DNA fragment
Previous work [10] has shown that tandem repeat DNA composed a high proportion of genomic DNA, and these repeats incorporate smaller repeat fragments of DNA. Small fragments of parasite-specific DNA, are found nested within tandem repeats. GenBank AY028262 is such a fragment. Primers for a 125-bp fragment were designed using PrimerQuest Tool (IDT) these are:
Forward (SSC-F) 5´-CTC AGC TCC AGT AAA GCA ACA G-3´
Reverse (SSC-R) 5´-AGC TGA ATC TGG AGA GTG AAG A-3´.
The sequence amplified by these primers was compared with a Blast search against total GenBank data and found only to amplify Str. stercoralis DNA. They were also tested against DNA from three Ancylostoma spp., Sch. mansoni and Sch. haematobium and found only to amplify a product from Str. stercoralis [9].
Amplification and visualization
PCR amplification in 15-µL volume with 2× Taq Mastermix (New England Biolabs), 0.75 µL of 10 µM of each primer, 1–2 µL (20–100 ng/µL) of product DNA made to volume with PCR-grade water (Sigma-Aldrich). The protocol, denaturation at 95 °C to 10 min and 35 cycles at 95 C for 1 min, 63 °C for 1 min 30s, 72 °C for 1 min and a final extension at 72 °C for 10 min. To confirm amplicon size products, were resolved on a 2% agarose gel and stained with Ethidium Bromide (Sigma-Aldrich) [9].
Results
Limits of detection
Genomic DNA from Str. stercoralis was diluted and titrated sequentially in concentration from 2 ng/µL to 2 fg/µL to determine the extinction level under standard amplification procedure. Amplifications were performed in duplicate to ensure reproducibility. Products amplified were cleaned with ExoSAP-IT (Afflymetrix Inc.), sequenced and compared with the Str. stercoralis repeat sequence in GenBank (AY028262) to ensure confirmation. In Figure 1 the limit of detection was 20 pg of target DNA.
Diagnostic efficacy
A study was conducted to compare the diagnostic efficacy of parasitological copro-diagnostic methods with DNA detection. For this specimens of stool and urine were collected from 125 individuals living in endemic regions of northern Argentina. The stool specimens were examined fresh using three parasitological tests, concentration- sedimentation, Harada-Mori and Baermann culture methods. Urine samples were filtered as outlined above, dried and sent to the laboratory at Johns Hopkins for DNA extraction and amplification.
The results are given in Table 1 comparing the results of stool versus urine analysis. The prevalence when stool only, 28% is compared with the DNA detection 44.8%, the difference in prevalence is a highly significant 62% difference (P=0.0058). With further analysis comparing the two procedures in the same community, detection of DNA in the urine is more sensitive with significant difference again, 87.5% (95% CL 76.8–94.4) against 56.5% (CL 42.3–69.0%). Specificity in both tests was 100%.
Discussion
There are important reasons for the development of highly sensitive and specific diagnostic tests for the neglected tropical diseases. These relate to modern attempts to limit or eliminate these diseases from much of the endemic areas [11]. However, most parasitic infections have been sustained in their communities for evolutionary time and the parasites have adapted effectively to sustain their populations. This has resulted in very high replicative stages in the life cycle, for instance schistosomes produce large and sustained numbers of cercariae in the snail intermediate hosts from a single miracidium [12]. With strongyloidiasis the multiplication occurs in the host through effective autoinfection; hence, effective control must identify all cases to eliminate the condition. In this work the difference between a prevalence of 28% and 44.8%, means missing almost half of the population at risk. Serological diagnoses are available, but authorities are not satisfied with either the specificity or sensitivity of these tests [13].
The work described there opens an avenue to help ameliorate these problems on two counts. First there is an improvement in sensitivity without loss of specificity, albeit the process requires the use of DNA amplification and detection equipment. This has been mentioned in numerous review articles, but in reality it is an excuse rather than a reason because there are few countries in the world now where there is no access to such equipment. Furthermore the ‘loop mediated amplification procedure’ (LAMP) has been applied to most of these diagnostic methods with success, so amplification is not a real problem. The main difficulty has been in collecting and storing specimens. This has been solved with the use of urine as a vehicle for diagnostic DNA. There are two reasons. First, urine can be obtained on demand; there is no need for a long wait. Second, the specimen is easily collected: the procedure is non-invasive and with simple equipment the sample can be filtered through standard Whatman No. 3 filter paper within minutes of collection. The collection of urine samples for DNA testing has already been done in Nigeria [8] and elsewhere, where colleagues have implemented the work.
Several laboratories are focusing on stool collections as so many soil-transmitted helminths are transmitted by feces. In a hospital environment, collection of a stool sample is a straightforward procedure that can be carried out under clean and safe-handling conditions. DNA detection can be carried out on preserved feces, and using real-time PCR multiplex procedures DNA from various sources (parasitic) can be identified from a single sample and the procedure is currently in use [14]. Although there is an advantage in multiple diagnoses from a single stool, the sensitivity will depend on whether there are actual organisms in the stool examined. In low-density infections, there are times when there is no parasite material in the feces, which will give a false negative response [15]. It has been shown with Sch. mansoni infections, DNA was detected in urine when there were no eggs of the parasite seen in stool [6].
Conclusions
Although this method may not be feasible for all soil-transmitted helminths, detection of parasite-specific DNA in urine seems the best way of achieving optimum sensitivity. The use of urine also has added advantages over stool collection, primarily because it is available more or less on demand, it is simple to handle, does not require fume extraction hoods, it is not dangerous to handle and can be processed in the field, and once collected on dry filter paper it is easily and economically transported.
References
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2. Krolewiecki AJ, Ramanathan R, Fink V, McAuliffe I, Cajal SP, Won K, Juarez M, Di Paolo A, Tapia L, et al. Improved diagnosis of Strongyloides stercoralis using recombinant antigen-based serologies in a community-wide study in northern Argentina. Clin Vaccine Immunol 2010; 17(10): 1624–1630.
3. Schad G, Aikens L, Smith G. Strongyloides stercoralis: is there a canonical migratory route through the host? J Parasitol 1989; 75: 740–749.
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6. Lodh N, Mwansa JC, Mutengo MM, Shiff CJ. Diagnosis of Schistosoma mansoni without the stool: comparison of three diagnostic tests to detect Schistosoma mansoni infection from filtered urine in Zambia. Am J Trop Med Hyg 2013 July; 89(1): 46–50.
7. Krolewiecki AJ, Lammie P, Jacobson J, Gabrielli AF, Levecke B, Socias E, Arias LM, Sosa N, Abraham D, et al. A public health response against Strongyloides stercoralis: time to look at soil-transmitted helminthiasis in full. PLoS Negl Trop Dis 2013; 7(5): e2165.
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11. Lo NC, Addiss DG, Hotez PJ, King CH, Stothard R, Evans DS, Colley DG, Lin W, Coulibaly JT, et al. A call to strengthen the global strategy against schistosomiasis and soil-transmitted helminthiasis: the time is now. Lancet Infect Dis 2016.
12. Shiff C. The importance of definitive diagnosis in chronic schistosomiasis, with reference to Schistosoma haematobium. J Parasitol Res 2012; 2012: 761269.
13. Bisoffi Z, Buonfrate D, Sequi M, Mejia R, Cimino RO, Krolewiecki AJ, Albonico M, Gobbo M, Bonafini S, et al. Diagnostic accuracy of five serologic tests for Strongyloides stercoralis infection. PLoS Negl Trop Dis 2014; 8(1): e2640.
14. Basuni M, Muhi J, Othman N, Verweij JJ, Ahmad M, Miswan N, Rahumatullah A, Aziz FA, Zainudin NS, Noordin R. A pentaplex real-time polymerase chain reaction assay for detection of four species of soil-transmitted helminths. Am J Trop Med Hyg 2011; 84(2): 338–343.
15. Lodh N, Naples JM, Bosompem KM, Quartey J, Shiff CJ. Detection of parasite-specific DNA in urine sediment obtained by filtration differentiates between single and mixed infections of Schistosoma mansoni and S. haematobium from endemic areas in Ghana. PLoS One 2014; 9: e91144.
The authors
Clive Shiff*1 PhD and Alejandro Krolewiecki2 MD, PhD
1Department of Molecular
Microbiology and Immunology,
Johns Hopkins Bloomberg
School of Public Health,
Baltimore, MD 21205, USA
2Instituto de Investigaciones en
Enfermedades Tropicales,
San Ramón de la Nueva Orán 4530,
Salta, Argentina
*Corresponding author
E-mail: cshiff1@jhu.edu
Cascade screening of relatives for familiar hypercholesterolemia: detection of low density lipoprotein receptor gene mutations using real-time PCR
, /in Featured Articles /by 3wmediaEarly detection of disease-associated mutations in patients with familial hypercholesterolemia (FH) is crucial for early interventions that can reduce the risk of cardiovascular disease. Here, we describe real-time PCR-based approaches for the rapid detection of single nucleotide substitutions or insertions of the low density lipoprotein receptor gene for cascade screening of relatives.
by Sarojini Pandey and Dimitris K. Grammatopoulos
Introduction
Familial hypercholesterolemia (FH) 5 (OMIM#606945) is an autosomal-dominant disorder associated with abnormally high serum concentrations of low density lipoprotein (LDL) cholesterol (LDL-C) [1]. FH is one of the most common inherited disorders, with a worldwide prevalence estimated at 1 in 200–500 [2]. Affected individuals have increased risk of premature coronary heart disease and death [3]; however, most remain undiagnosed, untreated or inadequately treated. It has been proven that early detection of the disease and treatment reduces morbidity and mortality [4]. The majority of FH cases are caused by genetic defects in the LDL receptor (LDLR) as well as apolipoprotein B, or proprotein convertase subtilisin/kexin type 9. More than 80% of FH patients have mutations in the LDLR gene [5]. Over 1400 different mutations are listed in the LDLR gene database of University College London to date.
To address the screening deficit, the National Institute for Health and Clinical Excellence (NICE) in the United Kingdom developed guidelines on FH management strongly recommending identification of causal mutations in suspected cases of FH phenotype and cascade screening of relatives using a combination of genetic testing and LDL-C concentration measurement to identify affected relatives of those index individuals with a clinical diagnosis of FH [6]. This approach of genetic testing of affected individuals and screening of relatives is considered the most cost-effective strategy for detecting cases of FH across the population [7]. However, the most appropriate and cost-effective diagnostic testing protocol for use across the FH clinical diagnostic services remains to be established. Here, we describe an experimental approach suitable for the rapid detection of known single nucleotide substitutions or insertions of the LDLR gene in suspected individuals using real-time based PCR.
Real-time PCR-based method for identifying LDLR gene mutations
Genomic DNA was extracted from saliva or EDTA-containing blood samples using a QIAamp DNA Blood Mini Kit (Qiagen), and DNA concentration was quantified by ND-1000 spectrophotometer (NanoDrop, Thermo Scientific).
Genomic DNA was amplified with specific oligonucleotide primers and fluorescently labelled probes to identify the PCR product (LC FastStart DNA Master Hybridization Probe kit, Roche). The specific genotype was determined by performing a melting-curve analysis based on fluorescence resonance energy transfer (FRET) technique. Each 10-μL reaction contained 1× LightCycler FastStart DNA Master HybProbe, 3 mmol/L MgCl2, 500 nmol/L of forward and reverse primers, and 200 nmol/L of each hybridization probe. The amplification conditions consisted of one denaturation/activation cycle of 10 min at 95 °C and 45 cycles of three-temperature amplification. Each cycle consisted of 95 °C for 10 seconds, 60 °C for 10 seconds, and 72 °C for 15 seconds with a single fluorescence acquisition step at the 60 °C hold. This was followed by a melting-curve analysis of 95 °C for 20 seconds, 40 °C for 20 seconds, and a slow ramp (0.2 °C/second) to 85 °C with continuous fluorescence acquisition [8].
For LDLR 2054C>T genotyping the LightSNP® Kit rs28942084 LDLR [P685L] from TIB MOLBIOL (Berlin, Germany) whereas LDLR c.1474G>A; c.1567G>C; c.487dupC and c.647G>C mutations were identified by custom-made assays as previously described [8].
Results
Repeatability/reproducibility studies using five replicates of the same DNA sample or different batches of DNAs of heterogeneous genotypes were analysed five times and showed no intra-patient or between-batch variation. All LightCycler assays consistently identified the genotype correctly, confirming their analytical reliability and suitability for routine use.
All PCR methods demonstrated excellent robustness and analytical performance characteristics even when processing genomic DNA of less than optimal DNA purity (absorbance ratio 260/280 <1.6) and quantity (2.5–50 ng/μL). The genotype of all patients tested was correctly identified.
Figure 1 shows examples of wild-type and heterozygous for the LDLR c.1474G>A mutation. Heterozygote patients showed two distinct melting peaks and the G>A nucleotide substitution was detected by a melting temperature (Tm) shift of 7 °C.
In addition to ease of use and cost-effectiveness, a major advantage of this methodology is the rapid turn-around time of 90 min from genomic DNA extraction to PCR genotyping. This identifies potential uses outside large specialist centres in local one-stop clinics.
Discussion
The UK National Institute for Health and Care Excellence (NICE) recommends genetic testing of candidate patients presenting with FH phenotype and, once a disease-causing mutation is identified, screening of relatives; this is considered as the most cost-effective strategy for early detection of unsuspected cases of FH [9], and for distinguishing monogenic FH from sporadic or polygenic hypercholesterolaemia [10]. Detection of unknown mutations in the LDLR gene, where the majority of disease-causing mutations are found, requires complex and specialized molecular methods suitable for comprehensive scanning of the nucleotide sequence [11]. In contrast, once the disease-causing mutation has been identified, screening of relatives for the presence of the mutation does not pose a significant analytical challenge and a number of methodologies are available to the diagnostic services. Selection of these methods ultimately depends on local clinical service configuration, available laboratory expertise and resources and budget constraints. Some of these test requirements can be addressed by real-time PCR methods, which provide a cost-effective (the cost of each PCR method is estimated below £20) and rapid method for screening mutations associated with FH in family studies. Thus, these methods have the potential to deliver the second line of investigations of the FH cascade testing NICE pathway. The fast turn-around time of the method offers a significant advantage allowing the provision of a faster service as well as supporting delivery models such as a one-stop lipid clinic. This would allow the fast-tracking of clinical decision-making and choice of treatment as well as patient convenience, thus offering additional financial savings to the healthcare provider.
References
1. Marks D, Thorogood M, Neil HA, Humphries SE. A review on the diagnosis, natural history, and treatment of familial hypercholesterolaemia. Atherosclerosis 2003; 168: 1–14.
2. Benn M, Watts GF, Tybjaerg-Hansen A, Nordestgaard BG. Familial hypercholesterolemia in the Danish general population: prevalence, coronary artery disease, and cholesterol-lowering medication. J Clin Endocrinol Metab 2012; 97: 3956–3964.
3. Austin MA, Hutter CM, Zimmern RL, Humphries SE. Familial hypercholesterolemia and coronary heart disease: a HuGE association review. Am J Epidemiol 2004; 160: 421–429.
4. Neil A, Cooper J, Betteridge J, Capps N, McDowell I, Durrington P, Seed M, Humphries SE. Reductions in all-cause, cancer, and coronary mortality in statin-treated patients with heterozygous familial hypercholesterolaemia: a prospective registry study. Eur Heart J 2008; 29: 2625–2633.
5. Usifo E, Leigh SE, Whittall RA, Lench N, Taylor A, Yeats C, Orengo CA, Martin AC, Celli J, Humphries SE. Low-density lipoprotein receptor gene familial hypercholesterolemia variant database: update and pathological assessment. Ann Hum Genet 2012; 76: 387–401.
6. Chiou KR, Charng MJ, Chang HM. Array-based resequencing for mutations causing familial hypercholesterolemia. Atherosclerosis 2011; 216: 383–389.
7. Hinchcliffe M, Le H, Fimmel A, Molloy L, Freeman L, Sullivan D, Trent RJ. Diagnostic validation of a familial hypercholesterolaemia cohort provides a model for using targeted next generation DNA sequencing in the clinical setting. Pathology 2014; 46: 60–68.
8. Pandey S, Leider M , Khan M , Grammatopoulos DK. Cascade screening for familiar hypercholesterolaemia: PCR methods with melting-curve genotyping for the targeted molecular detection of apolipoprotein B and low density lipoprotein receptor gene mutations to identify affected relatives. JALM 2016; 02: 109–118.
9. Nherera L, Marks D, Minhas R, Thorogood M, Humphries SE. Probabilistic cost-effectiveness analysis of cascade screening for familial hypercholesterolaemia using alternative diagnostic and identification strategies. Heart 2011; 97: 1175–1181.
10. Talmud PJ, Shah S, Whittall R, Futema M, Howard P, Cooper JA, Harrison SC, Li K, Drenos F, et al. Use of low-density lipoprotein cholesterol gene score to distinguish patients with polygenic and monogenic familial hypercholesterolaemia: a case-control study. Lancet 2013; 381: 1293–1301.
11. Hollants S1, Redeker EJ, Matthijs G. Microfluidic amplification as a tool for massive parallel sequencing of the familial hypercholesterolemia genes. Clin Chem 2012; 58: 717–724.
The authors
Sarojini Pandey1 MSc and Dimitris K. Grammatopoulos*1,2 PhD, FRCPath
1Department of Clinical Biochemistry,
University Hospital Coventry and Warwickshire, Coventry CV2 2DX, UK
2Division of Translational and Systems Medicine, Warwick Medical School,
Coventry CV4 7AL,
UK
*Corresponding author
E-mail: Sarojini.Pandey@uhcw.nhs.uk