Keynote sessions on antimicrobial resistance, the microbiome and systems vaccinology as well as presentations on late-breaking research on refugee health and colistin resistance at ECCMID 2016
The annual meeting of the European Society of Clinical Microbiology and Infectious Diseases is taking place this year from April 9 – 12 in Amsterdam. At the world’s largest congress focused on infectious diseases and medical microbiology researchers will present more than 3,000 abstracts with the latest findings and recommendations, which are set to help improve diagnosis, prevention and the clinical care given to patients. Discussions on this vibrant platform not only help translate the research findings into diagnostic tools, guidelines, best practices, and international policies; they also raise awareness of emerging healthcare challenges. The congress offers more than 150 oral presentations, including keynote lectures, symposia, oral sessions, educational workshops and meet-the-experts session as well as more than 2,000 poster presentations. The event also provides mini-oral e-poster presentations. Posters are presented as printed posters, but also on e-poster viewing stations, where visitors can scroll through abstracts presented as papers. The main topics are strategies to detect and tackle antimicrobial resistance in various settings, approaches for prevention involving vaccines and infection control as well as descriptions of novel diagnostic technologies. The most popular sessions include lectures by winners of the ESCMID Award for Excellence and the Young Investigator Awards, as well as oral presentations on ground-breaking research approaches and findings, and the late-breaking abstracts. The keynote speeches include presentations on innovative approaches to vaccines; the microbiome and tuberculosis therapies; lectures on how non-human antibiotics affect public health; and an economic perspective on antimicrobial resistance.
This year, the ECCMID Programme Committee has decided to offer two special tracks for the late-breaking abstract sessions, focused on two topics, requiring a coordinated response from infection specialists across all disciplines. The first topic is refugee and migrant health. The thousands of people who are currently migrating challenge public health systems in transition and host countries. Clinicians and public health specialists need to develop strategies for the screening, the prevention, and the treatment of infectious diseases some of which were largely eradicated in Europe are now gradually being reintroduced. The second focus of the late-breaking abstracts is on emerging colistin resistance. Reports about the emergence of plasmid-borne resistance to this last-resort antibiotic have reached us from China, Canada, the UK and most countries in continental Europe.
Hala Audi, head of the UK government review on antimicrobial resistance (AMR review) will examine not only the long-term consequences of increasing antibiotic resistance in terms of healthcare, but also its economic cost. If the present situation fails to improve, the impact could be as high as ten million lives lost every year and €90 trillion in lost productivity by 2050. Hala Audi will present her findings on how we can address this, and describe new financial models, which may be necessary to start developing newer classes of antibiotics. Another keynote session by Prof. Lance B. Price of George Washington University will address how the use of antibiotics in animal food production is significantly contributing to antimicrobial resistance. Notably, he is pioneering the use of genomic epidemiology to understand how the misuse of antibiotics in animal feed affects public health. Prof. Price found that by analysing the genomes of bacteria – in human and animals – one is able to trace strains of antibiotic-resistant pathogens to industrial livestock productions. In light of this association, it is alarming that many companies are still using antibiotics to prevent infection spread – what is not clear, is how endemic this use is and to what extent antibiotic use can be minimized and avoided in livestock production.
In terms of viral infections, experts at the congress will evaluate HIV and hepatitis C treatments in several sessions. At the same time, researchers will present results on emerging infections including those caused by the Zika virus. The problem with the current outbreak of the Zika virus is that we do not yet have any definitive evidence on how it is affecting their hosts – particularly on its potential link to microcephaly and Guillain-Barré syndrome – or on how this outbreak is different from previous outbreaks, and most crucially of all, on how to prevent transmission. Recent reports from the U.S. have indicated that the virus may be transmitted sexually – yet only a few weeks ago the CDC was stating this as ‘only a theoretical risk’. It is important that infectious disease specialists get together and discuss how to best tackle outbreaks of emerging or re-emerging infectious diseases. ECCMID offers an interdisciplinary platform for these debates.
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Low-complexity detection of infectious diseases with high sensitivity and specificity is urgently needed, especially in resource-limited settings. Optofluidic integration combines clinical sample preparation with optical sensing on a single chip-scale system, enabling the direct, amplification-free detection of single RNA from Ebola viruses. The optofluidic system fulfils all key requirements for chip-based clinical analysis, including a low limit of detection, wide dynamic range, and the ability to detect multiple pathogens simultaneously.
by Dr Hong Cai, Prof. Aaron R. Hawkins and Prof. Holger Schmidt
Introduction The recent Ebola and Zika outbreaks [1, 2] have made it clear that viral infections continue to pose diverse and widespread threats to humanity. Resource-limited settings, in particular, call for diagnostic devices and technologies that are robust and feature relatively low complexity for easy handling by potentially unskilled personnel. At the same time, such instruments need to fulfil all the technical requirements for accurate and reliable diagnosis. These include a limit of detection and dynamic range that are compatible with clinically observed viral loads as well as the ability to carry out multiplexed differential detection by screening simultaneously for several pathogens with similar clinical symptoms.
The ‘gold standard’ test for hemorrhagic fevers as well as other infectious diseases is real-time polymerase chain reaction (RT-PCR) [3]. PCR fulfils the sensitivity and specificity requirement for clinical testing. However, it is not ideal for resource-limited environments and point-of-care applications because of to its complexity. An alternative economic and portable option is antigen-capture enzyme-linked immunosorbent assay (ELISA) testing. However, ELISA requires more highly concentrated samples and thus its clinical application, especially for early disease detection, is restricted.
For the last two decades, the lab-on-chip approach, which features a small footprint and sample volume, has been considered as a promising candidate for the next generation low-complexity medical diagnostics [4]. Among all the approaches, optofluidics, which integrates optics and microfluidics in the same platform, has received increased attention [5, 6]. Microfluidics is ideal for performing biological sample processing on a chip-scale level and leads to miniaturization and simplification of the current diagnostic system. If it can be integrated with an optical sensing/read-out platform that enables high detection sensitivity down to the single pathogen level, an analytic system for which nucleic acid amplification is no longer needed becomes possible.
In order to detect single molecular biomarkers and bioparticles, an in-flow based detection scheme is preferred. In a typical in-flow detection scheme, bioparticles are transported to the sensing region in a stream of gas or liquid where they are detected in transient fashion as they pass an optical interrogation region [7, 8]. Therefore, fast read-out of the optical signal from single bioparticles in sequence can be achieved, and many concerns associated with traditional surface-based sensing schemes such as unwanted nonspecific binding, probe photobleaching, and diffusion-limited transportation are eliminated.
Anti-resonant reflecting optical waveguides (ARROWs) have been proven to be highly efficient in detecting single bioparticles. By properly designing a Fabry–Perot reflector surrounding a hollow channel, light can propagate inside the ARROWs. Therefore, ARROWs confine both liquid and light in the same microfluidic channel, such that light and matter have near-perfect overlap and the sensing capability is maximized [8, 9]. Figure 1(a) shows a cross-section of a liquid-core ARROW using state-of-the-art fabrication technology [9]. Moreover, a two-dimensional photonic sensing platform can be constructed with lithography patterning. Figure 1(b) shows an ARROW platform with solid-core and liquid-core ARROWs crossing orthogonally. Excitation light from an external laser is confined in the solid-core ARROW, producing a few-micron-wide optical mode in the intersecting region. Liquid flow is generated inside the liquid-core ARROW to transport the bioparticles to the excitation volume which is of the order of femtolitres for typical waveguide and channel dimensions of a few micrometers. Optical read-out is extracted orthogonally through the liquid-core ARROW to achieve a low-noise signal, sufficient for reaching single particle fluorescence detection.
Besides the optical sensing aspect, miniaturizing and optimizing sample preparation is equally important in order to achieve a complete bioanalysis detection system. The ARROW-based optofluidic system is particularly well suited for such hybrid integration strategies. The planar optical layout based on intersecting solid-core and target-carrying liquid-core waveguides leaves the third dimension open for vertical integration of other functionalities. A separate microfluidic sample processing layer can be made and optimized and then connected to the ARROW platform (Fig. 1c) [10, 11]. Through this approach, we can perform multiple sample preparation steps, such as mixing, distributing, sorting and pre-concentrating on the microfluidic layer and transfer the sample to the ARROW chip for sensing without compromising each of the layers’ performance [10]. Amplification-free detection of Ebola nucleic acids on an opto- fluidic system In our recent work, Zaire Ebola virus RNA detection from clinical samples has been demonstrated in a hybrid optofluidic ARROW system [12]. Through a strain-specific solid-phase extraction method, we extracted and labelled target RNA from Ebola infected Vero cells and put them through the optofluidic chip for detection. The ARROW chip provided a sequence of optical signals when individual fluorescent virus RNAs passed through the small excitation volume. Figure 1(d) shows the recorded digital RNA counts at low concentration levels of from 2.1×102 to ~2.1×104 pfu/mL within one second. We were able to detect six orders of magnitude of the clinical concentration range using the ARROW chip only. The lower concentration limit is determined by the detection time, which was set to be 10 min maximum. As a negative control, we used the same method to test for Sudan Ebola virus and Marburg virus. Our results showed no detectable signals and thus our method is target specific.
In order to incorporate critical sample processing steps and detect RNA at even lower concentrations within 10 min, we adapted a programmable microfluidic chip – an automaton – to handle processing of larger sample volumes (Fig. 1b). The polydimethylsiloxane (PDMS) based automaton chip consists of a two-layer microvalve array. Each valve’s state is controlled individually by the top pneumatic layer through a reprogrammable software program. We used the automaton chip to perform an extra pre-concentration step by processing a large amount of clinical sample. We washed, released and labelled the RNA on the same automaton chip after pre-concentration. Through ~460× concentration, the virus detection limit was improved down to 0.2 pfu/mL, with seven orders of magnitude of concentration range (Fig. 1e). This demonstration exhibits an amplification-free chip-based virus and nucleic acid analysis technique with high sensitivity and wide dynamic range, whose performance is comparable with the gold standard, more complex PCR technique.
Wavelength division multiplexing detection ARROWs also enable simultaneous detection of multiple pathogens through the wavelength division multiplexing (WDM) technique [13]. WDM is generated using a multi-mode interferometer waveguide (MMI). When an MMI is excited by a single optical mode, all of the modes inside the MMI propagate at different phase velocities. When a constructive interference condition is satisfied, various numbers of self-imaging spots which resemble the excitation mode are formed along the MMI. This allows us to design an MMI section that intersects the fluidic channel, where multiple excitation spots are generated (Fig. 2a). As a fluorescent target flows past this excitation region, multi-peak signals are recorded in the time domain. For a single wavelength excitation, the fixed pattern multi-peak detection enables a signal-to-noise improvement compared to single-mode detection [14].
For a given MMI, the number of the self-imaging spots is wavelength dependent. We can generate various spot patterns at various laser wavelengths. For example, 9, 8 and 7 excitation spots are generated using 488nm, 553nm and 633nm lasers (Fig. 2b). With this approach, multiple targets labelled with different dye can be distinguished by the number and spacing of the peaks in the detected signal. Figure 2(c) shows influenza virus H1N1 and H3N2, which were labelled with different dye, generating 9 and 6 peaks in the time domain, respectively. We also labelled H2N2 virus with a combination of these two dyes which resulted in a superposition of the 9-spot and 6-spot fluorescence signals (Fig. 2c, bottom). A signal-processing algorithm checks for the presence of signals at the two characteristic time delays and can easily identify the mixed-labelled virus particle. This technique was shown to discriminate between three influenza subtypes, again with single virus sensitivity, using only two excitation colours. Thus, the ARROW-based platform has now met all the fundamental requirements for clinical virus detection using single particle sensing.
Conclusion For the next generation of medical diagnostic devices, low-complexity detection with high sensitivity and specificity is required on the detection side, along with small footprint and multi-functional analyte handling on the sample processing side. In-flow based optofluidic devices in which both analyte handling and optical sensing are carried out on the chip scale are promising candidates. Using our ARROW-based optofluidic system, we demonstrated multi-stage sample processing and detection of clinical Zaire Ebola virus samples using hybrid integration. We also demonstrated wavelength multiplex detection of multiple analytes at the same time. This fulfils all quantitative requirements for clinical virus detection. Therefore, a fully integrated microsystem for front-to-back amplification-free virus analysis is within reach.
References 1. Fact sheet. The top 10 causes of death. World Health Organization 2014. (http://www.who.int/mediacentre/factsheets/fs310/en/). 2. Fact sheet. Zika virus. World Health Organization 2016. (http://www.who.int/mediacentre/factsheets/zika/en/). 3. Kuypers J, Wright N, Morrow R. Evaluation of quantitative and type-specific real-time RT-PCR assays for detection of respiratory syncytial virus in respiratory specimens from children. J Clin Virol. 2004; 31: 123–129. 4. Craighead H. Future lab-on-a-chip technologies for interrogating individual molecules. Nature 2006; 442: 387–393. 5. Fan X, White IM. Optofluidic microsystems for chemical and biological analysis. Nature Photon. 2011; 5: 591–607. 6. Schmidt H, Hawkins AR. The photonic integration of non-solid media using optofluidics. Nature Photon. 2011; 5: 598–604. 7. Zhu H, White IM, Suter JD, Zourob M, Fan X. Opto-fluidic micro-ring resonator for sensitive label-free viral detection. Analyst 2008; 133: 356–360. 8. Bernini R, Campopiano S, Zeni L, Sarro PM. ARROW optical waveguides based sensors. Sensors and Actuators B 2004; 100: 143–146. 9. Yin D, Barber JP, Hawkins AR, Deamer DW, Schmidt H. Integrated optical waveguides with liquid cores. Appl Phys Lett. 2004; 85: 3477–3479. 10. Parks JW, Cai H, Zempoaltecatl L, Yuzvinsky TD, Leake K, Hawkins AR, Schmidt H. Hybrid optofluidic integration. Lab Chip 2013; 13: 4118–4123. 11. Testa G, Persichetti G, Sarro, PM, Bernini R. A hybrid silicon-PDMS optofluidic platform for sensing applications. Biomed Opt Express 2014; 5: 417–426. 12. Cai H, Parks JW, Wall TA, Stott MA, Stambaugh A, Alfson K, Griffiths A, Mathies RA, Carrion R, Patterson JL, Hawkins AR, Schmidt H. Optofluidic analysis system for amplification-free, direct detection of Ebola infection. Scientific Reports 2015; 5: 14494. 13. Ozcelik D, Parks JW, Wall TA, Stott MA, Cai H, Parks JW, Hawkins AR, Schmidt H. Optofluidic wavelength division multiplexing for single-virus detection. Proc Nat Acad Sci U S A 2015; 112: 12933–12937. 14. Ozcelik D, Stott MA, Parks JW, Black JA, Wall TA, Hawkins AR, Schmidt H. Signal-to-noise enhancement in optical detection of single viruses with multi-spot excitation, IEEE J Sel Top Quant Elec. 2016; DOI: 10.1109/JSTQE.2015.2503321.
The authors Hong Cai1 PhD, Aaron R. Hawkins2 PhD, Holger Schmidt*1 PhD 1School of Engineering, University of California Santa Cruz, Street, Santa Cruz, CA 95064 USA 2ECEn Department, 459 Clyde Building, Brigham Young University, Provo, UT 84602 USA
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Ebola virus (EBOV) can lead to severe hemorrhagic fever with a high risk of death in humans and other primates. More recently, reverse transcription loop-mediated isothermal amplification (RT-LAMP) has become readily available for the diagnosis of EBOV, and is a suitable tool for clinical screening, diagnosis and primary quarantine purposes.
by H. Li, W. Lin, X. Wang, X. Wei, E. Li, P. Li, J. Chen, S. Qi, Y. Ma, L. Cui, X. Hu, Dr X. Zhao, Prof. J. Yuan
The 2014 Ebola virus (EBOV; one of the world’s most virulent viruses) caused an outbreak of human disease with widespread transmission in multiple West African countries and sporadic cases in Europe and North America [1, 2]. The numbers of people infected and deaths were the most severe in history. However, the massive public health response has been limited, in part, by the inability to rapidly detect the presence of EBOV in potential patients living in remote areas [3].
EBOV, (species Zaire ebolavirus from the family Filoviridae), was first identified in Zaire in 1976 and named after the River Ebola in Zaire [4]. However, EBOV could not be detected rapidly in many potential patients living in remote and developing areas. The EBOV genome is approximately 19 kb, and encodes the seven proteins in the following order from the 3’-UTR: nucleoprotein (NP), viral structural protein (VSP)35, VSP40, glycoprotein (GP), VP30, VP24, and RNA-dependent RNA polymerase (L) [5]. As the NP gene is highly conserved among EBOV species, it is, therefore, recommended by the World Health Organization (WHO) for use as a target gene for the reverse transcription (RT)-PCR assay. The initial symptoms of EBOV infection could be confused with those of other febrile illnesses such as endemic malaria [6].
Current approaches for the laboratory diagnosis of EBOV infection include virus isolation, electron microscopy, immunohistochemistry, antigen-capture ELISA testing, IgM ELISA, RT-PCR, and serologic testing for IgM or IgG virus-specific antibodies. In 2015, Baca et al. presented a rapid detection of EBOV with a reagent-free, point-of-care biosensor. In general, the detection of EBOV antigens by antigen-capture ELISA is suitable as a method of laboratory diagnosis when the viral load in the blood reaches a very much higher case fatality rate. Thus, real-time (q)RT-PCR has taken over as a first choice diagnostic technique for detection of EBOV recommended by WHO [3]. However, Taq DNA polymerase in PCR-based techniques can be inactivated by inhibitors present in crude biological samples. Moreover, these methods are relatively complex and require specialized high-cost instruments.
Loop-mediated isothermal amplification (LAMP) is a one-step nucleic acid detection method developed by Notomi et al., which relies on autocycling strand displacement DNA synthesis [7]. This novel method is highly specific and sensitive, takes advantage of four or six specific primers to recognize six or eight different sequences of the target gene, and is performed under isothermal conditions in less than 1 h using Bst DNA polymerase. Kurosaki et al. developed a simple reverse transcription (RT)-LAMP assay for the detection of EBOV, targeting the trailer region of the viral genome. However, this method has yet to be tested in clinical samples [8].
To develop an RT-LAMP for clinical screening and rapid diagnosis of EBOV, we first selected potential target regions based on the NP sequences of the EBOV variant Mayinga (GenBank Accession no. AF086833), which were further analysed with Primer Explorer V4 software (http:/primerexplorer.jp/lamp) and subsequently the sequences were aligned with other species of EBOV. A total of five sets of primers were initially designed to detect artificially synthesized EBOV RNA using a real-time turbidimeter. To compare the sensitivity and specificity of RT-LAMP, normal RT-PCR was performed with the primers.
The RT-LAMP reactions were carried out in a 25-μl reaction mixture with an RNA amplification kit (Eiken Chemical Co. Ltd), in accordance with the manufacturer’s protocol. The reaction mixture contained the following reagents (final concentration): RT-LAMP mixture and 8 U Bst DNA polymerase. The amount of primer needed for one reaction was 80 pmol of forward and backward inner primers (FIP and BIP), 40 pmol of loop primer (LB), and 10 pmol of outer forward primer (F3) and outer backward primer (B3). Finally, an appropriate amount of genomic template DNA was added to the reaction tube. The reaction was carried out in the reaction tube at 61 °C, 60–80 min, in dry bath incubators.
Two different methods were used to detect RT-LAMP products. For direct visual inspection, 1 μl of calcein (fluorescent detection reagent; Eiken Chemical Co. Ltd) was added to 25 μl of LAMP products. For a positive reaction, the colour changed from orange to green, whereas a negative reaction remained orange. The colour change could be observed by the naked eye under natural light or with the aid of UV light at 365 nm. For monitoring turbidity, real-time amplification by the RT-LAMP assay was monitored by spectrophotometry, recording the optical density at 650 nm every 6 s with the help of a Loopamp Realtime Turbidimeter (LA-230; Eiken Chemical Co. Ltd) [9]. Assay validation 1. Optimal primer choice and reaction temperature conditions for the RT-LAMP assay As shown in Figure 1A, the EBL-2 primer set amplified the NP gene using the shortest time of about 10min; therefore, this was chosen as the optimal primer set for EBOV detection of RT-LAMP (Table 1). To further optimize the amplification, reaction temperatures were compared ranging from 59 °C to 69 °C at 2 °C intervals. Ultimately, 61 °C was chosen as the optimal reaction temperature (Fig. 1B).
2. Specificity of NP detection by RT-LAMP using the artificial in vitro transcribed RNA Twenty-five other non-EBOV viruses were also tested. As shown in Figure 2, the EBOV RNA was identified positively by a successful RT-LAMP reaction with EBL-2 primer set using both methods of analysis. All non-EBOV strains tested negative, including the blank control, indicating that the RT-LAMP method was specific for EBOV. 3. Sensitivity of NP detection by RT-LAMP A 10-fold serial dilution of artificial EBOV RNA was tested by real-time turbidity monitoring (Fig. 3A), visual detection method (Fig. 3B), and qRT-PCR (Fig. 3C). The limit of detection by the visual method was 10-fold lower compared with the qRT-PCR assay.
4. Clinical sample detection The 417 clinical blood or swab samples were analysed by RT-LAMP and qRT-PCR simultaneously. The RT-LAMP and qRT-PCR detections both showed that 307 patients were confirmed cases of EBOV infections and 106 patients tested negative for EBOV.
Summary Zaire ebolavirus is a key member of the Filoviridae family and causes highly lethal hemorrhagic fever in human beings with extremely high morbidity and mortality. As a typical negative-sense single-stranded RNA (ssRNA) virus, EBOV possesses a nucleoprotein (NP) to facilitate genomic RNA encapsidation to form a viral ribonucleoprotein complex (RNP) together with genome RNA and polymerase, which plays the most essential role in virus proliferation cycle. EBOV is found in Central Africa, but re-emerged in Western Africa in 2014 to cause an outbreak that threatened to spread worldwide. Up until 10 January 2016, 28 601 total cases (including suspected, probable, and confirmed) and 11 300 deaths were reported in Guinea and Sierra Leone (http://www.cdc.gov/vhf/ebola/outbreaks/2014-west-africa/case-counts.html). Although several chemical agents, antibodies and vaccines are found to inhibit EBOV in animals or humans, there is no therapeutic with high efficacy that can be provided for clinical usage.
To combat the increasing incidence of EBOV infections, we developed and optimized a novel RT-LAMP assay specific for EBOV diagnosis using primers spanning the 663 bp NP sequence of the viral genome. In the RT-LAMP assay, the reverse transcription reaction and DNA amplification proceed in a single step and with incubation of the reaction mixture at a constant 61°C temperature for a given time period using a temperature-controlled water bath (or other devices that can provide a stable heat are also sufficient). Moreover, LAMP reaction primers specifically recognize five independent regions of the target sequence, compared to PCR primers that recognize two independent regions of the target sequence. The sensitivity of the PCR reaction can be greatly reduced by the presence of exogenous DNA and inhibitors. Therefore, the RT-LAMP method is more suitable for rapid detection of NP in clinical samples. Conclusion In conclusion, a specific, sensitive, rapid and cost effective RT-LAMP assay for NP detection in EBOV was established, which is as sensitive as other available technologies, highly specific and extremely rapid in the provision of molecular diagnosis of EBOV infections. The assay can provide accurate results in a short time frame. This makes it potentially useful for clinical diagnosis of EBOV in developing countries. Acknowledgment This article is based on one previously published by the authors: Li H, Wang X, Liu W, Wei X, Lin W, Li E, Li P, Dong D, Cui L, Hu X, Li B, Ma Y, Zhao X, Liu C, Yuan J. Survey and Visual detection of Zaire ebolavirus in clinical samples targeting the nucleoprotein gene in Sierra Leone. Frontiers in Microbiology 2015; 6: 1332 [10].
References 1. Frieden TR, Damon I, Bell BP, Kenyon T, Nichol S. 2014. Ebola 2014—New challenges, new global response and responsibility. N Engl J Med. 371(13): 1177–1180. 2. Hampton T. Largest-ever outbreak of Ebola virus disease thrusts experimental therapies, vaccines into spotlight. JAMA 2014; 312(10): 987–989. 3. Urgently needed: rapid, sensitive, safe and simple Ebola diagnostic tests. World Health Organization 2014. (http://www.who.int/mediacentre/news/ebola/18-november-2014-diagnostics/en/). 4. MacNeil A, Rollin PE. Ebola and Marburg hemorrhagic fevers: Neglected tropical diseases? PLoS Negl Trop Dis. 2012; 6(6): e1546. 5. Ali MT, Islam MO. A highly conserved GEQYQQLR epitope has been identified in the nucleoprotein of Ebola virus by using an in silico approach. Adv Bioinformatics 2015; 2015: 278197–278203. 6. Grolla A, Lucht A, Dick D, Strong JE, Feldmann H. Laboratory diagnosis of Ebola and Marburg hemorrhagic fever. Bull Soc Pathol Exot. 2005; 98(3):205–209. 7. Notomi T, Okayama H, Masubuchi H, Yonekawa T, Watanabe K, Amino N, Hase T. Loop-mediated isothermal amplification of DNA. Nucleic Acids Res. 2000; 28, E63. 8. Kurosaki Y, Takada A, Ebihara H, Grolla A, Kamo N, Feldmann H, Kawaoka Y, Yasuda J. Rapid and simple detection of Ebola virus by reverse transcription-loop-mediated isothermal amplification. J Virol Methods 2007; 141(1): 78–83. 9. Mori Y, Nagamine K, Tomita N, Notomi T. Detection of loop-mediated isothermal amplification reaction by turbidity derived from magnesium pyrophosphate formation. Biochem Biophys Res Commun. 2001; 289: 150–154. 10. Li H, Wang X, Liu W, Wei X, Lin W, Li E, Li P, Dong D, Cui L, Hu X, Li B, Ma Y, Zhao X, Liu C, Yuan J. Survey and Visual detection of Zaire ebolavirus in clinical samples targeting the nucleoprotein gene in Sierra Leone. Frontiers in Microbiology 2015; 6: 1332.
The authors Huan Li# MMed, Weishi Lin# MMed, Xuesong Wang MMed, Xiao Wei MMed, Erna Li MMed, Puyuan Li MMed, Jun Chen MMed, Silei Qi MMed, Yanyan Ma MMed, Lifei Cui MMed, Xuan Hu MMed, Xiangna Zhao PhD, Jing Yuan PhD* Institute of Disease Control and Prevention, Academy of Military Medical Sciences, Beijing, 100071, PR China
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Biochemical markers of alcohol intake can be separated into two categories: direct markers of ethanol metabolism and indirect markers. The different alcohol markers have varying time windows of detection and are a useful additional tool to detect alcohol intake in alcohol-dependent clients.
by Jane Armer and Rebecca Allcock
Introduction Alcohol dependence is characterized by craving, tolerance, a preoccupation with alcohol and continued drinking in spite of harmful consequences. The World Health Organization Alcohol Use Disorders Identification Test (AUDIT) is recommended for the identification of individuals that are dependent on alcohol [1]. The prevalence of alcohol use disorders (including dependence and harmful use of alcohol) is 11.1% in the UK compared to 7.5% across Europe [2]. In England, 250 000 people are believed to be moderately or severely dependent and require intensive treatment [3].
Alcohol use is the third leading risk factor contributing to the global burden of disease after high blood pressure and tobacco smoking [4]. In 2012, 3.3 million deaths (5.9% of all global deaths) were attributable to alcohol consumption [2]. It is estimated that the UK National Health Service (NHS) spends £3.5 billion/year in costs related to alcohol and the number of alcohol-related admissions has doubled over the last 15 years [3].
In the UK, one unit equals 10 mL or 8 g of pure alcohol, which is around the amount of alcohol the average adult can process in an hour. The latest UK recommendations are to not regularly drink more than 14 units per week (men and women) and to limit the total amount of alcohol consumed on a single occasion [5].
The most common entry into alcohol treatment services in England is either self-referral or referral by the GP [3]. Services have a limited number of options to determine if an individual in treatment for alcohol dependence is continuing to drink alcohol. They rely on self-report by the individuals in the form of alcohol diaries and breathalyser tests. There is no regular schedule for biochemical markers. If a client is found to be drinking alcohol during the treatment programme, an assessment is done of the amount of alcohol consumed, the pattern of alcohol consumption and how it will impact on their treatment. This is factored into the recovery plan and there is a re-assessment of the support and interventions needed for that client. Possible interventions include cognitive behavioural therapies, pharmacological therapies or in-patient assisted withdrawal. In 2013/14, only 38% of clients in alcohol treatment in England successfully completed their treatment [3]. Monitoring clients in alcohol treatment Diaries that record alcohol intake are commonly used to monitor the progress of clients. However, this relies on accurate self-reporting of alcohol intake by the client and under reporting is a common problem. Biochemical markers of alcohol intake can provide a more comprehensive assessment of a client’s progress.
Direct markers of alcohol intake Direct markers of alcohol intake include ethanol, ethyl glucuronide (EtG), ethyl sulphate (EtS), fatty acid ethyl esters (FAEE) and phosphatidylethanol (PEth).
Following the ingestion of ethanol, >95% is metabolized in the liver by alcohol dehydrogenase to acetaldehyde then by aldehyde dehydrogenase to acetic acid [14]. Less than 5% is excreted unchanged in the urine, breath and sweat. A small amount of ethanol is conjugated to form EtG and EtS (Fig. 1). Ethanol is usually only detectable in breath and urine after very recent alcohol consumption and the detection time window depends on the amount of alcohol consumed. In comparison, urine EtG and EtS remain detectable for around 24 hours after moderate alcohol intake and for up to 130 hours in subjects admitted for alcohol detoxification [6, 7]. Liquid chromatography-tandem mass spectrometry (LC-MS/MS) methods have been developed for EtG and EtS. An immunoassay is also available for EtG [8, 9].
Many studies have demonstrated the benefit of measuring EtG and EtS in clients in alcohol treatment. Continued alcohol consumption can be detected by the measurement of urine EtG and EtS in clients who do not admit to consuming alcohol and provide a negative breathalyser test. This is due to the increased time window of detection for urine EtG and EtS compared to breath ethanol. This demonstrates the unreliability of self-reporting of alcohol intake and the benefit of biochemical markers to detect clients that are continuing to drink alcohol [10].
As with urine testing for drugs of abuse, it is possible for a client to consume a large volume of water to dilute the sample and produce negative EtG and EtS results. Creatinine should always be measured to check for adulteration and it may be beneficial to report EtG and EtS as creatinine ratios to overcome this problem. Further work is required to define cut-offs for EtG and EtS as creatinine ratios.
False negative EtG results can be caused by the presence of Escherichia coli in urine as glucuronidase is present with high activity in most strains. False positive EtG and EtS results have also been reported following use of ethanol based mouthwash or hand gels and after the consumption of non-alcoholic beers (up to 0.5% alcohol). Due to the risk of positive results due to unintentional alcohol exposure, particularly for urine EtG, it is important that clinical cut-offs used are clearly defined and LC-MS/MS methods that measure both EtG and EtS are preferred [11]. In the USA, the Substance Abuse and Mental Health Administration (SAMHSA) have suggested that EtG results >1.0 mg/L are consistent with alcohol intake and that results between 0.1 and 1.0 mg/L should be interpreted with caution. It is accepted that further work is required to clearly define cut-offs for EtG and EtS and that other biomarkers may be useful when interpreting borderline positive results in the range 0.10–0.50 mg/L [12].
Methods for the measurement of EtG and FAEEs in hair have been developed allowing a longer term assessment of alcohol intake. Hair analysis is most suitable for subjects where longer term abstinence needs to be demonstrated such as in patients awaiting liver transplantation. EtG cut-offs have been suggested by the Society of Hair Testing for chronic excessive alcohol consumption (30 pg/mg) and abstinence assessment (7 pg/mg). However, results may be influenced by hair products and this needs to be taken into account when interpreting results.
PEth is formed from ethanol and phosphatidylcholine in cell membranes. The reaction is catalysed by phospholipase D and occurs in the cell membranes of erythrocytes; therefore, PEth is found in the red blood cell fraction of blood rather than in serum or plasma. PEth is a group of phospholipids with varying carbon lengths and LC-MS/MS methods to detect the major forms of PEth in whole blood have been developed. A single dose of ethanol does not produce a measurable amount of PEth and it has been demonstrated that approximately 50 g of ethanol/day (6.25 UK units) is required to provide a positive PEth result. In comparison to serum carbohydrate deficient transferrin (CDT; see ‘Indirect markers of alcohol intake’ below), urine EtG and urine EtS, PEth demonstrated the highest sensitivity for regular alcohol consumption in clients in alcohol treatment and was found to be positive twice as often as CDT [13]. Further work is required to understand how PEth can be used optimally in combination with other alcohol markers in clients in treatment for alcohol dependence [14].
Indirect markers of alcohol intake The indirect markers include mean corpuscular volume (MCV), gamma glutamyl transferase (GGT) and CDT. These markers increase following significant alcohol intake over a prolonged time period and are not useful for detecting a single alcohol ‘binge’. MCV and GGT are not specific markers of alcohol intake.
CDT refers to altered glycoforms of transferrin as a result of alcohol-induced changes in the carbohydrate composition of transferrin. The main component of serum transferrin is tetrasialotransferrin, which makes up approximately 80% of the total. Normal samples usually contain approximately 15%, 4–5%, 1–1.5% and 1% of pentasialotransferrin, trisialotransferrin, disialotransferrin and hexasialotransferrin, respectively. An alcohol consumption of at least 60 g/day (7.5 UK units) for 2 weeks is required to increase the disialotransferrin [15]. CDT may also be increased if genetic variants are present and in advanced liver disease. The International Federation of Clinical Chemistry and Laboratory Medicine (IFCC) has recently proposed a reference measurement procedure for CDT and more studies assessing the diagnostic performance of CDT to detect alcohol dependence are now needed using methods harmonized to the international reference measurement procedure.
Table 1 summarizes the time window of detection and limitations of the alcohol markers discussed.
Conclusions Currently, the assessment of clients in alcohol treatment relies largely on self-reporting and limited biochemical testing, which makes assessment of a client’s progress challenging. There are a number of available biochemical markers that could improve the detection of alcohol use in clients with alcohol dependence and ultimately lead to initiation of early intervention and altered treatment strategies. This in turn could improve the numbers successfully completing treatment. A combination of short-term and longer term biochemical markers is likely to be the most useful approach depending on the treatment setting. The advantage of the breathalyser test over biochemical markers that require laboratory analysis is the immediate availability of the result which allows an immediate intervention for a client with a positive result. Laboratory tests need to be available in a timely manner and with appropriate and well-defined cut-offs. The clinical benefit of alcohol markers in improving the number of clients that successfully complete their treatment for alcohol dependency has not yet been demonstrated. Randomized controlled trials comparing outcomes with or without the use of biochemical markers are required.
References 1. Babor TF, Higgins-Biddle JC, Saunders JB, Monteiro MG. Alcohol use disorders identification test (AUDIT). World Health Organization, 2001. (http://www.alcohollearningcentre.org.uk/Topics/Browse/BriefAdvice/?parent=4444&child=4896) 2. Global status report on alcohol and health. World Health Organization, 2014. (http://www.who.int/substance_abuse/publications/global_alcohol_report/msb_gsr_2014_2.pdf?ua=1) 3. Alcohol Treatment England 2013–14. Public Health England, 2014. (http://www.nta.nhs.uk/uploads/adult-alcohol-statistics-2013-14-commentary.pdf ) 4. Lim S, Vos T, Flaxman A, Danaei G, Shibuya K, Adair-Rohani H, Amann M, Anderson HR, Andrews KG, et al. A comparative risk assessment of burden of disease and injury attributable to 67 risk factors and risk factor clusters in 21 regions, 1990-2010: a systematic analysis for the Global Burden of Disease Study 2010. Lancet 2012; 380: 2224–2260. 5. UK Chief Medical Officers’ Alcohol Guidelines Review. Department of Health, 2016. (https://www.gov.uk/government/uploads/system/uploads/attachment_data/file/489795/summary.pdf) 6. Dahl H, Stephanson N, Beck O, Helander A. Comparison of urinary excretion characteristics of ethanol and ethyl glucuronide. J Anal Toxicol. 2002; 26: 201–204. 7. Helander A, Bottcher M, Fehr C, Dahmen N, Beck A. Detection times for urinary ethyl glucuronide and ethyl sulphate in heavy drinkers during alcohol detoxification. J Anal Toxicol. 2009; 44: 55–61. 8. Politi L, Morini L, Groppi A, Poloni V, Pozzi F, Polettini A. Direct determination of the ethanol metabolites ethyl glucuronide and ethyl sulphate in urine by liquid chromatography/electrospray tandem mass spectrometry. Rapid Commun Mass Spectrom. 2005; 19: 1321–1331. 9. Bottcher M, Beck O, Helander A. Evaluation of a new immunoassay for urinary ethyl glucuronide testing. Alcohol Alcohol. 2008; 43: 46–48. 10. Junghanns K, Graf I, Pfluger J, Wetterling G, Ziems C, Ehrenthal D, Zöllner M, Dibbelt L, Backhaus J, Weinmann W, Wurst FM. Urinary ethyl glucuronide (EtG) and ethyl sulphate (EtS) assessment: valuable tools to improve verification of abstention in alcohol-dependent patients during in-patient treatment and at follow ups. Addiction 2009; 104: 921–926. 11. Wurst F, Thon N, Yegles M, Schruck A, Preuss UW, Weinmann W. Ethanol metabolites: their role in the assessment of alcohol intake. Alcohol Clin Exp Res. 2015; 39: 2060–2072. 12. The role of biomarkers in the treatment of alcohol use disorders. SAMHSA, 2012. (http://store.samhsa.gov/product/The-Role-of-Biomarkers-in-the-Treatment-of-Alcohol-Use-Disorders-2012-Revision/SMA12-4686) 13. Helander A, Peter O, Zheng Y. Monitoring of the alcohol biomarkers PEth, CDT and EtG/EtS in an outpatient treatment setting. Alcohol Alcohol. 2012; 47: 552–557. 14. Viel G, Boscalo-Berto R, Cecchetto G, Fais P, Nalesso A, Ferrara SD. Phosphatidylethanol in blood as a marker of chromic alcohol use: a systematic review and emta-analysis. Int J Mol Sci. 2012; 13: 14788–14812. 15. Stibler H. Carbohydrate Deficient Transferrin in serum: a new marker of potentially harmful alcohol consumption reviewed. Clin Chem. 1991; 37: 2029–2037.
The authors Jane Armer*1 BA MSc FRCPath and Rebecca Allcock2 BSc MSc FRCPath 1Department of Blood Sciences, East Lancashire Hospitals NHS Trust, Blackburn, UK 2Department of Clinical Biochemistry, Lancashire Teaching Hospitals NHS Foundation Trust, Preston, UK
Niguarda Hospital in Milan is one of Italy’s leading general hospitals, and provides an extensive range of medical disciplines for adults and children throughout the Lombardy region and beyond.
Our hospital’s Department of Laboratory Medicine aim is to offer a complete, continuous and prompt diagnostic laboratory testing service, in order to guarantee effective support for this widespread clinical demand, and is committed to research into automation and analysis to ensure this is maintained. Our busy Molecular Biology Laboratory performed an estimated 40,000 tests in 2015, which is approximately 10% increase on the previous year.
The growing annual molecular workload is attributed, in part, to the development of new therapeutic strategies. Our staff, consisting of 8 laboratory technicians, one director and one manager, work 5 days per week and are expected to cope with increased workloads and demands for reduced turnaround times without any increase in resources, in terms of the number of staff and costs.
A large proportion of the molecular biology workload consists of viral load measurements for human immunodeficiency virus type 1 (HIV-1), hepatitis C virus (HCV), hepatitis B virus (HBV) and cytomegalovirus (CMV) (figure 1).
With a very important Italian transplant centre located at Niguarda Hospital, CMV analyses are vital and results are needed quickly, without delay. In addition, the laboratory performs viral load measurements for HIV-1, HCV, and HBV in order to evaluate and monitor therapeutic response. In these instances, rapid results are extremely important for patient management decisions, for example to maintain or change treatment.
Since 2005, these measurements have been performed using our laboratory’s current method, which has separate sample preparation and amplification/detection platforms. These are situated on separate benches within the same room, with one sample preparation system in another room. The accuracy and precision of this method is good, however, in order to be cost effective, it is necessary to optimize the size of the batches. Since they can’t be processed in the same day, sample test tubes often need to be collected and stored for several days, which increases the turnaround time considerably. In addition, this method involves many manual steps, which demand time, space and coordination of work between different members of staff.
A new automated molecular diagnostics method As part of our Department of Laboratory Medicine’s investigations into increased automation in the laboratory, Niguarda Hospital became a beta trial site for the new DxN VERIS Molecular Diagnostics System (Beckman Coulter), which consolidates DNA extraction, nucleic acid amplification, quantification and detection onto a single automated instrument for a number of molecular targets, including HIV-1, HCV, HBV and CMV.
The first step in assessing the DxN VERIS was to validate the assays in order to determine whether their performance is comparable with our laboratory’s existing method. Daily quality control measurements demonstrated good performance of the VERIS HBV assay for high level, low level and negative HBV samples (table 1). This assay was also shown to have excellent linearity within the range of 1.68 – 8.82 Log IU/mL, a limit of detection of 6.82 IU/mL, and good precision, achieving within run and between run mean standard deviations of less than 0.16 (table 2).
A series of performance evaluation studies, conducted in several laboratories around the world, have demonstrated that the VERIS HBV, HCV, HIV-1 and CMV assays have comparable precision, sensitivity and linearity to a range of alternative, commercially available viral load methods [1-13]. In accordance with these findings, the VERIS HBV assay correlated well with the existing method at Niguarda Hospital (Abbott m2000) and, indeed, detected HBV DNA in 23 samples that were negative using the current method, 22 of which were found to be positive by one or more serology assay (table 3). Regarding the 55 specimens that were quantified both with DxN VERIS and Abbott m2000, 7 of them had an HBV DNA concentration discordant for more than 1 Log.
Comparable performance, including sensitivity and specificity, was achieved for each of the DxN VERIS assays: HIV-1, HCV, HBV and CMV.
Workflow improvements In addition to validating the performance of the VERIS assays, a time/workflow analysis study was performed at Niguarda Hospital by Nexus Global Solutions (Plano, Texas, USA). The study compared workflows and time to results between the current viral load method for HIV-1, HCV, HBV and CMV (Abbott m2000sp and m2000rt systems) and the new DxN VERIS Molecular Diagnostic System.
By reducing manual intervention and automating processes from sample loading to reporting of results, the DxN VERIS offers the potential to transform clinical laboratory workflows. Each assay is supplied in a unique, single cartridge system, and all consumables and reagents are stored on board the system, which cuts preparation time compared to alternative methods. In addition, unlike traditional plate-based systems, there is no need to batch assays. The DxN VERIS allows true, single sample random access, which means that viral load assays can be performed as soon as they arrive in the laboratory. This, combined with short assay runtimes, ensures rapid turnaround of results and, since there are no empty plate wells, wastage and consumable costs are reduced.
The comparative time/workflow analysis in our study revealed that DxN VERIS involved only 10 steps and required just five reagents, compared to 26 steps and over 20 consumables for the current method, and required much less hands-on time for each of the viral load assays (figure 2). Notably, by consolidating the assay menu, time savings of up to 2 hours could be achieved.
In addition to an increase in productivity (achieving more results in an 8-hour working day), the time to the first result for the DxN VERIS was greatly reduced compared to the current method, with subsequent results available every 2.5 minutes. This is in contrast to the current method, where results are not available until the end of the assay run (table 4).
With these time savings, and by eliminating the need to batch samples, the DxN VERIS allowed much faster turnaround of results in a normal working week, with all results being reported within 8 hours of receipt, unlike the current method, which often required several days (figure 3).
The true single sample random access capability of the DxN VERIS has the potential to simplify sample management in the laboratory and to make the organization of viral load assays more fluid. It increases productivity by allowing the continuous loading of samples for different assays, eliminating the need for batching and reducing turnaround times. This is the most important advantage of random access testing for us because it increases the availability of medical reports to the different departments and is a great benefit to patient management and care by allowing more timely clinical decisions.
The DxN VERIS is easy to use with its few consumables, reduced maintenance requirements, complete automation and intuitive computer interface. By improving laboratory organization and workflows and reducing manual intervention, viral loads (which account for about 50% of the molecular workload) could be completed in a single day using the DxN VERIS. Requiring fewer people to be dedicated to this purpose, this makes it possible to accomplish more work with the same number of staff.
For further information about the DxN VERIS Molecular Diagnostic System and the VERIS assays currently available, please contact: Tiffany Page, Senior Pan European Marketing Manager Molecular Diagnostics, Email: info@beckmanmolecular.com or visit www.beckmancoulter.com/moleculardiagnostics References 1. Williams, JA, Rodriguez, J, Wang, Z et al (2014) Poster presentation, ESCV, Prague. 2. Drago, M, Franchetti, E, Fanti, D and Gesu, GP (2015) Poster presentation, EuroMedLab, Paris. 3. Zurita, S, Gutiérrez, F, Folgueira, MD et al (2015) Poster presentation, EuroMedLab, Paris. 4. Christenson, R, Maggert, K, Ruiz, RM et al (2015) Poster presentation, ECCMID, Copenhagen. 5. Trimoulet, P, Tauzin, B, Belloc, E et al (2015) Poster presentation, EuroMedLab, Paris. 6. Gilfillan, R, Wang, Z, Xu, Y et al (2014) Poster presentation, ECCMID, Barcelona. 7. Xu, Y, Gilfillan, R, Wang, Z et al (2014) Poster presentation, ESCV, Prague. 8. Mengelle, C, Sauné, K, Haslé, C et al (2014) Poster presentation, RICAI. 9. Mengelle, C, Sauné, K, Haslé, C et al (2015) Poster presentation, ECCMID, Copenhagen. 10. Silvestro, A, Duan, H, Lim, S et al (2014) Poster presentation, ECCMID, Barcelona. 11. Li, Q, Williams, J, Maggert, K et al (2014) Poster presentation, ECCMID, Barcelona. 12. Xu, Y, Dineen, S, Annese, V et al (2014) Poster presentation, ESCV, Prague. 13. Williams, JA, Rodriguez, J, Wang, Z et al (2014) Poster presentation, ECCMID, Barcelona. The author Diana Fanti, Molecular Biology Laboratory Manager Department of Laboratory Medicine, Niguarda Hospital, Milan, Italy
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Trends in infectious diseases and clinical microbiology at ECCMID 2016
, /in Featured Articles /by 3wmediaKeynote sessions on antimicrobial resistance, the microbiome and systems vaccinology as well as presentations on late-breaking research on refugee health and colistin resistance at ECCMID 2016
The annual meeting of the European Society of Clinical Microbiology and Infectious Diseases is taking place this year from April 9 – 12 in Amsterdam. At the world’s largest congress focused on infectious diseases and medical microbiology researchers will present more than 3,000 abstracts with the latest findings and recommendations, which are set to help improve diagnosis, prevention and the clinical care given to patients. Discussions on this vibrant platform not only help translate the research findings into diagnostic tools, guidelines, best practices, and international policies; they also raise awareness of emerging healthcare challenges.
The congress offers more than 150 oral presentations, including keynote lectures, symposia, oral sessions, educational workshops and meet-the-experts session as well as more than 2,000 poster presentations. The event also provides mini-oral e-poster presentations. Posters are presented as printed posters, but also on e-poster viewing stations, where visitors can scroll through abstracts presented as papers.
The main topics are strategies to detect and tackle antimicrobial resistance in various settings, approaches for prevention involving vaccines and infection control as well as descriptions of novel diagnostic technologies. The most popular sessions include lectures by winners of the ESCMID Award for Excellence and the Young Investigator Awards, as well as oral presentations on ground-breaking research approaches and findings, and the late-breaking abstracts.
The keynote speeches include presentations on innovative approaches to vaccines; the microbiome and tuberculosis therapies; lectures on how non-human antibiotics affect public health; and an economic perspective on antimicrobial resistance.
This year, the ECCMID Programme Committee has decided to offer two special tracks for the late-breaking abstract sessions, focused on two topics, requiring a coordinated response from infection specialists across all disciplines.
The first topic is refugee and migrant health. The thousands of people who are currently migrating challenge public health systems in transition and host countries. Clinicians and public health specialists need to develop strategies for the screening, the prevention, and the treatment of infectious diseases some of which were largely eradicated in Europe are now gradually being reintroduced.
The second focus of the late-breaking abstracts is on emerging colistin resistance. Reports about the emergence of plasmid-borne resistance to this last-resort antibiotic have reached us from China, Canada, the UK and most countries in continental Europe.
Hala Audi, head of the UK government review on antimicrobial resistance (AMR review) will examine not only the long-term consequences of increasing antibiotic resistance in terms of healthcare, but also its economic cost. If the present situation fails to improve, the impact could be as high as ten million lives lost every year and €90 trillion in lost productivity by 2050. Hala Audi will present her findings on how we can address this, and describe new financial models, which may be necessary to start developing newer classes of antibiotics.
Another keynote session by Prof. Lance B. Price of George Washington University will address how the use of antibiotics in animal food production is significantly contributing to antimicrobial resistance. Notably, he is pioneering the use of genomic epidemiology to understand how the misuse of antibiotics in animal feed affects public health. Prof. Price found that by analysing the genomes of bacteria – in human and animals – one is able to trace strains of antibiotic-resistant pathogens to industrial livestock productions. In light of this association, it is alarming that many companies are still using antibiotics to prevent infection spread – what is not clear, is how endemic this use is and to what extent antibiotic use can be minimized and avoided in livestock production.
In terms of viral infections, experts at the congress will evaluate HIV and hepatitis C treatments in several sessions. At the same time, researchers will present results on emerging infections including those caused by the Zika virus. The problem with the current outbreak of the Zika virus is that we do not yet have any definitive evidence on how it is affecting their hosts – particularly on its potential link to microcephaly and Guillain-Barré syndrome – or on how this outbreak is different from previous outbreaks, and most crucially of all, on how to prevent transmission. Recent reports from the U.S. have indicated that the virus may be transmitted sexually – yet only a few weeks ago the CDC was stating this as ‘only a theoretical risk’. It is important that infectious disease specialists get together and discuss how to best tackle outbreaks of emerging or re-emerging infectious diseases. ECCMID offers an interdisciplinary platform for these debates.
Amplification-free direct detection of Ebola virus on a hybrid optofluidic platform
, /in Featured Articles /by 3wmediaLow-complexity detection of infectious diseases with high sensitivity and specificity is urgently needed, especially in resource-limited settings. Optofluidic integration combines clinical sample preparation with optical sensing on a single chip-scale system, enabling the direct, amplification-free detection of single RNA from Ebola viruses. The optofluidic system fulfils all key requirements for chip-based clinical analysis, including a low limit of detection, wide dynamic range, and the ability to detect multiple pathogens simultaneously.
by Dr Hong Cai, Prof. Aaron R. Hawkins and Prof. Holger Schmidt
Introduction
The recent Ebola and Zika outbreaks [1, 2] have made it clear that viral infections continue to pose diverse and widespread threats to humanity. Resource-limited settings, in particular, call for diagnostic devices and technologies that are robust and feature relatively low complexity for easy handling by potentially unskilled personnel. At the same time, such instruments need to fulfil all the technical requirements for accurate and reliable diagnosis. These include a limit of detection and dynamic range that are compatible with clinically observed viral loads as well as the ability to carry out multiplexed differential detection by screening simultaneously for several pathogens with similar clinical symptoms.
The ‘gold standard’ test for hemorrhagic fevers as well as other infectious diseases is real-time polymerase chain reaction (RT-PCR) [3]. PCR fulfils the sensitivity and specificity requirement for clinical testing. However, it is not ideal for resource-limited environments and point-of-care applications because of to its complexity. An alternative economic and portable option is antigen-capture enzyme-linked immunosorbent assay (ELISA) testing. However, ELISA requires more highly concentrated samples and thus its clinical application, especially for early disease detection, is restricted.
For the last two decades, the lab-on-chip approach, which features a small footprint and sample volume, has been considered as a promising candidate for the next generation low-complexity medical diagnostics [4]. Among all the approaches, optofluidics, which integrates optics and microfluidics in the same platform, has received increased attention [5, 6]. Microfluidics is ideal for performing biological sample processing on a chip-scale level and leads to miniaturization and simplification of the current diagnostic system. If it can be integrated with an optical sensing/read-out platform that enables high detection sensitivity down to the single pathogen level, an analytic system for which nucleic acid amplification is no longer needed becomes possible.
In order to detect single molecular biomarkers and bioparticles, an in-flow based detection scheme is preferred. In a typical in-flow detection scheme, bioparticles are transported to the sensing region in a stream of gas or liquid where they are detected in transient fashion as they pass an optical interrogation region [7, 8]. Therefore, fast read-out of the optical signal from single bioparticles in sequence can be achieved, and many concerns associated with traditional surface-based sensing schemes such as unwanted nonspecific binding, probe photobleaching, and diffusion-limited transportation are eliminated.
Anti-resonant reflecting optical waveguides (ARROWs) have been proven to be highly efficient in detecting single bioparticles. By properly designing a Fabry–Perot reflector surrounding a hollow channel, light can propagate inside the ARROWs. Therefore, ARROWs confine both liquid and light in the same microfluidic channel, such that light and matter have near-perfect overlap and the sensing capability is maximized [8, 9]. Figure 1(a) shows a cross-section of a liquid-core ARROW using state-of-the-art fabrication technology [9]. Moreover, a two-dimensional photonic sensing platform can be constructed with lithography patterning. Figure 1(b) shows an ARROW platform with solid-core and liquid-core ARROWs crossing orthogonally. Excitation light from an external laser is confined in the solid-core ARROW, producing a few-micron-wide optical mode in the intersecting region. Liquid flow is generated inside the liquid-core ARROW to transport the bioparticles to the excitation volume which is of the order of femtolitres for typical waveguide and channel dimensions of a few micrometers. Optical read-out is extracted orthogonally through the liquid-core ARROW to achieve a low-noise signal, sufficient for reaching single particle fluorescence detection.
Besides the optical sensing aspect, miniaturizing and optimizing sample preparation is equally important in order to achieve a complete bioanalysis detection system. The ARROW-based optofluidic system is particularly well suited for such hybrid integration strategies. The planar optical layout based on intersecting solid-core and target-carrying liquid-core waveguides leaves the third dimension open for vertical integration of other functionalities. A separate microfluidic sample processing layer can be made and optimized and then connected to the ARROW platform (Fig. 1c) [10, 11]. Through this approach, we can perform multiple sample preparation steps, such as mixing, distributing, sorting and pre-concentrating on the microfluidic layer and transfer the sample to the ARROW chip for sensing without compromising each of the layers’ performance [10].
Amplification-free detection of Ebola nucleic acids on an opto-
fluidic system
In our recent work, Zaire Ebola virus RNA detection from clinical samples has been demonstrated in a hybrid optofluidic ARROW system [12]. Through a strain-specific solid-phase extraction method, we extracted and labelled target RNA from Ebola infected Vero cells and put them through the optofluidic chip for detection. The ARROW chip provided a sequence of optical signals when individual fluorescent virus RNAs passed through the small excitation volume. Figure 1(d) shows the recorded digital RNA counts at low concentration levels of from 2.1×102 to ~2.1×104 pfu/mL within one second. We were able to detect six orders of magnitude of the clinical concentration range using the ARROW chip only. The lower concentration limit is determined by the detection time, which was set to be 10 min maximum. As a negative control, we used the same method to test for Sudan Ebola virus and Marburg virus. Our results showed no detectable signals and thus our method is target specific.
In order to incorporate critical sample processing steps and detect RNA at even lower concentrations within 10 min, we adapted a programmable microfluidic chip – an automaton – to handle processing of larger sample volumes (Fig. 1b). The polydimethylsiloxane (PDMS) based automaton chip consists of a two-layer microvalve array. Each valve’s state is controlled individually by the top pneumatic layer through a reprogrammable software program. We used the automaton chip to perform an extra pre-concentration step by processing a large amount of clinical sample. We washed, released and labelled the RNA on the same automaton chip after pre-concentration. Through ~460× concentration, the virus detection limit was improved down to 0.2 pfu/mL, with seven orders of magnitude of concentration range (Fig. 1e). This demonstration exhibits an amplification-free chip-based virus and nucleic acid analysis technique with high sensitivity and wide dynamic range, whose performance is comparable with the gold standard, more complex PCR technique.
Wavelength division multiplexing detection
ARROWs also enable simultaneous detection of multiple pathogens through the wavelength division multiplexing (WDM) technique [13]. WDM is generated using a multi-mode interferometer waveguide (MMI). When an MMI is excited by a single optical mode, all of the modes inside the MMI propagate at different phase velocities. When a constructive interference condition is satisfied, various numbers of self-imaging spots which resemble the excitation mode are formed along the MMI. This allows us to design an MMI section that intersects the fluidic channel, where multiple excitation spots are generated (Fig. 2a). As a fluorescent target flows past this excitation region, multi-peak signals are recorded in the time domain. For a single wavelength excitation, the fixed pattern multi-peak detection enables a signal-to-noise improvement compared to single-mode detection [14].
For a given MMI, the number of the self-imaging spots is wavelength dependent. We can generate various spot patterns at various laser wavelengths. For example, 9, 8 and 7 excitation spots are generated using 488nm, 553nm and 633nm lasers (Fig. 2b). With this approach, multiple targets labelled with different dye can be distinguished by the number and spacing of the peaks in the detected signal. Figure 2(c) shows influenza virus H1N1 and H3N2, which were labelled with different dye, generating 9 and 6 peaks in the time domain, respectively. We also labelled H2N2 virus with a combination of these two dyes which resulted in a superposition of the 9-spot and 6-spot fluorescence signals (Fig. 2c, bottom). A signal-processing algorithm checks for the presence of signals at the two characteristic time delays and can easily identify the mixed-labelled virus particle. This technique was shown to discriminate between three influenza subtypes, again with single virus sensitivity, using only two excitation colours. Thus, the ARROW-based platform has now met all the fundamental requirements for clinical virus detection using single particle sensing.
Conclusion
For the next generation of medical diagnostic devices, low-complexity detection with high sensitivity and specificity is required on the detection side, along with small footprint and multi-functional analyte handling on the sample processing side. In-flow based optofluidic devices in which both analyte handling and optical sensing are carried out on the chip scale are promising candidates. Using our ARROW-based optofluidic system, we demonstrated multi-stage sample processing and detection of clinical Zaire Ebola virus samples using hybrid integration. We also demonstrated wavelength multiplex detection of multiple analytes at the same time. This fulfils all quantitative requirements for clinical virus detection. Therefore, a fully integrated microsystem for front-to-back amplification-free virus analysis is within reach.
References
1. Fact sheet. The top 10 causes of death. World Health Organization 2014. (http://www.who.int/mediacentre/factsheets/fs310/en/).
2. Fact sheet. Zika virus. World Health Organization 2016. (http://www.who.int/mediacentre/factsheets/zika/en/).
3. Kuypers J, Wright N, Morrow R. Evaluation of quantitative and type-specific real-time RT-PCR assays for detection of respiratory syncytial virus in respiratory specimens from children. J Clin Virol. 2004; 31: 123–129.
4. Craighead H. Future lab-on-a-chip technologies for interrogating individual molecules. Nature 2006; 442: 387–393.
5. Fan X, White IM. Optofluidic microsystems for chemical and biological analysis. Nature Photon. 2011; 5: 591–607.
6. Schmidt H, Hawkins AR. The photonic integration of non-solid media using optofluidics. Nature Photon. 2011; 5: 598–604.
7. Zhu H, White IM, Suter JD, Zourob M, Fan X. Opto-fluidic micro-ring resonator for sensitive label-free viral detection. Analyst 2008; 133: 356–360.
8. Bernini R, Campopiano S, Zeni L, Sarro PM. ARROW optical waveguides based sensors. Sensors and Actuators B 2004; 100: 143–146.
9. Yin D, Barber JP, Hawkins AR, Deamer DW, Schmidt H. Integrated optical waveguides with liquid cores. Appl Phys Lett. 2004; 85: 3477–3479.
10. Parks JW, Cai H, Zempoaltecatl L, Yuzvinsky TD, Leake K, Hawkins AR, Schmidt H. Hybrid optofluidic integration. Lab Chip 2013; 13: 4118–4123.
11. Testa G, Persichetti G, Sarro, PM, Bernini R. A hybrid silicon-PDMS optofluidic platform for sensing applications. Biomed Opt Express 2014; 5: 417–426.
12. Cai H, Parks JW, Wall TA, Stott MA, Stambaugh A, Alfson K, Griffiths A, Mathies RA, Carrion R, Patterson JL, Hawkins AR, Schmidt H. Optofluidic analysis system for amplification-free, direct detection of Ebola infection. Scientific Reports 2015; 5: 14494.
13. Ozcelik D, Parks JW, Wall TA, Stott MA, Cai H, Parks JW, Hawkins AR, Schmidt H. Optofluidic wavelength division multiplexing for single-virus detection. Proc Nat Acad Sci U S A 2015; 112: 12933–12937.
14. Ozcelik D, Stott MA, Parks JW, Black JA, Wall TA, Hawkins AR, Schmidt H. Signal-to-noise enhancement in optical detection of single viruses with multi-spot excitation, IEEE J Sel Top Quant Elec. 2016; DOI: 10.1109/JSTQE.2015.2503321.
The authors
Hong Cai1 PhD, Aaron R. Hawkins2 PhD, Holger Schmidt*1 PhD
1School of Engineering, University of California Santa Cruz, Street, Santa Cruz, CA 95064 USA
2ECEn Department, 459 Clyde Building, Brigham Young University, Provo, UT 84602 USA
*Corresponding author
E-mail: hschmidt@soe.ucsc.edu
Visual detection of Ebola virus: targeting the NP gene by RT-LAMP
, /in Featured Articles /by 3wmediaEbola virus (EBOV) can lead to severe hemorrhagic fever with a high risk of death in humans and other primates. More recently, reverse transcription loop-mediated isothermal amplification (RT-LAMP) has become readily available for the diagnosis of EBOV, and is a suitable tool for clinical screening, diagnosis and primary quarantine purposes.
by H. Li, W. Lin, X. Wang, X. Wei, E. Li, P. Li, J. Chen, S. Qi, Y. Ma, L. Cui, X. Hu, Dr X. Zhao, Prof. J. Yuan
The 2014 Ebola virus (EBOV; one of the world’s most virulent viruses) caused an outbreak of human disease with widespread transmission in multiple West African countries and sporadic cases in Europe and North America [1, 2]. The numbers of people infected and deaths were the most severe in history. However, the massive public health response has been limited, in part, by the inability to rapidly detect the presence of EBOV in potential patients living in remote areas [3].
EBOV, (species Zaire ebolavirus from the family Filoviridae), was first identified in Zaire in 1976 and named after the River Ebola in Zaire [4]. However, EBOV could not be detected rapidly in many potential patients living in remote and developing areas. The EBOV genome is approximately 19 kb, and encodes the seven proteins in the following order from the 3’-UTR: nucleoprotein (NP), viral structural protein (VSP)35, VSP40, glycoprotein (GP), VP30, VP24, and RNA-dependent RNA polymerase (L) [5]. As the NP gene is highly conserved among EBOV species, it is, therefore, recommended by the World Health Organization (WHO) for use as a target gene for the reverse transcription (RT)-PCR assay. The initial symptoms of EBOV infection could be confused with those of other febrile illnesses such as endemic malaria [6].
Current approaches for the laboratory diagnosis of EBOV infection include virus isolation, electron microscopy, immunohistochemistry, antigen-capture ELISA testing, IgM ELISA, RT-PCR, and serologic testing for IgM or IgG virus-specific antibodies. In 2015, Baca et al. presented a rapid detection of EBOV with a reagent-free, point-of-care biosensor. In general, the detection of EBOV antigens by antigen-capture ELISA is suitable as a method of laboratory diagnosis when the viral load in the blood reaches a very much higher case fatality rate. Thus, real-time (q)RT-PCR has taken over as a first choice diagnostic technique for detection of EBOV recommended by WHO [3]. However, Taq DNA polymerase in PCR-based techniques can be inactivated by inhibitors present in crude biological samples. Moreover, these methods are relatively complex and require specialized high-cost instruments.
Loop-mediated isothermal amplification (LAMP) is a one-step nucleic acid detection method developed by Notomi et al., which relies on autocycling strand displacement DNA synthesis [7]. This novel method is highly specific and sensitive, takes advantage of four or six specific primers to recognize six or eight different sequences of the target gene, and is performed under isothermal conditions in less than 1 h using Bst DNA polymerase. Kurosaki et al. developed a simple reverse transcription (RT)-LAMP assay for the detection of EBOV, targeting the trailer region of the viral genome. However, this method has yet to be tested in clinical samples [8].
To develop an RT-LAMP for clinical screening and rapid diagnosis of EBOV, we first selected potential target regions based on the NP sequences of the EBOV variant Mayinga (GenBank Accession no. AF086833), which were further analysed with Primer Explorer V4 software (http:/primerexplorer.jp/lamp) and subsequently the sequences were aligned with other species of EBOV. A total of five sets of primers were initially designed to detect artificially synthesized EBOV RNA using a real-time turbidimeter. To compare the sensitivity and specificity of RT-LAMP, normal RT-PCR was performed with the primers.
The RT-LAMP reactions were carried out in a 25-μl reaction mixture with an RNA amplification kit (Eiken Chemical Co. Ltd), in accordance with the manufacturer’s protocol. The reaction mixture contained the following reagents (final concentration): RT-LAMP mixture and 8 U Bst DNA polymerase. The amount of primer needed for one reaction was 80 pmol of forward and backward inner primers (FIP and BIP), 40 pmol of loop primer (LB), and 10 pmol of outer forward primer (F3) and outer backward primer (B3). Finally, an appropriate amount of genomic template DNA was added to the reaction tube. The reaction was carried out in the reaction tube at 61 °C, 60–80 min, in dry bath incubators.
Two different methods were used to detect RT-LAMP products. For direct visual inspection, 1 μl of calcein (fluorescent detection reagent; Eiken Chemical Co. Ltd) was added to 25 μl of LAMP products. For a positive reaction, the colour changed from orange to green, whereas a negative reaction remained orange. The colour change could be observed by the naked eye under natural light or with the aid of UV light at 365 nm. For monitoring turbidity, real-time amplification by the RT-LAMP assay was monitored by spectrophotometry, recording the optical density at 650 nm every 6 s with the help of a Loopamp Realtime Turbidimeter (LA-230; Eiken Chemical Co. Ltd) [9].
Assay validation
1. Optimal primer choice and reaction temperature conditions for the RT-LAMP assay
As shown in Figure 1A, the EBL-2 primer set amplified the NP gene using the shortest time of about 10min; therefore, this was chosen as the optimal primer set for EBOV detection of RT-LAMP (Table 1). To further optimize the amplification, reaction temperatures were compared ranging from 59 °C to 69 °C at 2 °C intervals. Ultimately, 61 °C was chosen as the optimal reaction temperature (Fig. 1B).
2. Specificity of NP detection by RT-LAMP using the artificial in vitro transcribed RNA
Twenty-five other non-EBOV viruses were also tested. As shown in Figure 2, the EBOV RNA was identified positively by a successful RT-LAMP reaction with EBL-2 primer set using both methods of analysis. All non-EBOV strains tested negative, including the blank control, indicating that the RT-LAMP method was specific for EBOV.
3. Sensitivity of NP detection by RT-LAMP
A 10-fold serial dilution of artificial EBOV RNA was tested by real-time turbidity monitoring (Fig. 3A), visual detection method (Fig. 3B), and qRT-PCR (Fig. 3C). The limit of detection by the visual method was 10-fold lower compared with the qRT-PCR assay.
4. Clinical sample detection
The 417 clinical blood or swab samples were analysed by RT-LAMP and qRT-PCR simultaneously. The RT-LAMP and qRT-PCR detections both showed that 307 patients were confirmed cases of EBOV infections and 106 patients tested negative for EBOV.
Summary
Zaire ebolavirus is a key member of the Filoviridae family and causes highly lethal hemorrhagic fever in human beings with extremely high morbidity and mortality. As a typical negative-sense single-stranded RNA (ssRNA) virus, EBOV possesses a nucleoprotein (NP) to facilitate genomic RNA encapsidation to form a viral ribonucleoprotein complex (RNP) together with genome RNA and polymerase, which plays the most essential role in virus proliferation cycle. EBOV is found in Central Africa, but re-emerged in Western Africa in 2014 to cause an outbreak that threatened to spread worldwide. Up until 10 January 2016, 28 601 total cases (including suspected, probable, and confirmed) and 11 300 deaths were reported in Guinea and Sierra Leone (http://www.cdc.gov/vhf/ebola/outbreaks/2014-west-africa/case-counts.html). Although several chemical agents, antibodies and vaccines are found to inhibit EBOV in animals or humans, there is no therapeutic with high efficacy that can be provided for clinical usage.
To combat the increasing incidence of EBOV infections, we developed and optimized a novel RT-LAMP assay specific for EBOV diagnosis using primers spanning the 663 bp NP sequence of the viral genome. In the RT-LAMP assay, the reverse transcription reaction and DNA amplification proceed in a single step and with incubation of the reaction mixture at a constant 61°C temperature for a given time period using a temperature-controlled water bath (or other devices that can provide a stable heat are also sufficient). Moreover, LAMP reaction primers specifically recognize five independent regions of the target sequence, compared to PCR primers that recognize two independent regions of the target sequence. The sensitivity of the PCR reaction can be greatly reduced by the presence of exogenous DNA and inhibitors. Therefore, the RT-LAMP method is more suitable for rapid detection of NP in clinical samples.
Conclusion
In conclusion, a specific, sensitive, rapid and cost effective RT-LAMP assay for NP detection in EBOV was established, which is as sensitive as other available technologies, highly specific and extremely rapid in the provision of molecular diagnosis of EBOV infections. The assay can provide accurate results in a short time frame. This makes it potentially useful for clinical diagnosis of EBOV in developing countries.
Acknowledgment
This article is based on one previously published by the authors: Li H, Wang X, Liu W, Wei X, Lin W, Li E, Li P, Dong D, Cui L, Hu X, Li B, Ma Y, Zhao X, Liu C, Yuan J. Survey and Visual detection of Zaire ebolavirus in clinical samples targeting the nucleoprotein gene in Sierra Leone. Frontiers in Microbiology 2015; 6: 1332 [10].
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2. Hampton T. Largest-ever outbreak of Ebola virus disease thrusts experimental therapies, vaccines into spotlight. JAMA 2014; 312(10): 987–989.
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4. MacNeil A, Rollin PE. Ebola and Marburg hemorrhagic fevers: Neglected tropical diseases? PLoS Negl Trop Dis. 2012; 6(6): e1546.
5. Ali MT, Islam MO. A highly conserved GEQYQQLR epitope has been identified in the nucleoprotein of Ebola virus by using an in silico approach. Adv Bioinformatics 2015; 2015: 278197–278203.
6. Grolla A, Lucht A, Dick D, Strong JE, Feldmann H. Laboratory diagnosis of Ebola and Marburg hemorrhagic fever. Bull Soc Pathol Exot. 2005; 98(3):205–209.
7. Notomi T, Okayama H, Masubuchi H, Yonekawa T, Watanabe K, Amino N, Hase T. Loop-mediated isothermal amplification of DNA. Nucleic Acids Res. 2000; 28, E63.
8. Kurosaki Y, Takada A, Ebihara H, Grolla A, Kamo N, Feldmann H, Kawaoka Y, Yasuda J. Rapid and simple detection of Ebola virus by reverse transcription-loop-mediated isothermal amplification. J Virol Methods 2007; 141(1): 78–83.
9. Mori Y, Nagamine K, Tomita N, Notomi T. Detection of loop-mediated isothermal amplification reaction by turbidity derived from magnesium pyrophosphate formation. Biochem Biophys Res Commun. 2001; 289: 150–154.
10. Li H, Wang X, Liu W, Wei X, Lin W, Li E, Li P, Dong D, Cui L, Hu X, Li B, Ma Y, Zhao X, Liu C, Yuan J. Survey and Visual detection of Zaire ebolavirus in clinical samples targeting the nucleoprotein gene in Sierra Leone. Frontiers in Microbiology 2015; 6: 1332.
The authors
Huan Li# MMed, Weishi Lin# MMed, Xuesong Wang MMed, Xiao Wei MMed, Erna Li MMed, Puyuan Li MMed, Jun Chen MMed, Silei Qi MMed, Yanyan Ma MMed, Lifei Cui MMed, Xuan Hu MMed, Xiangna Zhao PhD, Jing Yuan PhD*
Institute of Disease Control and Prevention, Academy of Military Medical Sciences, Beijing, 100071, PR China
#These authors contributed equally to this work
*Corresponding author
E-mail: yuanjing6216@163.com
Biochemical markers of alcohol intake
, /in Featured Articles /by 3wmediaBiochemical markers of alcohol intake can be separated into two categories: direct markers of ethanol metabolism and indirect markers. The different alcohol markers have varying time windows of detection and are a useful additional tool to detect alcohol intake in alcohol-dependent clients.
by Jane Armer and Rebecca Allcock
Introduction
Alcohol dependence is characterized by craving, tolerance, a preoccupation with alcohol and continued drinking in spite of harmful consequences. The World Health Organization Alcohol Use Disorders Identification Test (AUDIT) is recommended for the identification of individuals that are dependent on alcohol [1]. The prevalence of alcohol use disorders (including dependence and harmful use of alcohol) is 11.1% in the UK compared to 7.5% across Europe [2]. In England, 250 000 people are believed to be moderately or severely dependent and require intensive treatment [3].
Alcohol use is the third leading risk factor contributing to the global burden of disease after high blood pressure and tobacco smoking [4]. In 2012, 3.3 million deaths (5.9% of all global deaths) were attributable to alcohol consumption [2]. It is estimated that the UK National Health Service (NHS) spends £3.5 billion/year in costs related to alcohol and the number of alcohol-related admissions has doubled over the last 15 years [3].
In the UK, one unit equals 10 mL or 8 g of pure alcohol, which is around the amount of alcohol the average adult can process in an hour. The latest UK recommendations are to not regularly drink more than 14 units per week (men and women) and to limit the total amount of alcohol consumed on a single occasion [5].
The most common entry into alcohol treatment services in England is either self-referral or referral by the GP [3]. Services have a limited number of options to determine if an individual in treatment for alcohol dependence is continuing to drink alcohol. They rely on self-report by the individuals in the form of alcohol diaries and breathalyser tests. There is no regular schedule for biochemical markers. If a client is found to be drinking alcohol during the treatment programme, an assessment is done of the amount of alcohol consumed, the pattern of alcohol consumption and how it will impact on their treatment. This is factored into the recovery plan and there is a re-assessment of the support and interventions needed for that client. Possible interventions include cognitive behavioural therapies, pharmacological therapies or in-patient assisted withdrawal. In 2013/14, only 38% of clients in alcohol treatment in England successfully completed their treatment [3].
Monitoring clients in alcohol treatment
Diaries that record alcohol intake are commonly used to monitor the progress of clients. However, this relies on accurate self-reporting of alcohol intake by the client and under reporting is a common problem. Biochemical markers of alcohol intake can provide a more comprehensive assessment of a client’s progress.
Direct markers of alcohol intake
Direct markers of alcohol intake include ethanol, ethyl glucuronide (EtG), ethyl sulphate (EtS), fatty acid ethyl esters (FAEE) and phosphatidylethanol (PEth).
Following the ingestion of ethanol, >95% is metabolized in the liver by alcohol dehydrogenase to acetaldehyde then by aldehyde dehydrogenase to acetic acid [14]. Less than 5% is excreted unchanged in the urine, breath and sweat. A small amount of ethanol is conjugated to form EtG and EtS (Fig. 1). Ethanol is usually only detectable in breath and urine after very recent alcohol consumption and the detection time window depends on the amount of alcohol consumed. In comparison, urine EtG and EtS remain detectable for around 24 hours after moderate alcohol intake and for up to 130 hours in subjects admitted for alcohol detoxification [6, 7]. Liquid chromatography-tandem mass spectrometry (LC-MS/MS) methods have been developed for EtG and EtS. An immunoassay is also available for EtG [8, 9].
Many studies have demonstrated the benefit of measuring EtG and EtS in clients in alcohol treatment. Continued alcohol consumption can be detected by the measurement of urine EtG and EtS in clients who do not admit to consuming alcohol and provide a negative breathalyser test. This is due to the increased time window of detection for urine EtG and EtS compared to breath ethanol. This demonstrates the unreliability of self-reporting of alcohol intake and the benefit of biochemical markers to detect clients that are continuing to drink alcohol [10].
As with urine testing for drugs of abuse, it is possible for a client to consume a large volume of water to dilute the sample and produce negative EtG and EtS results. Creatinine should always be measured to check for adulteration and it may be beneficial to report EtG and EtS as creatinine ratios to overcome this problem. Further work is required to define cut-offs for EtG and EtS as creatinine ratios.
False negative EtG results can be caused by the presence of Escherichia coli in urine as glucuronidase is present with high activity in most strains. False positive EtG and EtS results have also been reported following use of ethanol based mouthwash or hand gels and after the consumption of non-alcoholic beers (up to 0.5% alcohol). Due to the risk of positive results due to unintentional alcohol exposure, particularly for urine EtG, it is important that clinical cut-offs used are clearly defined and LC-MS/MS methods that measure both EtG and EtS are preferred [11]. In the USA, the Substance Abuse and Mental Health Administration (SAMHSA) have suggested that EtG results >1.0 mg/L are consistent with alcohol intake and that results between 0.1 and 1.0 mg/L should be interpreted with caution. It is accepted that further work is required to clearly define cut-offs for EtG and EtS and that other biomarkers may be useful when interpreting borderline positive results in the range 0.10–0.50 mg/L [12].
Methods for the measurement of EtG and FAEEs in hair have been developed allowing a longer term assessment of alcohol intake. Hair analysis is most suitable for subjects where longer term abstinence needs to be demonstrated such as in patients awaiting liver transplantation. EtG cut-offs have been suggested by the Society of Hair Testing for chronic excessive alcohol consumption (30 pg/mg) and abstinence assessment (7 pg/mg). However, results may be influenced by hair products and this needs to be taken into account when interpreting results.
PEth is formed from ethanol and phosphatidylcholine in cell membranes. The reaction is catalysed by phospholipase D and occurs in the cell membranes of erythrocytes; therefore, PEth is found in the red blood cell fraction of blood rather than in serum or plasma. PEth is a group of phospholipids with varying carbon lengths and LC-MS/MS methods to detect the major forms of PEth in whole blood have been developed. A single dose of ethanol does not produce a measurable amount of PEth and it has been demonstrated that approximately 50 g of ethanol/day (6.25 UK units) is required to provide a positive PEth result. In comparison to serum carbohydrate deficient transferrin (CDT; see ‘Indirect markers of alcohol intake’ below), urine EtG and urine EtS, PEth demonstrated the highest sensitivity for regular alcohol consumption in clients in alcohol treatment and was found to be positive twice as often as CDT [13]. Further work is required to understand how PEth can be used optimally in combination with other alcohol markers in clients in treatment for alcohol dependence [14].
Indirect markers of alcohol intake
The indirect markers include mean corpuscular volume (MCV), gamma glutamyl transferase (GGT) and CDT. These markers increase following significant alcohol intake over a prolonged time period and are not useful for detecting a single alcohol ‘binge’. MCV and GGT are not specific markers of alcohol intake.
CDT refers to altered glycoforms of transferrin as a result of alcohol-induced changes in the carbohydrate composition of transferrin. The main component of serum transferrin is tetrasialotransferrin, which makes up approximately 80% of the total. Normal samples usually contain approximately 15%, 4–5%, 1–1.5% and 1% of pentasialotransferrin, trisialotransferrin, disialotransferrin and hexasialotransferrin, respectively. An alcohol consumption of at least 60 g/day (7.5 UK units) for 2 weeks is required to increase the disialotransferrin [15]. CDT may also be increased if genetic variants are present and in advanced liver disease. The International Federation of Clinical Chemistry and Laboratory Medicine (IFCC) has recently proposed a reference measurement procedure for CDT and more studies assessing the diagnostic performance of CDT to detect alcohol dependence are now needed using methods harmonized to the international reference measurement procedure.
Table 1 summarizes the time window of detection and limitations of the alcohol markers discussed.
Conclusions
Currently, the assessment of clients in alcohol treatment relies largely on self-reporting and limited biochemical testing, which makes assessment of a client’s progress challenging. There are a number of available biochemical markers that could improve the detection of alcohol use in clients with alcohol dependence and ultimately lead to initiation of early intervention and altered treatment strategies. This in turn could improve the numbers successfully completing treatment. A combination of short-term and longer term biochemical markers is likely to be the most useful approach depending on the treatment setting. The advantage of the breathalyser test over biochemical markers that require laboratory analysis is the immediate availability of the result which allows an immediate intervention for a client with a positive result. Laboratory tests need to be available in a timely manner and with appropriate and well-defined cut-offs. The clinical benefit of alcohol markers in improving the number of clients that successfully complete their treatment for alcohol dependency has not yet been demonstrated. Randomized controlled trials comparing outcomes with or without the use of biochemical markers are required.
References
1. Babor TF, Higgins-Biddle JC, Saunders JB, Monteiro MG. Alcohol use disorders identification test (AUDIT). World Health Organization, 2001. (http://www.alcohollearningcentre.org.uk/Topics/Browse/BriefAdvice/?parent=4444&child=4896)
2. Global status report on alcohol and health. World Health Organization, 2014. (http://www.who.int/substance_abuse/publications/global_alcohol_report/msb_gsr_2014_2.pdf?ua=1)
3. Alcohol Treatment England 2013–14. Public Health England, 2014. (http://www.nta.nhs.uk/uploads/adult-alcohol-statistics-2013-14-commentary.pdf )
4. Lim S, Vos T, Flaxman A, Danaei G, Shibuya K, Adair-Rohani H, Amann M, Anderson HR, Andrews KG, et al. A comparative risk assessment of burden of disease and injury attributable to 67 risk factors and risk factor clusters in 21 regions, 1990-2010: a systematic analysis for the Global Burden of Disease Study 2010. Lancet 2012; 380: 2224–2260.
5. UK Chief Medical Officers’ Alcohol Guidelines Review. Department of Health, 2016. (https://www.gov.uk/government/uploads/system/uploads/attachment_data/file/489795/summary.pdf)
6. Dahl H, Stephanson N, Beck O, Helander A. Comparison of urinary excretion characteristics of ethanol and ethyl glucuronide. J Anal Toxicol. 2002; 26: 201–204.
7. Helander A, Bottcher M, Fehr C, Dahmen N, Beck A. Detection times for urinary ethyl glucuronide and ethyl sulphate in heavy drinkers during alcohol detoxification. J Anal Toxicol. 2009; 44: 55–61.
8. Politi L, Morini L, Groppi A, Poloni V, Pozzi F, Polettini A. Direct determination of the ethanol metabolites ethyl glucuronide and ethyl sulphate in urine by liquid chromatography/electrospray tandem mass spectrometry. Rapid Commun Mass Spectrom. 2005; 19: 1321–1331.
9. Bottcher M, Beck O, Helander A. Evaluation of a new immunoassay for urinary ethyl glucuronide testing. Alcohol Alcohol. 2008; 43: 46–48.
10. Junghanns K, Graf I, Pfluger J, Wetterling G, Ziems C, Ehrenthal D, Zöllner M, Dibbelt L, Backhaus J, Weinmann W, Wurst FM. Urinary ethyl glucuronide (EtG) and ethyl sulphate (EtS) assessment: valuable tools to improve verification of abstention in alcohol-dependent patients during in-patient treatment and at follow ups. Addiction 2009; 104: 921–926.
11. Wurst F, Thon N, Yegles M, Schruck A, Preuss UW, Weinmann W. Ethanol metabolites: their role in the assessment of alcohol intake. Alcohol Clin Exp Res. 2015; 39: 2060–2072.
12. The role of biomarkers in the treatment of alcohol use disorders. SAMHSA, 2012. (http://store.samhsa.gov/product/The-Role-of-Biomarkers-in-the-Treatment-of-Alcohol-Use-Disorders-2012-Revision/SMA12-4686)
13. Helander A, Peter O, Zheng Y. Monitoring of the alcohol biomarkers PEth, CDT and EtG/EtS in an outpatient treatment setting. Alcohol Alcohol. 2012; 47: 552–557.
14. Viel G, Boscalo-Berto R, Cecchetto G, Fais P, Nalesso A, Ferrara SD. Phosphatidylethanol in blood as a marker of chromic alcohol use: a systematic review and emta-analysis. Int J Mol Sci. 2012; 13: 14788–14812.
15. Stibler H. Carbohydrate Deficient Transferrin in serum: a new marker of potentially harmful alcohol consumption reviewed. Clin Chem. 1991; 37: 2029–2037.
The authors
Jane Armer*1 BA MSc FRCPath and
Rebecca Allcock2 BSc MSc FRCPath
1Department of Blood Sciences,
East Lancashire Hospitals NHS Trust,
Blackburn, UK
2Department of Clinical Biochemistry,
Lancashire Teaching Hospitals NHS
Foundation Trust, Preston, UK
*Corresponding author
E-mail: jane.armer@elht.nhs.uk
Meeting clinical needs with high performance viral load assays, workflow improvements and reduced turnaround times
, /in Featured Articles /by 3wmediaNiguarda Hospital in Milan is one of Italy’s leading general hospitals, and provides an extensive range of medical disciplines for adults and children throughout the Lombardy region and beyond.
Our hospital’s Department of Laboratory Medicine aim is to offer a complete, continuous and prompt diagnostic laboratory testing service, in order to guarantee effective support for this widespread clinical demand, and is committed to research into automation and analysis to ensure this is maintained. Our busy Molecular Biology Laboratory performed an estimated 40,000 tests in 2015, which is approximately 10% increase on the previous year.
The growing annual molecular workload is attributed, in part, to the development of new therapeutic strategies. Our staff, consisting of 8 laboratory technicians, one director and one manager, work 5 days per week and are expected to cope with increased workloads and demands for reduced turnaround times without any increase in resources, in terms of the number of staff and costs.
A large proportion of the molecular biology workload consists of viral load measurements for human immunodeficiency virus type 1 (HIV-1), hepatitis C virus (HCV), hepatitis B virus (HBV) and cytomegalovirus (CMV) (figure 1).
With a very important Italian transplant centre located at Niguarda Hospital, CMV analyses are vital and results are needed quickly, without delay. In addition, the laboratory performs viral load measurements for HIV-1, HCV, and HBV in order to evaluate and monitor therapeutic response. In these instances, rapid results are extremely important for patient management decisions, for example to maintain or change treatment.
Since 2005, these measurements have been performed using our laboratory’s current method, which has separate sample preparation and amplification/detection platforms. These are situated on separate benches within the same room, with one sample preparation system in another room. The accuracy and precision of this method is good, however, in order to be cost effective, it is necessary to optimize the size of the batches. Since they can’t be processed in the same day, sample test tubes often need to be collected and stored for several days, which increases the turnaround time considerably. In addition, this method involves many manual steps, which demand time, space and coordination of work between different members of staff.
A new automated molecular diagnostics method
As part of our Department of Laboratory Medicine’s investigations into increased automation in the laboratory, Niguarda Hospital became a beta trial site for the new DxN VERIS Molecular Diagnostics System (Beckman Coulter), which consolidates DNA extraction, nucleic acid amplification, quantification and detection onto a single automated instrument for a number of molecular targets, including HIV-1, HCV, HBV and CMV.
The first step in assessing the DxN VERIS was to validate the assays in order to determine whether their performance is comparable with our laboratory’s existing method. Daily quality control measurements demonstrated good performance of the VERIS HBV assay for high level, low level and negative HBV samples (table 1). This assay was also shown to have excellent linearity within the range of 1.68 – 8.82 Log IU/mL, a limit of detection of 6.82 IU/mL, and good precision, achieving within run and between run mean standard deviations of less than 0.16 (table 2).
A series of performance evaluation studies, conducted in several laboratories around the world, have demonstrated that the VERIS HBV, HCV, HIV-1 and CMV assays have comparable precision, sensitivity and linearity to a range of alternative, commercially available viral load methods [1-13]. In accordance with these findings, the VERIS HBV assay correlated well with the existing method at Niguarda Hospital (Abbott m2000) and, indeed, detected HBV DNA in 23 samples that were negative using the current method, 22 of which were found to be positive by one or more serology assay (table 3). Regarding the 55 specimens that were quantified both with DxN VERIS and Abbott m2000, 7 of them had an HBV DNA concentration discordant for more than 1 Log.
Comparable performance, including sensitivity and specificity, was achieved for each of the DxN VERIS assays: HIV-1, HCV, HBV and CMV.
Workflow improvements
In addition to validating the performance of the VERIS assays, a time/workflow analysis study was performed at Niguarda Hospital by Nexus Global Solutions (Plano, Texas, USA). The study compared workflows and time to results between the current viral load method for HIV-1, HCV, HBV and CMV (Abbott m2000sp and m2000rt systems) and the new DxN VERIS Molecular Diagnostic System.
By reducing manual intervention and automating processes from sample loading to reporting of results, the DxN VERIS offers the potential to transform clinical laboratory workflows. Each assay is supplied in a unique, single cartridge system, and all consumables and reagents are stored on board the system, which cuts preparation time compared to alternative methods. In addition, unlike traditional plate-based systems, there is no need to batch assays. The DxN VERIS allows true, single sample random access, which means that viral load assays can be performed as soon as they arrive in the laboratory. This, combined with short assay runtimes, ensures rapid turnaround of results and, since there are no empty plate wells, wastage and consumable costs are reduced.
The comparative time/workflow analysis in our study revealed that DxN VERIS involved only 10 steps and required just five reagents, compared to 26 steps and over 20 consumables for the current method, and required much less hands-on time for each of the viral load assays (figure 2). Notably, by consolidating the assay menu, time savings of up to 2 hours could be achieved.
In addition to an increase in productivity (achieving more results in an 8-hour working day), the time to the first result for the DxN VERIS was greatly reduced compared to the current method, with subsequent results available every 2.5 minutes. This is in contrast to the current method, where results are not available until the end of the assay run (table 4).
With these time savings, and by eliminating the need to batch samples, the DxN VERIS allowed much faster turnaround of results in a normal working week, with all results being reported within 8 hours of receipt, unlike the current method, which often required several days (figure 3).
The true single sample random access capability of the DxN VERIS has the potential to simplify sample management in the laboratory and to make the organization of viral load assays more fluid. It increases productivity by allowing the continuous loading of samples for different assays, eliminating the need for batching and reducing turnaround times. This is the most important advantage of random access testing for us because it increases the availability of medical reports to the different departments and is a great benefit to patient management and care by allowing more timely clinical decisions.
The DxN VERIS is easy to use with its few consumables, reduced maintenance requirements, complete automation and intuitive computer interface. By improving laboratory organization and workflows and reducing manual intervention, viral loads (which account for about 50% of the molecular workload) could be completed in a single day using the DxN VERIS. Requiring fewer people to be dedicated to this purpose, this makes it possible to accomplish more work with the same number of staff.
For further information about the DxN VERIS Molecular Diagnostic System and the VERIS assays currently available, please contact: Tiffany Page, Senior Pan European Marketing Manager Molecular Diagnostics, Email: info@beckmanmolecular.com or visit www.beckmancoulter.com/moleculardiagnostics
References
1. Williams, JA, Rodriguez, J, Wang, Z et al (2014) Poster presentation, ESCV, Prague.
2. Drago, M, Franchetti, E, Fanti, D and Gesu, GP (2015) Poster presentation, EuroMedLab, Paris.
3. Zurita, S, Gutiérrez, F, Folgueira, MD et al (2015) Poster presentation, EuroMedLab, Paris.
4. Christenson, R, Maggert, K, Ruiz, RM et al (2015) Poster presentation, ECCMID, Copenhagen.
5. Trimoulet, P, Tauzin, B, Belloc, E et al (2015) Poster presentation, EuroMedLab, Paris.
6. Gilfillan, R, Wang, Z, Xu, Y et al (2014) Poster presentation, ECCMID, Barcelona.
7. Xu, Y, Gilfillan, R, Wang, Z et al (2014) Poster presentation, ESCV, Prague.
8. Mengelle, C, Sauné, K, Haslé, C et al (2014) Poster presentation, RICAI.
9. Mengelle, C, Sauné, K, Haslé, C et al (2015) Poster presentation, ECCMID, Copenhagen.
10. Silvestro, A, Duan, H, Lim, S et al (2014) Poster presentation, ECCMID, Barcelona.
11. Li, Q, Williams, J, Maggert, K et al (2014) Poster presentation, ECCMID, Barcelona.
12. Xu, Y, Dineen, S, Annese, V et al (2014) Poster presentation, ESCV, Prague.
13. Williams, JA, Rodriguez, J, Wang, Z et al (2014) Poster presentation, ECCMID, Barcelona.
The author
Diana Fanti, Molecular Biology Laboratory Manager
Department of Laboratory Medicine, Niguarda Hospital,
Milan, Italy
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