Article:Find the right sample: A study on the versatility of saliva and urine samples for the diagnosis of emerging viruses (6311079)

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Journal Information

Title: BMC Infectious Diseases

Find the right sample: A study on the versatility of saliva and urine samples for the diagnosis of emerging viruses

  • Matthias Niedrig
  • Pranav Patel
  • Ahmed Abd El Wahed
  • Regina Schädler
  • Sergio Yactayo

Publication date (epub): 12/2018

Publication date (pmc-release): 12/2018

Publication date (collection): /2018



The emergence of different viral infections during the last decades like dengue, West Nile, SARS, chikungunya, MERS-CoV, Ebola, Zika and Yellow Fever raised some questions on quickness and reliability of laboratory diagnostic tests for verification of suspected cases. Since sampling of blood requires medically trained personal and comprises some risks for the patient as well as for the health care personal, the sampling by non-invasive methods (e.g. saliva and/ or urine) might be a very valuable alternative for investigating a diseased patient.

Main body

To analyse the usefulness of alternative non­invasive samples for the diagnosis of emerging infectious viral diseases, a literature search was performed on PubMed for alternative sampling for these viral infections. In total, 711 papers of potential relevance were found, of which we have included 128 in this review.


Considering the experience using non-invasive sampling for the diagnostic of emerging viral diseases, it seems important to perform an investigation using alternative samples for routine diagnostics. Moreover, during an outbreak situation, evaluation of appropriate sampling and further processing for laboratory analysis on various diagnostic platforms are very crucial. This will help to achieve optimal diagnostic results for a good and reliable case identification.

Electronic supplementary material

The online version of this article (10.1186/s12879-018-3611-x) contains supplementary material, which is available to authorized users.



The emergence of several virus infections during the last decades like dengue, West Nile (WN), Severe acute respiratory syndrome (SARS), chikungunya (CHIK), Middle East respiratory syndrome coronavirus (MERS-CoV), Ebola (EBO), Zika (ZIK) and Yellow Fever (YF) raised some questions on quickness and reliability of laboratory diagnostic tests for verification of suspected cases. All important questions arising with an emerging infection like source and route of transmission, infectivity of patients, recommendation of safety measures for health care personal and distribution in an animal reservoir or other vectors require the performance of laboratory diagnosis. To investigate the patient’s infections most often a blood sample is taken not only for the analysis of blood parameters but also for pathogen detection mostly by PCR or for testing the presence of specific antibodies. However, sampling of blood requires medically trained personal and comprises some risks for the patient as well as for the health care personal. In particular the blood sampling of patients with highly infectious haemorrhagic fever like Ebola involve a great risk for the physician and the patient. Therefore, the sampling by non-invasive methods (e.g. saliva and/ or urine) might be a very valuable alternative for investigating a diseased patient. The use of saliva and urine for the diagnosis for many viral infections has been reported, however, it is often not performed in the broad routine diagnostics.

After a viral infection it usually takes some days until the propagation of the virus causes symptoms and a clinical investigation of a patient is indicated. Blood sampling for analysing blood parameters belongs to a routine procedure in developed countries. This is different in developing countries, where extensive analyses of clinical parameters are not performed routinely due to financial and technical constraints. Even though the sampling of swaps from the respiratory tracks is a common procedure for influenza, SARS, MERS-CoV and other respiratory infections, the sampling of blood is the prevalent procedure for all other recently emerged viruses. However, with the introduction of the highly sensitive Nucleic Acid Tests (NAT) for diagnostic also additional samples like saliva and/or urine were included in diagnostic sample collections. The advantage of using non-invasive samples like saliva for dengue serology was successfully proven 18 years ago by Cuzzubbo et al. who discovered that dengue primary and secondary infections can be distinguished by the level of IgG in saliva [[1]]. The specificity of the diagnosis of acute flavivirus infections like dengue, WN and YF has been improved by applying the NAT because it is not impeded by the widespread cross-reactivity found for most serological assays [[2]]. Since urine, saliva and other non-invasive samples were not part of the routine investigation, the sampling procedures and the samples containment are also very different. These differences complicate the interpretation of diagnostic findings and therefore an optimization for these samplings is required. An improvement of the sampling and stabilization of viral RNA in the diagnostic material will help to increase significantly the diagnostic sensitivity.

To analyse the usefulness of alternative non-invasive samples for diagnostic of infectious viral diseases like SARS, WN, dengue, MERS-CoV, Ebola, Zika and YF, we reviewed the importance of saliva and urine for the diagnosis of recently emerging viruses in comparison to conventional blood sampling. Since the extensive evaluation of different samples for human patients is often not performed, also investigations of animal models or vectors are considered. Therefore, we performed an electronic literature review on PubMed for alternative sampling for SARS, WN, dengue, MERS-CoV, Ebola, Zika and YF.

Main text

A review of the literature for urine and saliva samples for diagnostic of CHIK, dengue, Ebola, WN, Zika, YF, SARS and MERS was performed using the electronic database PubMed. The period for searches was 1st January 1980 – 26th August 2016.

Appropriate articles were selected according to their abstracts and further reviewed for reason for diagnostic analysis, sample types used and the methods applied for diagnosis focusing on saliva and urine as test samples.

Search profile used for this review in PubMed for CHIK as an example:

"chikungunya virus"[MeSH Terms] OR ("chikungunya"[All Fields] AND "virus"[All Fields]) OR "chikungunya virus"[All Fields] AND ("saliva"[MeSH Terms] OR "saliva"[All Fields])

"chikungunya virus"[MeSH Terms] OR ("chikungunya"[All Fields] AND "virus"[All Fields]) OR "chikungunya virus"[All Fields] AND ("urine"[Subheading] OR "urine"[All Fields] OR "urine"[MeSH Terms])

Another 42 references were considered for analysing the preparation and storage of diagnostic samples as demonstrated in Table 1, see Additional file 1 for detailed methodology.Table 1

Overview of selected publications describing samples used for viral diagnostic

Type of specimens Arboviruses VHFV Coronaviruses
Blood derived samples whole blood [[139]] [[143]] [[147], [148]] n.r. [[88]] [[160]] [[165]] [[170]]
peripheral blood (cells) [[139]] [[144]] [[147], [148]] n.r. n.r. [[54]] [[166]]
peripheral blood (plasma) [[140]] [[13]] [[149]] [[152]] [[89]] [[54]] [[112], [166]]
seruma [[141]] [[144]] [[149]] [[152]] [[97]] [[52]] [[167]] [[169]]
umbilical cord blood n.r. n.r. n.r. n.r. n.r. n.r. n.r. n.r.
Body fluids saliva [[19]] [[13]] n.r. ? [[88], [89], [97]] [[52]] n.r. n.r.
urine [[19]] [[19]] [[75]] [[104]] [[88], [89], [97]] [[54]] [[110], [112]] [[124]]

VHF Viral haemorrhagic Fever, a = NAT & Immunological test, n.r. not reported

The literature research revealed 711 papers, of which we considered 128 in this study (Additional file 1: Figure S1). Furthermore 43 publication analysing the sample processing and preparation were included.

Our analysis shows that there should be put more emphasis on non-invasive sampling to disclose more easy alternatives for collecting human samples for clinical and routine investigations. The present review provides some indications that non-invasive sampling offers a promising alternative for routine diagnostic as well as a favourable diagnostic approach for surveillance studies with high numbers of samples. The review provides an overview of the usability of non-invasive sample for diagnostics of emerging viruses and tries to give recommendation on how this experience could be helpful for future possible outbreaks of emerging viruses.


After an epidemic at La Reunion in 2005, an autochthone outbreak in Italy in 2007 and, spread in Caribbean islands in December 2013, chikungunya (CHIK) emerged in 2014 for the first time in Central America spreading into South America and North America [[3][7]]. Typical clinical specimens for CHIKV are blood, serum and plasma, but also urine, saliva and in rare cases CSF (Table 1, Additional file 1: Table S1). A systematic evaluation of the CHIK infection in mice and Cynomolgus macaques revealed the infectivity of different body fluids like saliva, urine and vaginal swaps [[8]]. This could be confirmed by analysis of saliva samples from 4 out of 13 acute CHIKV patients with haemorrhagic manifestations, which were found to contain viral RNA and infectious virus. Another study shows that RNA of chikungunya virus can be detected in the saliva at the first seven days of infection but with a lower efficacy than blood samples [[9]]. Also the detection of CHIKV genome by RT-PCR in urine was successful in one returning traveller, which is a valuable confirmation of the positive serology [[10]]. Based on the experience with other viral infections, the analysis of throat swabs and urine are part of recommendation for CHIK diagnostic [[11]].


Common clinical specimens used for dengue infection are blood, plasma and serum, but also in some cases CSF and breast milk (Table 1, Additional file 1: Table S2). Interestingly, the twenty-two selected publications described very well the progress of using saliva and urine as non- invasive samples. Based on the existing IgG ELISA for serology, they found that the IgG levels in saliva could be applied to differentiate between dengue primary and secondary infection [[1]]. One year later, Artimos de Oliveira et al., used saliva for detection of specific IgM of recent infections [[12]]. In 2000 Torres et al., found for the first time evidence of DENV RNA in saliva and plasma by RT-PCR [[13]]. The evaluation of saliva samples as alternative approaches for serological diagnosis (IgM, IgG, IgA) was further investigated and complemented by more systematic kinetics analysis of 1st and 2nd infections also comprising urine as an additional alternative sample [[14], [15]]. Since the sensitivity (93.3%), specificity (100%), positive predictive value (100%), and negative predictive value (83.3%) of the detection of IgG in saliva are high, it was considered as promising sample type for dengue diagnostics [[16]]. Also in case reports analysing acute infections, the detection of DENV in urine and saliva by RT-PCR as well as the application of filter-paper for saliva sampling prove the usefulness of these non-invasive samples [[17][19]]. The utility of saliva in an assay that detects DENV-specific IgA in the early phase of a 2nd dengue infection shows 100% sensitivity from day-one after fever onset with a good correlation to IgA levels in serum and 97% specificity [[20]]. DEN NS1 antigen was detected in urine in around two third of dengue fever and DEN haemorrhagic fever (HF) using ELISA. Follow up the excretion of NS1 in urine is very important particularly in patients developing DF or DHF as an indication of protein leakage. The study demonstrates that the NS1 strip assay is less sensitive for urine compared to the ELISA and both assays need further adaptation to improve the sensitivity [[21]]. This was confirmed by a later study from Saito et al. showing that urine samples could be used for NS1 antigen ELISA but only in case of serum samples are very limited or in poor resource settings [[22]].

From 2012, more studies were performed using saliva and urine samples besides serum for quantitative RT-PCR and other parameters (NS1, IgM, IgG) for dengue diagnostic during the course of the disease [[23][25]]. Further studies underline the benefit of saliva as a diagnostic tool for use in developing tropical countries like India nevertheless further research is encouraged before implementation [[26]]. The delayed release of DENV in urine allows the diagnosis of an acute dengue infection by RT-PCR and sequencing on day 8 and 18, while the respective serum samples collected on the same period were negative for DENV [[27]].

All these promising results using urine and saliva samples for diagnostics stimulated further investigations on DENV RNA stability in urine and serum as well as establishing novel diagnostic device (stacking flow platform) for single-step detection of a target antibody in salivary fluid [[28], [29]]. Both studies investigated different conditions using urine or saliva samples for the diagnostics. One study focused on the storage conditions for these samples while the other improved the sample processing to optimize diagnostics using saliva. Andries et al. performed a representative study by analysing plasma, saliva and urine in 267 patients by applying various molecular and serological analyses. Unfortunately, the performances of these assays were better upon using plasma samples [[30]]. Analysing commercial rapid diagnostic tests (RDT) for detection of NS1/IgG/IgA in urine and saliva samples suggested that these RDT kits are of higher specificity and limited sensitivity [[31]]. Interestingly, in dengue secondary infection, the IgA in urine at 4-7 days of disease onset is higher than those with 1st infections indicating severe Dengue [[32]]. Therefore, DENV IgA in urine is an indicator for the disease severity and must be investigated.

Ebola and other viral hemorrhagic fevers (VHFs)

Ebola virus and other viral hemorrhagic fevers are mostly detected in blood derived samples such as whole blood, plasma or serum. Recent studies reported Ebola detection also in body fluids including saliva, urine, sweat, tears, semen and vaginal discharge as well as feces, ammonitic fluids, placenta and breast milk (Table 1 and Additional file 1: Table S3).The search for alternative sampling for Ebola yielded 34 publications from which 11 were performed during the recent Ebola outbreak from 2015/2016. Other publication were found for other VHF viruses like Hanta, 9 hits; Marburg, 4; Junin, 3; VHFs in general 2; Lassa, 2; Crimean Congo hemorrhagic fever (CCHF), 2; Rift Valley fever (RVF), 1; Guanarito, 1 and 1 for a novel Bunya virus.The first report dated from 1980 for RVF virus detection in saliva based on experimentally infected sheep and showed successful virus isolation from saliva [[33]]. This was completed by the extensive analysis of infected rodents as main transmission vectors for Hantavirus [[34]]. Besides virus excretion, Hanta virus could be demonstrated in several organs (lung, kidney, salivary glands and liver) by virus isolation from saliva, urine and feces for up to one year post inoculation. Also the intensive investigation of an imported Lassa case to the UK demonstrated the high viral load in blood taken eleven days post onset of the disease as well as significant virus count in urine on day 36, while throat swabs did not yield any detectable virus [[35]]. These findings were confirmed during the intensive analysis of an acute Lassa virus infected patient when the use of urine as additional sample to serum was successfully investigated for viral RNA by PCR [[36]].

Vereta and colleagues used urine and urine sediment epithelium for the identification of acute Hanta virus patients showing hemorrhagic fever with renal syndrome (HFRS) [[37], [38]]. Urine becomes an important sample for serological studies since antibody excretion in the urine coincided with the period of renal structure damage and stopped when the normal renal function was restored. The first successful Hantavirus isolation of a fatal HFRS case used urine and brain tissue and was further confirmed by RT-PCR [[39]]. Henceforward, urine was used for antigen and genome detection as well as for serology in experimentally infected pigs demonstrating that pigs can be the host for the viral transmission of HFRS virus [[40]]. Petterson et al. could demonstrate that hantavirus RNA was found in saliva few days after disease onset [[41]]. The association between the anti-Hantavirus IgA and Hantavirus RNA in saliva was inverse, the same is true for detecting RNA and antigen in endothelial cells within the parotid gland of HFRS patients [[42]]. In a recent study, the anti-Hantavirus antibodies in saliva was detected in a risk group in farms in Yorkshire, UK [[43]]. Hereby, the non-invasive saliva sampling confirms the usefulness of Hantavirus surveillance studies for analyzing the presence of Hantavirus in previously unknown risk areas. Also a recent investigation of Hanta virus persistence in wild rodents is based on the analysis of viral RNA in saliva, urine and feces show a peak during the first month after seroconversion continuing throughout the 8 month study [[44]].

Urine and saliva as non-invasive samples were also successfully used for analyzing the pathogenesis of Junin virus in the rodent vector responsible for virus transmission for up to 480 days post infection [[45], [46]]. Surviving animals showed a viral dissemination in brain, spleen, kidney and salivary glands with half of the animals shedding Junin virus and the other half being negative for infectious virus, while seroconverted. These findings were successfully used for analyzing the prevalence of Junin virus in the rodent population in endemic areas defining risk areas for the Junin virus infection [[47]].

Non-invasive samples like urine were used for the analysis of guinea pig and Rhesus macaques infectivity experiments with highly infectious and deadly Marburg virus [[48]]. In particular for dangerous biosafety level 4 pathogens the sampling of blood is very risky and laboratory infections are possible, which can be avoided by sampling urine. In the guinea pig model, the persistent shedding of Marburg virus in saliva, urine and feces showed that as early as by the end of incubation period and throughout the disease, the virus could be found in the feces and saliva virtually in the same concentrations [[49]], while in the blood the content of the virus was high and increased by the end of the disease. A similar approach was used investigating bat fecal and urine samples for Marburg virus after infection of a tourist visiting a cave in Uganda [[50]]. Around 2.5% of liver/spleen tissues of captured bats were positive in RT-PCR for Marburg virus. Moreover, the virus was detected in 15 different tissues and plasma of Egyptian fruit bats subcutaneously inoculated with Marburg virus, while limited results were obtained applying mucosal swab samples, urine and fecal samples [[51]].

Antibodies against Ebola virus could not be found in oral fluids, while 100% agreement between the presence of RNA in serum and oral fluids by RT-PCR was observed [[52]]. Moreover, in 26 confirmed Ebola cases, 16 samples including saliva, stool, semen, breast milk, tears, nasal blood and a skin swab were positive in Ebola culture and/or RT-PCR [[53]]. In case of EBOV, all samples including body fluids must be handled applying the WHO recommendations, since they are highly infectious.The recent Ebolavirus outbreak in Serra Leone, Liberia und Guinea with 11,325 deaths initiated several intensive investigations of different patient specimens like blood, urine, sweat, saliva, conjunctival swaps, stool and semen for infectious virus [[54], [55]]. These became a major interest since transmission routes were not always obvious from previous experience. Moreover, public health measures require clear references, when recovered patients are not infectious anymore and can be released from quarantine. The excretion of EBOV in body fluids from infected or recovered patients create a great risk for person in contact [[54], [56]] as the Ebola virus RNA was detected in the following body fluids weeks up to several months after infection: saliva, conjunctiva/tears, stool, vaginal fluid, sweat, urine, amniotic fluid, aqueous humor, cerebrospinal fluid, breast milk, and semen raised many concerns regarding the long term transmissibility of infectious virus causing a recrudesce of an outbreak [[56][58]]. Comparing whole-blood with urine specimen’s analysis from Ebola patients in a novel diagnostic film-array with RT-PCR shows a reasonable match of 90% for whole blood and 85% for urine [[59]]. Testing the stability of Ebola virus RNA in human blood and urine under environmental African conditions shows that viral RNA testing from blood samples stored in EDTA buffer is more sensitive [[60]]. Viral RNA in urine seems less stable compared to blood since urine samples were found positive by RT-PCR by day 10-14 compared to blood until at least day-18. Since the CCHF outbreak in Turkey in 2002, the detection of virus in other body fluids received attention to avoid secondary infections. RNA of CCHF virus was identified in the saliva and the urine of 5/6 and 2/3 patients, respectively [[61]]. This was confirmed by a later study, which detected CCHF virus in urine samples of patients with prolonged viremia [[62]].

Furthermore, nasopharyngeal aspirates and/or urine were considered as diagnostic samples evaluating a multiplex hybridization array for detecting various infectious diseases [[63]]. Interestingly, the Plasmodium falciparum was identified as a cause of death in a case of hemorrhagic fever like illness during the Marburg hemorrhagic fever outbreak in Angola in 2004-2005. Also a novel diagnostic array based on beads technology for detection of multiple bat-borne viruses including Ebola and Nipha viruses used urine from wild bats from Australia and Bangladesh for a surveillance study [[64]].

Analyzing cases in China of a hemorrhagic fever due to a novel Bunya virus, the RNA was identifed in the blood as well as in urine, throat, and fecal specimens [[65]]. Further studies are necessary to investigate the significance of non-invasive samples for analyzing the virus pathogenesis and the infectivity of different body fluids.

West Nile

Clinical specimens used for WNV detection are blood derived samples and CSF (Table 1 and Additional file 1: Table S4). Since the safety of blood banks was important, testing of blood samples was the evident choice. Even though Steele et al. found infectious WN in several organs (brain, heart, spleen, liver, kidney, adrenal intestines, pancreas, lung and ovary) of birds, it took four years until Tesh and others published the investigation of urine samples used for pathogenesis studies in hamsters and birds [[66][69]]. These findings were later confirmed in different animal models like chipmunks, fox squirrels, hamsters and mice analyzed for WNV pathogenesis research [[70][74]]. The first report of WN detection in urine was in an acute severe case with encephalitis found positive eight days after symptoms onset [[75]]. Further studies detected WNV RNA up to 6.7 years in urine indicating for a persistent renal infection for several years [[76]]. This was confirmed by a number of studies demonstrating the long persistence of WNV particularly in patients with WN fever compared to patients with WN neuro-invasive disease, which had a higher vireamia for a shorter period than those with West Nile fever [[77][80]]. In the meantime, the advantage of using urine instead or additionally to serum is demonstrated by many investigations and could prove its value also for routine diagnostic [[81][86]]. Also WNV could be successfully isolated from urine samples and further analyzed by sequencing.

In summary, one could state that the use of urine is a suitable alternative diagnostic sample compared to serum and should be listed in fact sheets from national and international organizations (WHO, PAHO, CDC, ECDC). Further studies are needed investigating if urine samples are a useful source for surveillance studies for WNV distribution and for analyzing the severity of the disease in patients with neuro-invasive and/or kidney manifestation.


The Zika virus was detected in blood, serum, saliva, and urine, as well as in semen, vaginal discharge, sweat, tears, ammonitic fluids, placenta and breast milk (Table 1 and Additional file 1: Table S5). The expansion of Zika to Brazil via New Caledonia 2015 stimulates the development of diagnostic assays for NAT and serology to analyze the increasing number of cases [[87]]. In a surveillance study, ZIKV was identified in 19.2% (total 182 patients) of saliva samples, but not in blood [[88]]. The detection of ZIKV in saliva samples increased the molecular detection rate of ZIKV in acute case but ZIKV did not persist for longer time frame in saliva as in urine or semen. Nevertheless, saliva sample was advantageous in children and neonates [[89]]. Interestingly, the ZIKV can be detected in urine samples for even longer. Combination of urine and saliva besides serum is of great significant in ZIKV diagnostics [[90]]. This was also true in a rhesus macaque model used for analyzing plasma, saliva, urine and cerebrospinal fluid by RT-PCR for ZIKV detection [[91]].

Moreover, RNA of ZIKV was detected in semen by RT-PCR on the 7th day of the appearance of symptoms but was not identified on the 21st day, which was confirmed by Reusken et al. [[92], [93]]. This finding consequently leads to the possibility of an infection via sexual intercourse as documented by D'Ortenzio et al. with a high viral load in the semen obtained on sample collected after 18 days [[94], [95]]. In several smaller and bigger studies, the usability of saliva and urine beside blood samples was demonstrated for the Zika virus diagnostics [[96][98]]. The excretion of viral RNA in urine and saliva was observed for up to 29 days after symptom onset and with higher viral load than in blood [[99], [100]]. The high risk for transmission by patient’s body fluids also raised concern for appropriate safety measures for health care personal as discussed in several publications [[101][103]].

Yellow Fever

Yellow Fever is generally detected in blood, plasma, serum and CSF as other flaviviruses (Table 1 and Additional file 1: Table S6). Although alternative samples demonstrate their usefulness for other flavivirus infections like urine or saliva samples for diagnostics of dengue or West Nile, YF were not investigated until 2011 [[104], [105]]. These are the first reports showing the usefulness of urine samples for diagnostic of adverse events after YF vaccination. For vaccinees developing severe side effects after vaccination it could be clearly shown that viral shedding is prolonged up to day 25 post vaccination. The release of YF RNA could be found in urine at day 198 after vaccination. If the detection of the YF attenuated 17D vaccine virus could be demonstrated in urine easily. However, representative data of alternative samples like urine or saliva from YF infections are missing. In particular for cases with a severe course of disease with numerous hemorrhagic bleedings the use of non-invasive sampling would be a real alternative preventing bodily injury of the patient.

Severe acute respiratory syndrome (SARS)

When SARS emerged in November 2002, it was obvious to analyze samples from the respiratory tract for diagnostic. However, even for the first SARS patients other samples beside blood and nasopharyngeal aspirate samples like feces and urine were found positive with 97% and 42% for viral RNA, respectively [[106]] (Table 1 and Additional file 1: Table S7). This was confirmed by analysis of throat wash and saliva showing a high virus load up to 6x106 and 6x108 RNA copies per ml, respectively [[107]]. Moreover, the detection of SARS in plasma, sputum, endotracheal aspirates, stool, throat swabs and saliva revealed significant differences between the types of samples [[108]] as all samples from the lower respiratory tract were tested positive. Cheng et al. found that death was associated with higher virus load in nasopharyngeal specimens obtained on day 10 after the onset of symptoms [[109]]. Different samples like serum, nasopharyngeal aspirates (NPA), throat swabs, nasal swabs, rectal swab, stool and urine were used for analysis of the viral pathogenesis during the course of the disease [[109][112]]. The diarrhea and hepatic dysfunction was associated high viral load in NPA, while in urine is associated with abnormal urinalysis findings. The extensive analysis of 415 SARS patients revealed the presence of SARS in respiratory specimens two weeks after disease onset, and in stool, rectal swab and urine specimens for up to four weeks. Throat and nasal swabs is less significant that the NPA specimens, therefore, the high viral load in NPA has a major impact on the airborne transmission, which played a major role during the outbreak in Hong Kong [[113]].

The evaluations of commercial PCR kits and in-house antigen-ELISAs for the quality of SARS diagnostic are an important task to select the most appropriate test for the selected sample. The sensitivity of RT-PCR is higher compared to two antigen-ELISAs [[114]] as RNA of SARS-CoV can be detected earlier in fecal specimens in around 80% of patients and in 25% of urine samples. Based on all these findings, RT-PCR is the method of choice for early diagnosis of SARS CoV infection [[115], [116]].

The co-presence of viral RNA and antibodies in plasma was observed over three weeks in two cases [[117]]. In addition, the SARS-CoV was isolated from stool or urine specimens for longer than 4 weeks [[118]]. Since the detection of SARS virus is possible in various clinical samples (NPA, throat-swab, fecal, cerebrospinal fluid, blood and urine) by viral culture or RT-PCR clear and significant recommendations for diagnostic of SARS based on the previous experiments worked out by the WHO and the Multi-Centre Collaborative Network require an appropriate update. Even though SARS did not appear any more after the outbreak except for the two laboratory infections thereafter [[119][171]].

Bats are the most likely animal vector for CoVs [[121]]. Also experiments in mice and rhesus macaque analyzing the viral pathogenesis for the development of effective strategies for diagnosis, prevention, therapy and vaccine design was further evaluated [[122], [123]]. Since salivary gland epithelial cells were affected as a first target after infection the immunization with virus like particles might be eliciting a protective immune response against SARS-CoV generating a mucosal immunity. This new vaccine approach provides important information for future vaccine design.

Middle East respiratory syndrome coronavirus (MERS-CoV)

A boost of nearly 50 publications on alternative sampling within three years demonstrates the importance and documents the public health concern with this respiratory disease caused by a new Corona virus (CoV) (Additional file 1: Table S8). The publications can be roughly categorized as follows 11 case reports, 7 descriptions of pathogenesis, 11 on recommendation and investigation of diagnostics, 7 on recommendation and analysis of risks for health care workers, 2 recommendations for patient management and 5 on control and prevention including treatment and vaccine development. Only a few selected publications were considered for this review. Bronchoalveolar and lower respiratory tract fluids were highly positive for MERS-CoV in the two identified cases [[124]], while urine samples were only positive on day 13. Stool samples and oro-nasal swabs contained very low MERS-CoV copy number on days 12 and 16, respectively, and no virus was detected in blood. In the beginning of the outbreak the way of transmission was rather unclear. Therefore, the virus replication in the kidney with potential shedding in urine was further investigated [[125]]. To clarify the different transmission routes a prediction model for oral-fecal and/or oral-saliva transmission routes for MERS-CoV was developed and evaluated [[126]]. Very soon camels as contributing animal host for MERS-CoV infection got into focus of investigation and it became obvious that also dromedary camels in Eastern Africa and the Arabian Peninsula have a very high seroprevalence of MERS-CoV antibodies [[127]]. For further surveillance studies in camels, hedgehogs, and bats very often non-invasive samples like urine, saliva, fecal nasal swaps, fecal swaps or camel milk were used to analyze for MERS-CoV RNA or specific antibodies [[128]]. In summer 2015, 183 confirmed cases of MERS and 33 fatalities were reported in the Republic of Korea caused by a nosocomial outbreak [[129]].

The timing and intensity of respiratory viral shedding in MERS-CoV infected patients is still of major interest as airborne transmission from human to human by droplets seems to be the dominant infection route [[130][132]]. It might be that the introduction of the isothermal amplification (LAMP or RPA) for detecting of emerging diseases provides a useful tool for quick and reliable diagnostic of acute cases and might lead to an immediate public health response including the appropriate safety precautions for the medical personal [[133], [134]].

Based on the experience with other respiratory infections and based on the lessons learned from the previous SARS outbreak the analysis of respiratory samples are parts of the routine samples taken for diagnostic of suspected MERS patients.


The analysis of using other non-invasive samples for virus diagnostics clearly shows that the procedures for the optimal sampling strategy of the different viral infections are not seriously investigated table S9. In the following overview it became obvious that for the newly emerging viruses like CHIK, MersCoV and Zika more and broader attempts for sampling strategies were performed and investigated than for well known viruses like YF and dengue.

The application of saliva as a sample of choice for the diagnosis of CHIK fever infections in small children or patients whose blood samples are difficult to obtain is of great significant. However, all recommendations/fact sheets for CHIK from national and international organizations (CDC, PAHO, WHO, ECDC) do not mention saliva or urine samples as suitable material for diagnosis because of the missing of meaningful studies that evaluate saliva and urine as diagnostic samples compared with blood. Hopefully, there will be studies performed answering these questions with a representative number of CHIK patients in the future.

The use of saliva and urine for the diagnosis of acute dengue infections is of great importance and requires more attention. Present studies show the usefulness for these samples for diagnostics and also the monitoring of the severity of infection. In particular for surveillance studies and/or investigation of small children the noninvasive approach using saliva or urine for the laboratory diagnosis of dengue cases should be considered when blood samples are difficult to obtain. Although the application of alternative samples for the diagnostic became more popular in the recent years, further studies and improvements are necessary to establish these samples equally to common plasma or serum samples. Commercial assays as well as appropriate sampling equipment have to be verified for diagnostic applicability in representative cohorts and during the course of the disease. It seems that the increased awareness of alternative samples stimulates such studies for dengue diagnosis as demonstrated in the recent publications mentioned above.

In particular for diagnostic of VHF cases, the investigation of the usefulness of non-invasive taken samples should have high priority since blood sampling from such patients is highly risky and often dangerous for patients with hemorrhagic symptoms while continuous bleeding occurs. The virus load in sever cases of VHF is very high in body fluid and the NAT is the method of choice since the production of specific IgM and/or IgG antibodies takes a few days. In very severe cases, the patient even can pass away before developing a detectable antibody reaction, while a general viremia affecting several organs and is detectable in most body fluids. In less severe cases, the exact moment becomes an important issue since viremia only remains for just a few days (3-4 days) and often patient samples were taken too late for successful NAT thus requiring laborious serology testing with in-house assays and risk of false positive reactivity. Onset of disease is also not a well-defined point of time because the perception of a disease might be different in developing compared to developed countries. This makes the comparison of clinical symptoms and diagnostic findings during the course of the disease even more difficult.

Although the analysis of different body fluids during the recent Ebola virus outbreak expands our knowledge of virus transmissibility via eye fluid and semen tremendously, systematic studies on which and for how long the different body fluids will contain viral RNA and/or infectious viruses are still missing. Recommendations on how non-invasive sampling should be performed, what kind of samples are necessary and how they should be stabilized and transported under African environmental conditions are still missing and require thoroughly investigations.

For WNV infections the use of urine is a suitable alternative diagnostic sample compared to serum and should be listed in fact sheets from national and international organizations. Further studies are needed investigating if urine samples are a useful source for surveillance studies on WNV distribution and for analyzing the severity of the disease in patients with neuro-invasive and/or kidney manifestation. The emergence of Zika virus tremendously stimulated the investigation of using non-invasive samples as saliva and urine for diagnostic of Zika virus infections. From the start of the Zika outbreak in French Polynesia saliva was considered as alternative material for the diagnostics. The investigation of urine and saliva shows a higher virus load compared to blood making these samples a suitable alternative to blood. However, this information has to be forwarded to the physicians as it is already part of the PAHO and WHO recommendations.

For YF diagnostic one can hope that the integration of saliva and urine as alternative samples into the recommendation worked out by the WHO for the recent outbreak will help to implement this method into the medical procedure and create some meaningful results for future outbreak scenarios.

Even although SARS seems to have disappeared from the globe, the experience and findings on which and when the different diagnostic samples should be used for a quick and reliable diagnostic should be carefully evaluated, since respiratory infections are on the rise as we have seen a few years later by the influenza and MERS-CoV outbreaks. The great variety on which, how and when respiratory samples have to be taken is reflected by the list of nomenclature found in the publications: for upper respiratory specimens (nasal swab, exudate nose, oronasal swab, pharyngeal swab, tracheal swab, exudate mouth, saliva and sputum) and lower respiratory specimens (bronchoalveolar fluid, bronchoalveolar lavage). Recommendations for standardized procedures for sampling and handling of the respiratory samples will help to allow a better comparison of the findings and provide a better preparedness for the next respiratory disease outbreak.

Because of a higher virus load, samples from the lower respiratory tract are more suitable for diagnostic. However, the sampling of bronchoalveolar-lavage fluid is also invasive and requires an experienced physician and holds a certain risk of nosocomial infections by aerosols. Other non-invasive samples like urine, nasal, pharyngeal, tracheal and rectal swabs were included in the diagnostic analysis for analyzing the pathogenesis and the kinetic of virus release by the different body fluids. Like for SARS standardized procedures for sampling and handling of the respiratory samples will help to allow a better comparison of the findings and provide a better preparedness for future respiratory disease outbreak.

The present review clearly shows that there is still the need to perform more diligent studies analyzing non-invasive patient samples in comparison with blood and serum samples, which used for routine diagnostic of emerging viral diseases. While for some diseases like the respiratory infections, the sampling of nasal swaps and saliva is part of routine sampling, for other viral infections, blood or serum seems the predominant and only type of sample used for laboratory diagnosis.

This is in particular disappointing because non-invasive sampling offers a real alternative avoiding additional risk for the patient and the medical personal. Patients suspected to be infected by VHF usually have a very high virus load. Therefore, the analysis of saliva or urine should be routinely performed in parallel to investigation of blood samples. Based on a solid database it will be possible to determine the most appropriate sample for a viral diagnosis before it gets implemented in the routine diagnostic procedure. Diagnostic laboratories and physicians have to gain knowledge when and how alternative samples have to be considered for analyzing an acute or convalescent viral infection. Even though, we have with NAT a very sensitive diagnostic tool, the best point in time during the course of the disease together with the optimal sampling, sample storage and sample preparation are essential to get optimal results for a laboratory diagnosis. In patients presenting with a moderate or mild disease, the viremia is mostly very short lasting just 1-3 days. For these patients the positive NAT is sometimes difficult to achieve and a negative finding is of no predictive value since the virus titer might be just below the detection limit or sample storage and sample preparation is not optimal in case of non-invasive sampling such as saliva and urine. Although the virus disappeared from the blood, the virus genome will be still detectable in the urine as consequence of the clearance process as we have found in Yellow Fever vaccinees or other for Zika [[97], [104]]. In contrast, we find a high viral load in patients developing a severe disease sometimes with a general infection affecting several organs. In particular in those patients, the detection of the pathogenic virus by NAT is not difficult analyzing different types of samples. Very often the viral load also reflects the severity of the disease and can be used as prognostic marker for progression of the disease in the patient. Exactly for these patients the non-invasive sampling is less stressful as frequent bleeding performed with the current procedure. This becomes even more important, if point of care diagnostic allows a more frequent monitoring of infection parameters of the diseased patient in the future. For the recently emerging CHIK and Zika in South America, the analysis of throat swaps, saliva and urine samples were already analyzed for their usability in diagnostics. However, the findings are based on case studies or small studies and require a more systematic approach. Preferably there should be performed a systematic analysis of different invasive samples (blood, bronchoalveolar lavage) and non-invasive samples (saliva, urine, etc.) for their diagnostic value using different diagnostic methods for patients with acute and convalescent disease when new infections emerge. This also implies that the procedure taking samples has to be somehow standardized. This became very obvious by the large number of terms for upper and lower respiratory samples most likely reflecting the great variety how the samples were taken. This makes the comparability of diagnostic findings presented in different studies very difficult. To address this problem a clear recommendation for the sampling procedure would help to overcome this imponderability. Furthermore, the methods for storage of the samples regarding temperature and stabilization needs to be worked out to allow an optimal transport and preparation of the diagnostic sample. For example there are special sampling tubes with stabilizer available for both saliva (DNAgard Saliva by biomatrica, USA) and urine (VACUETTE Urine CCM tubes by Greiner Bio-One, Germany and Urine Monovette by Sarstedt, Germany), which will help preserves the integrity of sample at room temperature. Sample preparation from urine and saliva could be also challenging especially for molecular diagnostics, therefore it is very important to optimize protocols using clinical samples rather than spiked mock-samples. Our personal experience showed that the optimized sample preparation method on real clinical samples was not as efficient as on spiked mock samples.

The increasing number of publications considering alternative sampling for diagnostic of the recent outbreaks MERS-CoV, CHIK, Ebola and Zika are promising. However, systematic studies are rare and for some infectious disease outbreaks like the recent yellow fever in Angola, non-invasive samples for diagnostic are still uncommon and require further investigation. Just for diseases like yellow fever, non-invasive samples like urine or saliva might be a very practical alternative, since the first index patients are detected often very late, while blood will be negative by NAT in patients with mild disease and from patients with a severe disease it is difficult taking blood because of a hemorrhage clinical picture. Non-invasive samples might be also a great alternative for investigations of children or other patients were collection of blood samples is difficult [[18], [30], [88]]. Also for large surveillance studies, non-invasive samples like saliva and/or urine could be performed without medical personal for collecting blood. However, the usability of these samples for serology or NAT has to be evaluated in comparison to conventional use of sera. Therefore, it is necessary to introduce non-invasive sampling strategies in recommendations as done for the “Yellow Fever Laboratory Diagnostic Algorithm” worked out for the epidemic and non-epidemic countries in Africa during the YF outbreak in Angola by WHO expert team [[135]]. Since these investigations could be only performed during an outbreak situation the preparation of a scheme for evaluation of sampling for respiratory as well as for other infectious diseases including the further processing for laboratory diagnostic is highly recommended. This more structured approach will help to improve the diagnostic of emerging infectious diseases significantly as new diagnostic methods and platforms are developed continuously and require effective evaluation under real conditions.

For confirmation of a clinical diagnosis a correct laboratory diagnosis is essential for all further steps of patient's treatment and handling. In particular for new and/or emerging diseases, the conditions for the best sampling regarding time and source (blood, serum, saliva, urine, etc.) are often not known. Since every disease has its own course of symptoms and affected organs, the optimal sampling schema might change and require different strategies regarding sample preparation and processing. While in the early phase when disease symptoms appear often the viral pathogen could be detected by nucleic acid testing in blood or other body fluids in the later phase only antibodies are detectable in serum.

In the present article, we analyzed over 700 scientific publications on diagnostic approaches for dengue, West Nile, SARS, chikungunya, MERS-CoV, Ebola, Zika and Yellow Fever regarding their relevance for using alternative samples like saliva and urine for their versatility for virus diagnostic. The use of non-invasive samples offers a suitable alternative for improving the diagnostic performance and significantly reduces the risk for medical personal and patients for routine diagnostic and during an outbreak situation. The advantage of using non-invasive samples for the diagnostic has to be seriously analyzed and investigated before implementing in the routine diagnostics.

Additional file

Additional file 1:

Figure S1. Selection scheme flowchart. Abbreviation: CHIV = Chikungunya Virus, VHF = Viral Hemorrhagic Fever, WN = West Nile, YF = Yellow fever. Table S1. Analysis of different sampling methods for diagnostic of CHIK virus infection. Table S2. Analysis of different sampling methods for diagnostic of Dengue virus infection. Table S3. Analysis of different sampling methods for diagnostic of Ebola/VHF virus infections. Table S4. Analysis of different sampling methods for diagnostic of West Nile virus infection. Table S5. Analysis of different sampling methods for diagnostic of Zika virus infection. Table S6. Analysis of different sampling methods for diagnostic of Yellow Fever virus infection. Table S7. Analysis of different sampling methods for diagnostic of severe acute respiratory syndrome (SARS) virus infection. Table S8. Analysis of different sampling methods for diagnostic of Middle East respiratory syndrome coronavirus (MERS-CoV) infection. (DOCX 162 kb)



The authors thank Véronique Millot from the World Health Organization for help with the literature search.


No specific funding was obtained for preparing the manuscript.

Availability of data and materials

All publications evaluated for the analysis are provided in the supplement part of the manuscript.


  1. AJ CuzzubboDW VaughnA NisalakS SuntayakornJ AaskovPL DevineDetection of specific antibodies in saliva during dengue infectionJ Clin Microbiol19983612373737399817913
  2. P KorakaH ZellerM NiedrigAD OsterhausJ GroenReactivity of serum samples from patients with a flavivirus infection measured by immunofluorescence assay and ELISAMicrobes Infect20024121209121510.1016/S1286-4579(02)01647-712467761
  3. M VazeilleS MoutaillerD CoudrierC RousseauxH KhunM HuerreJ ThiriaJS DehecqD FontenilleI SchuffeneckerTwo Chikungunya isolates from the outbreak of La Reunion (Indian Ocean) exhibit different patterns of infection in the mosquito, Aedes albopictusPLoS One2007211e116810.1371/journal.pone.000116818000540
  4. AM PowersCH LogueChanging patterns of chikungunya virus: re-emergence of a zoonotic arbovirusJ Gen Virol200788Pt 92363237710.1099/vir.0.82858-017698645
  5. G RezzaL NicolettiR AngeliniR RomiAC FinarelliM PanningP CordioliC FortunaS BorosF MaguranoInfection with chikungunya virus in Italy: an outbreak in a temperate regionLancet200737096021840184610.1016/S0140-6736(07)61779-618061059
  6. F SimonP ParolaM GrandadamS FourcadeM OliverP BrouquiP HanceP KraemerA Ali MohamedX de LamballerieChikungunya infection: an emerging rheumatism among travelers returned from Indian Ocean islands. Report of 47 casesMedicine (Baltimore)200786312313710.1097/MD/0b013e31806010a517505252
  7. DL VanlandinghamC HongK KlinglerK TsetsarkinKL McElroyAM PowersMJ LehaneS HiggsDifferential infectivities of o'nyong-nyong and chikungunya virus isolates in Anopheles gambiae and Aedes aegypti mosquitoesAm J Trop Med Hyg200572561662110.4269/ajtmh.2005.72.61615891138
  8. J GardnerPA RuddNA ProwE BelarbiP RoquesT LarcherL GreshA BalmasedaE HarrisWA SchroderInfectious Chikungunya Virus in the Saliva of Mice, Monkeys and HumansPLoS One20151010e013948110.1371/journal.pone.013948126447467
  9. D MussoA TeissierE RouaultS TeururaiJJ de PinaTX NhanDetection of chikungunya virus in saliva and urineVirol J20161310210.1186/s12985-016-0556-927306056
  10. M KondoS AkachiK AndoT NomuraK YamanakaH MizutaniTwo Japanese siblings affected with Chikungunya fever with different clinical courses: Imported infections from the Cook IslandsJ Dermatol201643669770010.1111/1346-8138.1325326813362
  11. CG RautH HanumaiahWC RautUtilization and Assessment of Throat Swab and Urine Specimens for Diagnosis of Chikungunya Virus InfectionMethods Mol Biol20161426758310.1007/978-1-4939-3618-2_727233262
  12. S Artimos de OliveiraCV RodriguesLA CamachoMP MiagostovichES AraújoRM NogueiraDiagnosis of dengue infection by detecting specific immunoglobulin M antibodies in saliva samplesJ Virol Methods.1999771818610.1016/S0166-0934(98)00139-610029328
  13. JR TorresF LiprandiAP GoncalvezAcute parotitis due to dengue virusClin Infect Dis2000315E28E2910.1086/31745411073786
  14. A BalmasedaMG GuzmanS HammondG RobletoC FloresY TellezE VideaS SaborioL PerezE SandovalDiagnosis of dengue virus infection by detection of specific immunoglobulin M (IgM) and IgA antibodies in serum and salivaClin Diagn Lab Immunol200310231732212626461
  15. S VazquezS CabezasAB PerezM PupoD RuizN CalzadaL BernardoO CastroD GonzalezT SerranoKinetics of antibodies in sera, saliva, and urine samples from adult patients with primary or secondary dengue 3 virus infectionsInt J Infect Dis200711325626210.1016/j.ijid.2006.05.00516914345
  16. A ChakravartiM MatlaniM JainImmunodiagnosis of dengue virus infection using salivaCurr Microbiol200755646146410.1007/s00284-007-9040-517899259
  17. A BalmasedaS SaborioY TellezJC MercadoL PerezSN HammondC RochaG KuanE HarrisEvaluation of immunological markers in serum, filter-paper blood spots, and saliva for dengue diagnosis and epidemiological studiesJ Clin Virol200843328729110.1016/j.jcv.2008.07.01618783984
  18. Y MizunoA KotakiF HaradaS TajimaI KuraneT TakasakiConfirmation of dengue virus infection by detection of dengue virus type 1 genome in urine and saliva but not in plasmaTrans R Soc Trop Med Hyg2007101773873910.1016/j.trstmh.2007.02.00717418320
  19. TR PoloniAS OliveiraHL AlfonsoLR GalvaoAA AmarillaDF PoloniLT FigueiredoVH AquinoDetection of dengue virus in saliva and urine by real time RT-PCRVirol J201072210.1186/1743-422X-7-2220105295
  20. G YapBK SilLC NgUse of saliva for early dengue diagnosisPLoS Negl Trop Dis201155e104610.1371/journal.pntd.000104621572982
  21. A ChuansumritW ChaiyaratanaK TangnararatchakitS YoksanM FlamandA SakuntabhaiDengue nonstructural protein 1 antigen in the urine as a rapid and convenient diagnostic test during the febrile stage in patients with dengue infectionDiagn Microbiol Infect Dis201171446746910.1016/j.diagmicrobio.2011.08.02021996098
  22. Y SaitoML MoiA KotakiM IkedaS TajimaH ShibaK HosonoM SaijoI KuraneT TakasakiDetecting Dengue Virus Nonstructural Protein 1 (NS1) in Urine Samples Using ELISA for the Diagnosis of Dengue Virus InfectionJpn J Infect Dis201568645546010.7883/yoken.JJID.2014.44125766601
  23. KL AndersNM NguyetNT QuyenTV NgocTV TramTT GanNT TungNT DungNV ChauB WillsAn evaluation of dried blood spots and oral swabs as alternative specimens for the diagnosis of dengue and screening for past dengue virus exposureAm J Trop Med Hyg201287116517010.4269/ajtmh.2012.11-071322764309
  24. T HirayamaY MizunoN TakeshitaA KotakiS TajimaT OmatsuK SanoI KuraneT TakasakiDetection of dengue virus genome in urine by real-time reverse transcriptase PCR: a laboratory diagnostic method useful after disappearance of the genome in serumJ Clin Microbiol20125062047205210.1128/JCM.06557-1122442323
  25. EM KorhonenE HuhtamoAM VirtalaA KanteleO VapalahtiApproach to non-invasive sampling in dengue diagnostics: exploring virus and NS1 antigen detection in saliva and urine of travelers with dengueJ Clin Virol201461335335810.1016/j.jcv.2014.08.02125242312
  26. S Ravi BanavarGS VidyaDiagnostic efficacy of saliva for dengue - a reality in near future? A piloting initiativeJ Clin Diagn Res20148322923224783144
  27. X MaW ZhenP YangX SunW NieL ZhangH XuK HuFirst confirmation of imported dengue virus serotype 2 complete genome in urine from a Chinese traveler returning from IndiaVirol J2014115610.1186/1743-422X-11-5624666930
  28. D Van den BosscheL CnopsM Van EsbroeckRecovery of dengue virus from urine samples by real-time RT-PCREur J Clin Microbiol Infect Dis20153471361136710.1007/s10096-015-2359-025794553
  29. Y ZhangJ BaiJY YingA stacking flow immunoassay for the detection of dengue-specific immunoglobulins in salivary fluidLab Chip20151561465147110.1039/C4LC01127A25608951
  30. AC AndriesV DuongS LyJ CappelleKS KimP Lorn TryS RosS OngR HuyP HorwoodValue of Routine Dengue Diagnostic Tests in Urine and Saliva SpecimensPLoS Negl Trop Dis201599e000410010.1371/journal.pntd.000410026406240
  31. AC AndriesV DuongS OngS RosA SakuntabhaiP HorwoodP DussartP BuchyEvaluation of the performances of six commercial kits designed for dengue NS1 and anti-dengue IgM, IgG and IgA detection in urine and saliva clinical specimensBMC Infect Dis20161620110.1186/s12879-016-1551-x27184801
  32. H ZhaoS QiuWX HongKY SongJ WangHQ YangYQ DengSY ZhuFC ZhangCF QinDengue Specific Immunoglobulin A Antibody is Present in Urine and Associated with Disease SeveritySci Rep201662729810.1038/srep2729827250703
  33. DG HarringtonHW LuptonCL CrabbsCJ PetersJA ReynoldsTW Slone JrEvaluation of a formalin-inactivated Rift Valley fever vaccine in sheepAm J Vet Res19804110155915647224281
  34. HW LeePW LeeLJ BaekCK SongIW SeongIntraspecific transmission of Hantaan virus, etiologic agent of Korean hemorrhagic fever, in the rodent Apodemus agrariusAm J Trop Med Hyg19813051106111210.4269/ajtmh.1981.30.11066116436
  35. RT EmondB BannisterG LloydTJ SoutheeET BowenA case of Lassa fever: clinical and virological findingsBr Med J (Clin Res Ed)198228563471001100210.1136/bmj.285.6347.1001
  36. K LunkenheimerFT HufertH SchmitzDetection of Lassa virus RNA in specimens from patients with Lassa fever by using the polymerase chain reactionJ Clin Microbiol19902812268926922279999
  37. LA VeretaElisova TD: [The use of the indirect fluorescent antibody technic for determining viral antigens in the urine of patients with hemorrhagic fever with renal syndrome]Lab Delo199057274
  38. LA VeretaTD ElisovaGM VoronkovaThe detection of antibodies to the hantaan virus in the urine of patients with hemorrhagic fever with renal syndromeVopr Virusol199338118217915448
  39. T Avsic-ZupancM PoljakP FurlanR KapsSY XiaoJW LeducIsolation of a strain of a Hantaan virus from a fatal case of hemorrhagic fever with renal syndrome in SloveniaAm J Trop Med Hyg199451439340010.4269/ajtmh.1994.51.3937943563
  40. ZQ YangSY YuJ NieQ ChenZF LiYX LiuJL ZhangJJ XuXM YuXP BuPrevalence of hemorrhagic fever with renal syndrome virus in domestic pigs: an epidemiological investigation in Shandong provinceDi Yi Jun Yi Da Xue Xue Bao200424111283128615567780
  41. L PetterssonJ KlingstromJ HardestamA LundkvistC AhlmM EvanderHantavirus RNA in saliva from patients with hemorrhagic fever with renal syndromeEmerg Infect Dis200814340641110.3201/eid1403.07124218325254
  42. L PetterssonJ RasmusonC AnderssonC AhlmM EvanderHantavirus-specific IgA in saliva and viral antigen in the parotid gland in patients with hemorrhagic fever with renal syndromeJ Med Virol201183586487010.1002/jmv.2204021360546
  43. LJ JamesonA NewtonL CooleEN NewmanMW CarrollNJ BeechingR HewsonRM ChristleyReply to comment--Clement et al.: (Prevalence of antibodies against hantaviruses in serum and saliva of adults living or working on farms in Yorkshire, United Kingdom)Viruses2014693425342710.3390/v609342525256390
  44. L VoutilainenT SironenE TonteriAT BackM RazzautiM KarlssonM WahlstromJ NiemimaaH HenttonenA LundkvistLife-long shedding of Puumala hantavirus in wild bank voles (Myodes glareolus)J Gen Virol201596Pt 61238124710.1099/vir.0.00007625701819
  45. AD VitulloVL HodaraMS MeraniEffect of persistent infection with Junin virus on growth and reproduction of its natural reservoir, Calomys musculinusAm J Trop Med Hyg198737366366910.4269/ajtmh.1987.37.6632825553
  46. AD VitulloMS MeraniVertical transmission of Junin virus in experimentally infected adult Calomys musculinusIntervirology199031633934410.1159/0001501702177742
  47. JN MillsBA EllisJE ChildsKT McKee JrJI MaizteguiCJ PetersTG KsiazekPB JahrlingPrevalence of infection with Junin virus in rodent populations in the epidemic area of Argentine hemorrhagic feverAm J Trop Med Hyg199451555456210.4269/ajtmh.1994.51.5547985747
  48. VA PokhodiaevNI GoncharVA PshenichnovAn experimental study of the contact transmission of the Marburg virusVopr Virusol19913665065081785188
  49. TS ChupurnovaVA PisankoLF BakulinaVA ZhukovAA ChepurnovAssay for level of Marburg virus in blood and isolates from experimentally infected animalsVopr Virusol2000452182010765545
  50. BR AmmanSA CarrollZD ReedTK SealyS BalinandiR SwanepoelA KempBR EricksonJA ComerS CampbellSeasonal pulses of Marburg virus circulation in juvenile Rousettus aegyptiacus bats coincide with periods of increased risk of human infectionPLoS Pathog2012810e100287710.1371/journal.ppat.100287723055920
  51. JT PaweskaP Jansen van VurenKA FentonK GravesAA GrobbelaarN MoollaP LemanJ WeyerN StormSD McCullochLack of Marburg Virus Transmission From Experimentally Infected to Susceptible In-Contact Egyptian Fruit BatsJ Infect Dis2015212Suppl 2S109S11810.1093/infdis/jiv13225838270
  52. P FormentyEM LeroyA EpelboinF LibamaM LenziH SudeckP YabaY AllarangarP BoumandoukiVB NkounkouDetection of Ebola virus in oral fluid specimens during outbreaks of Ebola virus hemorrhagic fever in the Republic of CongoClin Infect Dis200642111521152610.1086/50383616652308
  53. DG BauschJS TownerSF DowellF KaducuM LukwiyaA SanchezST NicholTG KsiazekPE RollinAssessment of the risk of Ebola virus transmission from bodily fluids and fomitesJ Infect Dis2007196Suppl 2S142S14710.1086/52054517940942
  54. B KreuelsD WichmannP EmmerichJ Schmidt-ChanasitG de HeerS KlugeA SowT RenneS GuntherAW LohseA case of severe Ebola virus infection complicated by gram-negative septicemiaN Engl J Med2014371252394240110.1056/NEJMoa141167725337633
  55. N PetrosilloE NicastriS LaniniMR CapobianchiA Di CaroM AntoniniV PuroFN LauriaN ShindoN MagriniEbola virus disease complicated with viral interstitial pneumonia: a case reportBMC Infect Dis20151543210.1186/s12879-015-1169-426471197
  56. P VetterWA Fischer 2ndM SchiblerM JacobsDG BauschL KaiserEbola Virus Shedding and Transmission: Review of Current EvidenceJ Infect Dis2016214suppl 3S177S18410.1093/infdis/jiw25427443613
  57. AA ChughtaiM BarnesCR MacintyrePersistence of Ebola virus in various body fluids during convalescence: evidence and implications for disease transmission and controlEpidemiol Infect201614481652166010.1017/S095026881600005426808232
  58. E GreenL HuntJC RossNM NissenT CurranA BadhanKA SutherlandJ RichardsJS LeeSH AllenViraemia and Ebola virus secretion in survivors of Ebola virus disease in Sierra Leone: a cross-sectional cohort studyLancet Infect Dis20161691052105610.1016/S1473-3099(16)30060-327197552
  59. TR SouthernLD RacsaCG AlbarinoPD FeySH HinrichsCN MurphyVL HerreraAR SambolCE HillEL RyanComparison of FilmArray and Quantitative Real-Time Reverse Transcriptase PCR for Detection of Zaire Ebolavirus from Contrived and Clinical SpecimensJ Clin Microbiol20155392956296010.1128/JCM.01317-1526157148
  60. F JanvierD DelauneT PoyotE ValadeA MerensPE RollinV FoissaudEbola Virus RNA Stability in Human Blood and Urine in West Africa's Environmental ConditionsEmerg Infect Dis201622229229410.3201/eid2202.15139526812135
  61. H BodurE AkinciP OnguruA CarhanY UyarA TanriciO CatalolukA KubarDetection of Crimean-Congo hemorrhagic fever virus genome in saliva and urineInt J Infect Dis2010143e247e24910.1016/j.ijid.2009.04.01819656706
  62. S ThomasG ThomsonS DowallC BruceN CookL EasterbrookL O'DonoghueS SummersL AjazajR HewsonReview of Crimean Congo hemorrhagic fever infection in Kosova in 2008 and 2009: prolonged viremias and virus detected in urine by PCRVector Borne Zoonotic Dis201212980080410.1089/vbz.2011.077622925025
  63. G PalaciosPL QuanOJ JabadoS ConlanDL HirschbergY LiuJ ZhaiN RenwickJ HuiH HegyiPanmicrobial oligonucleotide array for diagnosis of infectious diseasesEmerg Infect Dis2007131738110.3201/eid1301.06083717370518
  64. V BoydI SmithG CrameriAL BurroughsPA DurrJ WhiteC CowledGA MarshLF WangDevelopment of multiplexed bead arrays for the simultaneous detection of nucleic acid from multiple viruses in bat samplesJ Virol Methods201522351210.1016/j.jviromet.2015.07.00426190638
  65. X ZhangY LiuL ZhaoB LiH YuH WenXJ YuAn emerging hemorrhagic fever in China caused by a novel bunyavirus SFTSVSci China Life Sci201356869770010.1007/s11427-013-4518-923917841
  66. X DingX WuT DuanM SiirinH GuzmanZ YangRB TeshSY XiaoNucleotide and amino acid changes in West Nile virus strains exhibiting renal tropism in hamstersAm J Trop Med Hyg200573480380710.4269/ajtmh.2005.73.80316222029
  67. RB TeshM SiirinH GuzmanAP Travassos da RosaX WuT DuanH LeiMR NunesSY XiaoPersistent West Nile virus infection in the golden hamster: studies on its mechanism and possible implications for other flavivirus infectionsJ Infect Dis2005192228729510.1086/43115315962223
  68. JH TonrySY XiaoM SiirinH ChenAP da RosaRB TeshPersistent shedding of West Nile virus in urine of experimentally infected hamstersAm J Trop Med Hyg200572332032410.4269/ajtmh.2005.72.32015772329
  69. KE SteeleMJ LinnRJ SchoeppN KomarTW GeisbertRM ManducaPP CalleBL RaphaelTL ClippingerT LarsenPathology of fatal West Nile virus infections in native and exotic birds during the 1999 outbreak in New York City, New YorkVet Pathol200037320822410.1354/vp.37-3-20810810985
  70. KB PlattBJ TuckerPG HalburBJ BlitvichFG FabiosaK MullinGR ParikhP KitikoonLC BartholomayWA RowleyFox squirrels (Sciurus niger) develop West Nile virus viremias sufficient for infecting select mosquito speciesVector Borne Zoonotic Dis20088222523310.1089/vbz.2007.018218240969
  71. KB PlattBJ TuckerPG HalburS TiawsirisupBJ BlitvichFG FabiosaLC BartholomayWA RowleyWest Nile virus viremia in eastern chipmunks (Tamias striatus) sufficient for infecting different mosquitoesEmerg Infect Dis200713683183710.3201/eid1306.06100817553220
  72. V SaxenaG XieB LiT FarrisT WelteB GongP BoorP WuSJ TangR TeshA hamster-derived West Nile virus isolate induces persistent renal infection in micePLoS Negl Trop Dis201376e227510.1371/journal.pntd.000227523785537
  73. S TiawsirisupBJ BlitvichBJ TuckerPG HalburLC BartholomayWA RowleyKB PlattSusceptibility of fox squirrels (Sciurus niger) to West Nile virus by oral exposureVector Borne Zoonotic Dis201010220720910.1089/vbz.2008.015819402765
  74. X WuL LuH GuzmanRB TeshSY XiaoPersistent infection and associated nucleotide changes of West Nile virus serially passaged in hamstersJ Gen Virol200889Pt 123073307910.1099/vir.0.2008/003210-019008395
  75. JH TonryCB BrownCB CroppJK CoSN BennettVR NerurkarT KuberskiDJ GublerWest Nile virus detection in urineEmerg Infect Dis20051181294129610.3201/eid1108.05023816102323
  76. K MurrayC WalkerE HerringtonJA LewisJ McCormickDW BeasleyRB TeshS Fisher-HochPersistent infection with West Nile virus years after initial infectionJ Infect Dis201020112410.1086/64873119961306
  77. L BarzonM PacentiE FranchinS PagniT MartelloM CattaiR CusinatoG PaluExcretion of West Nile virus in urine during acute infectionJ Infect Dis201320871086109210.1093/infdis/jit29023821721
  78. SA BatyKB GibneyJE StaplesAB PattersonC LevyJ LehmanT WadleighJ FeldR LanciottiCT NugentEvaluation for West Nile Virus (WNV) RNA in urine of patients within 5 months of WNV infectionJ Infect Dis201220591476147710.1093/infdis/jis22122438324
  79. KB GibneyRS LanciottiJJ SejvarCT NugentJM LinnenMJ DeloreyJA LehmanEN BoswellJE StaplesM FischerWest nile virus RNA not detected in urine of 40 people tested 6 years after acute West Nile virus diseaseJ Infect Dis2011203334434710.1093/infdis/jiq05721208926
  80. F MaguranoME RemoliM BaggieriC FortunaA MarchiC FiorentiniP BucciE BenedettiMG CiufoliniC RizzoCirculation of West Nile virus lineage 1 and 2 during an outbreak in ItalyClin Microbiol Infect20121812E545E54710.1111/1469-0691.1201823020657
  81. L BarzonM PacentiE FranchinL SquarzonA SinigagliaS UlbertR CusinatoG PaluIsolation of West Nile virus from urine samples of patients with acute infectionJ Clin Microbiol20145293411341310.1128/JCM.01328-1424951801
  82. L BarzonM PacentiG PaluWest Nile virus and kidney diseaseExpert Rev Anti Infect Ther201311547948710.1586/eri.13.3423627854
  83. L BarzonM PacentiS UlbertG PaluLatest developments and challenges in the diagnosis of human West Nile virus infectionExpert Rev Anti Infect Ther201513332734210.1586/14787210.2015.100704425641365
  84. K ErgunayA KaragulA AbudalalS HaciogluD UsY ErdemA OzkulProspective investigation of the impact of West Nile Virus infections in renal diseasesJ Med Virol201587101625163210.1002/jmv.2422625965349
  85. A NagyE BanO NagyE FerencziA FarkasK BanyaiS FarkasM TakacsDetection and sequencing of West Nile virus RNA from human urine and serum samples during the 2014 seasonal periodArch Virol201616171797180610.1007/s00705-016-2844-527038827
  86. A PapaT TestaE PapadopoulouDetection of West Nile virus lineage 2 in the urine of acute human infectionsJ Med Virol201486122142214510.1002/jmv.2394924760617
  87. D MussoZika Virus Transmission from French Polynesia to BrazilEmerg Infect Dis20152110188710.3201/eid2110.15112526403318
  88. D MussoC RocheTX NhanE RobinA TeissierVM Cao-LormeauDetection of Zika virus in salivaJ Clin Virol201568535510.1016/j.jcv.2015.04.02126071336
  89. C FourcadeJM MansuyM DutertreM DelpechB MarchouP DelobelJ IzopetG Martin-BlondelViral load kinetics of Zika virus in plasma, urine and saliva in a couple returning from Martinique, French West IndiesJ Clin Virol2016821410.1016/j.jcv.2016.06.01127389909
  90. J ZhangX JinZ ZhuL HuangS LiangY XuR LiaoL ZhouY ZhangA Wilder-SmithEarly detection of Zika virus infection among travellers from areas of ongoing transmission in ChinaJ Travel Med20162351310.1093/jtm/taw047
  91. DM DudleyMT AliotaEL MohrAM WeilerG Lehrer-BreyKL WeisgrauMS MohnsME BreitbachMN RasheedCM NewmanA rhesus macaque model of Asian-lineage Zika virus infectionNat Commun201671220410.1038/ncomms1220427352279
  92. HC JangWB ParkUJ KimJY ChunSJ ChoiPG ChoeSI JungY JeeNJ KimEH ChoiFirst Imported Case of Zika Virus Infection into KoreaJ Korean Med Sci20163171173117710.3346/jkms.2016.31.7.117327366020
  93. C ReuskenS PasC GeurtsvanKesselR MoglingJ van KampenT LangerakM KoopmansA van der EijkE van GorpLongitudinal follow-up of Zika virus RNA in semen of a traveller returning from Barbados to the Netherlands with Zika virus disease, March 2016Euro Surveill201621231410.2807/1560-7917.ES.2016.21.23.30251
  94. E D'OrtenzioS MatheronY YazdanpanahX de LamballerieB HubertG PiorkowskiM MaquartD DescampsF DamondI Leparc-GoffartEvidence of Sexual Transmission of Zika VirusN Engl J Med2016374222195219810.1056/NEJMc160444927074370
  95. A DavidsonS SlavinskiK KomotoJ RakemanD WeissSuspected Female-to-Male Sexual Transmission of Zika Virus - New York City, 2016MMWR Morb Mortal Wkly Rep2016652871671710.15585/mmwr.mm6528e227442327
  96. L BarzonM PacentiA BertoA SinigagliaE FranchinE LavezzoP BrugnaroG PaluIsolation of infectious Zika virus from saliva and prolonged viral RNA shedding in a traveller returning from the Dominican Republic to Italy, January 2016Euro Surveill201621101510.2807/1560-7917.ES.2016.21.10.30159
  97. AM BinghamM ConeV MockL Heberlein-LarsonD StanekC BlackmoreA LikosComparison of Test Results for Zika Virus RNA in Urine, Serum, and Saliva Specimens from Persons with Travel-Associated Zika Virus Disease - Florida, 2016MMWR Morb Mortal Wkly Rep2016651847547810.15585/mmwr.mm6518e227171533
  98. MC BonaldoIP RibeiroNS LimaAA Dos SantosLS MenezesSO da CruzIS de MelloND FurtadoEE de MouraL DamascenoIsolation of Infective Zika Virus from Urine and Saliva of Patients in BrazilPLoS Negl Trop Dis2016106e000481610.1371/journal.pntd.000481627341420
  99. F GrischottM PuhanC HatzP SchlagenhaufNon-vector-borne transmission of Zika virus: A systematic reviewTravel Med Infect Dis201614431333010.1016/j.tmaid.2016.07.00227425793
  100. JC LeaoLA GueirosG LodiNA RobinsonC ScullyZika virus: oral healthcare implicationsOral Dis2017231121710.1111/odi.1251227232461
  101. CK OlsonM IwamotoKM PerkinsKN PolenJ HagemanD Meaney-DelmanII IgbinosaS KhanMA HoneinM BellPreventing Transmission of Zika Virus in Labor and Delivery Settings Through Implementation of Standard Precautions - United States, 2016MMWR Morb Mortal Wkly Rep2016651129029210.15585/mmwr.mm6511e327010422
  102. C ScullyA RobinsonCheck before you travel: Zika virus--another emerging global health threatBr Dent J2016220526526710.1038/sj.bdj.2016.18226964604
  103. WL SiqueiraEB MoffaMC MussiMA MachadoZika virus infection spread through saliva--a truth or myth?Braz Oral Res2016301210.1590/1807-3107BOR-2016.vol30.0046
  104. C DomingoS YactayoE AgbenuM DemanouAR SchulzK DaskalowM NiedrigDetection of yellow fever 17D genome in urineJ Clin Microbiol201149276076210.1128/JCM.01775-1021106799
  105. MJ MartinezA VilellaT PumarolaM RoldanVG SequeraI VeraEB HayesPersistence of yellow fever vaccine RNA in urineVaccine201129183374337610.1016/j.vaccine.2011.02.07521385635
  106. JS PeirisCM ChuVC ChengKS ChanIF HungLL PoonKI LawBS TangTY HonCS ChanClinical progression and viral load in a community outbreak of coronavirus-associated SARS pneumonia: a prospective studyLancet200336193711767177210.1016/S0140-6736(03)13412-512781535
  107. WK WangSY ChenIJ LiuYC ChenHL ChenCF YangPJ ChenSH YehCL KaoLM HuangDetection of SARS-associated coronavirus in throat wash and saliva in early diagnosisEmerg Infect Dis20041071213121910.3201/eid1007.03111315324540
  108. C DrostenLL ChiuM PanningHN LeongW PreiserJS TamS GuntherS KrammeP EmmerichWL NgEvaluation of advanced reverse transcription-PCR assays and an alternative PCR target region for detection of severe acute respiratory syndrome-associated coronavirusJ Clin Microbiol20044252043204710.1128/JCM.42.5.2043-2047.200415131168
  109. VC ChengIF HungBS TangCM ChuMM WongKH ChanViral replication in the nasopharynx is associated with diarrhea in patients with severe acute respiratory syndromeClin Infect Dis.200438446747510.1086/38268114765337
  110. PK ChanTo WKKC NgRK LamTK NgRC ChanA WuWC YuN LeeDS HuiLaboratory diagnosis of SARSEmerg Infect Dis200410582583110.3201/eid1005.03068215200815
  111. CM ChuLL PoonVC ChengKS ChanIF HungMM WongKH ChanWS LeungBS TangVL ChanInitial viral load and the outcomes of SARSCMAJ2004171111349135210.1503/cmaj.104039815557587
  112. IF HungVC ChengAK WuBS TangKH ChanCM ChuViral loads in clinical specimens and SARS manifestationsEmerg Infect Dis.20041091550155710.3201/eid1009.04005815498155
  113. CM ChuVC ChengIF HungKS ChanBS TangTH TsangKH ChanKY YuenViral load distribution in SARS outbreakEmerg Infect Dis200511121882188610.3201/eid1112.04094916485474
  114. JB MahonyA PetrichL LouieX SongS ChongM SmiejaM CherneskyM LoebS RichardsonOntario Laboratory Working Group for the Rapid Diagnosis of Emerging I: Performance and Cost evaluation of one commercial and six in-house conventional and real-time reverse transcription-pcr assays for detection of severe acute respiratory syndrome coronavirusJ Clin Microbiol20044241471147610.1128/JCM.42.4.1471-1476.200415070991
  115. SK LauXY ChePC WooBH WongVC ChengGK WooIF HungRW PoonKH ChanJS PeirisSARS coronavirus detection methodsEmerg Infect Dis20051171108111110.3201/eid1107.04104516022791
  116. SC WongJK ChanKC LeeES LoDN TsangDevelopment of a quantitative assay for SARS coronavirus and correlation of GAPDH mRNA with SARS coronavirus in clinical specimensJ Clin Pathol200558327628010.1136/jcp.2004.01659215735160
  117. D XuZ ZhangL JinF ChuY MaoH WangM LiuM WangL ZhangGF GaoPersistent shedding of viable SARS-CoV in urine and stool of SARS patients during the convalescent phaseEur J Clin Microbiol Infect Dis200524316517110.1007/s10096-005-1299-515789222
  118. XW WangJS LiTK GuoB ZhenQX KongB YiZ LiN SongM JinXM WuExcretion and detection of SARS coronavirus and its nucleic acid from digestive systemWorld J Gastroenterol200511284390439510.3748/wjg.v11.i28.439016038039
  119. A GoffardM LazrekC SchanenPE LobertL BocketA DewildeD HoberEmergent viruses: SARS-associate coronavirus and H5N1 influenza virusAnn Biol Clin (Paris)200664319520816698555
  120. D NormileSecond Lab Accident Fuels Fears About SARSScience200430356542610.1126/science.303.5654.2614704402
  121. EF DonaldsonAN HaskewJE GatesJ HuynhCJ MooreMB FriemanMetagenomic analysis of the viromes of three North American bat species: viral diversity among different bat species that share a common habitatJ Virol20108424130041301810.1128/JVI.01255-1020926577
  122. L LiuQ WeiX AlvarezH WangY DuH ZhuH JiangJ ZhouP LamL ZhangEpithelial cells lining salivary gland ducts are early target cells of severe acute respiratory syndrome coronavirus infection in the upper respiratory tracts of rhesus macaquesJ Virol20118584025403010.1128/JVI.02292-1021289121
  123. B LuY HuangL HuangB LiZ ZhengZ ChenJ ChenQ HuH WangEffect of mucosal and systemic immunization with virus-like particles of severe acute respiratory syndrome coronavirus in miceImmunology2010130225426110.1111/j.1365-2567.2010.03231.x20406307
  124. C DrostenM SeilmaierVM CormanW HartmannG ScheibleS SackW GuggemosR KalliesD MuthS JunglenClinical features and virological analysis of a case of Middle East respiratory syndrome coronavirus infectionLancet Infect Dis201313974575110.1016/S1473-3099(13)70154-323782859
  125. I EckerleMA MullerS KalliesDN GotthardtC DrostenIn-vitro renal epithelial cell infection reveals a viral kidney tropism as a potential mechanism for acute renal failure during Middle East Respiratory Syndrome (MERS) Coronavirus infectionVirol J20131035910.1186/1743-422X-10-35924364985
  126. GK GohAK DunkerV UverskyPrediction of Intrinsic Disorder in MERS-CoV/HCoV-EMC Supports a High Oral-Fecal TransmissionPLoS Curr201351223
  127. AS OmraniJA Al-TawfiqZA MemishMiddle East respiratory syndrome coronavirus (MERS-CoV): animal to human interactionPathog Glob Health2015109835436210.1080/20477724.2015.112285226924345
  128. VM CormanS OlschlagerCM WendtnerJF DrexlerM HessC DrostenPerformance and clinical validation of the RealStar MERS-CoV Kit for detection of Middle East respiratory syndrome coronavirus RNAJ Clin Virol201460216817110.1016/j.jcv.2014.03.01224726679
  129. RH ChaJS JohI JeongJY LeeHS ShinG KimY KimCritical Care Team of National Medical C: Renal Complications and Their Prognosis in Korean Patients with Middle East Respiratory Syndrome-Coronavirus from the Central MERS-CoV Designated HospitalJ Korean Med Sci201530121807181410.3346/jkms.2015.30.12.180726713056
  130. JA Al-TawfiqZA MemishManaging MERS-CoV in the healthcare settingHosp Pract (1995)201543315816310.1080/21548331.2015.107402926224424
  131. VM CormanAM AlbarrakAS OmraniMM AlbarrakME FarahM AlmasriD MuthA SiebergB MeyerAM AssiriViral Shedding and Antibody Response in 37 Patients With Middle East Respiratory Syndrome Coronavirus InfectionClin Infect Dis201662447748326565003
  132. C GossnerN DanielsonA GervelmeyerF BertheB FayeK Kaasik AaslavC AdlhochH ZellerP PenttinenD CoulombierHuman-Dromedary Camel Interactions and the Risk of Acquiring Zoonotic Middle East Respiratory Syndrome Coronavirus InfectionZoonoses Public Health20166311910.1111/zph.1217125545147
  133. Y DuRA HughesS BhadraYS JiangAD EllingtonB LiA Sweet Spot for Molecular Diagnostics: Coupling Isothermal Amplification and Strand Exchange Circuits to GlucometersSci Rep201551103910.1038/srep1103926050646
  134. A Abd El WahedP PatelD HeidenreichFT HufertM WeidmannReverse transcription recombinase polymerase amplification assay for the detection of middle East respiratory syndrome coronavirusPLoS Curr2013516
  135. Yellow Fever laboratory diagnostic testing algorithm for Africa - Interim guidance- July 2016.
  136. CF FulhorstTG KsiazekCJ PetersRB TeshExperimental infection of the cane mouse Zygodontomys brevicauda (family Muridae) with guanarito virus (Arenaviridae), the etiologic agent of Venezuelan hemorrhagic feverJ Infect Dis.1999180496696910.1086/31502910479119
  137. V CormanR KalliesH PhilippsG GöpnerMA MüllerI EckerleCharacterization of a Novel Betacoronavirus Related to Middle East Respiratory Syndrome Coronavirus in European HedgehogsJ of Virol201488171772410.1128/JVI.01600-1324131722
  138. J PoissyA.Goffard, E.Parmentier-Decrucq, R.Favory, M.Kauv, E.Kipnis, et al: Kinetics andpatternofviralexcretioninbiologicalspecimensoftwo MERS-CoV casesJCV20146127527825073585
  139. Z HerB MalleretM ChanEK OngSC WongDJ KwekActive infection of human blood monocytes by Chikungunya virus triggers an innate immune responseJ Immunol.2010184105903591310.4049/jimmunol.090418120404274
  140. V ReddyRS ManiA DesaiV RaviCorrelation of plasma viral loads and presence of Chikungunya IgM antibodies with cytokine/chemokine levels during acute Chikungunya virus infectionJ Med Virol.20148681393140110.1002/jmv.2387524523146
  141. MM ParidaSR SanthoshPK DashNK TripathiV LakshmiN MamidiRapid and real-time detection of Chikungunya virus by reverse transcription loop-mediated isothermal amplification assayJ Clin Microbiol.200745235135710.1128/JCM.01734-0617135444
  142. P LewthwaiteR VasanthapuramJC OsborneA BegumJL PlankMV ShankarChikungunya virus and central nervous system infections in children, IndiaEmerg Infect Dis.200915232933110.3201/eid1502.08090219193287
  143. C KlungthongRV GibbonsB ThaisomboonsukA NisalakS KalayanaroojV ThirawuthDengue virus detection using whole blood for reverse transcriptase PCR and virus isolationJ Clin Microbiol.20074582480248510.1128/JCM.00305-0717522268
  144. A SrikiatkhachornS WichitRV GibbonsS GreenDH LibratyTP EndyDengue viral RNA levels in peripheral blood mononuclear cells are associated with disease severity and preexisting dengue immune statusPLoS One.2012712e5133510.1371/journal.pone.005133523284680
  145. T SolomonNM DungDW VaughnR KneenLT ThaoB RaengsakulrachNeurological manifestations of dengue infectionLancet.200035592091053105910.1016/S0140-6736(00)02036-510744091
  146. A BarthelAC GourinatC CazorlaC JoubertM Dupont-RouzeyrolE DesclouxBreast milk as a possible route of vertical transmission of dengue virus?Clin Infect Dis.201357341541710.1093/cid/cit22723575200
  147. M RiosS DanielC ChanceyIK HewlettSL StramerWest Nile virus adheres to human red blood cells in whole bloodClin Infect Dis.200745218118610.1086/51885017578776
  148. MC LanteriTH LeeL WenZ KaidarovaMD BravoNE KielyWest Nile virus nucleic acid persistence in whole blood months after clearance in plasma: implication for transfusion and transplantation safetyTransfusion.201454123232324110.1111/trf.1276424965017
  149. PA TilleyJD FoxGC JayaramanJK PreiksaitisNucleic acid testing for west nile virus RNA in plasma enhances rapid diagnosis of acute infection in symptomatic patientsJ Infect Dis.2006193101361136410.1086/50357716619182
  150. KL TylerJ PapeRJ GoodyM CorkillBK Kleinschmidt-DeMastersCSF findings in 250 patients with serologically confirmed West Nile virus meningitis and encephalitisNeurology.200666336136510.1212/01.wnl.0000195890.70898.1f16382032
  151. AF HinckleyDR O'LearyEB HayesTransmission of West Nile virus through human breast milk seems to be rarePediatrics.20071193e666e67110.1542/peds.2006-210717332186
  152. C DomingoP PatelJ YillahM WeidmannJA MendezER NakouneAdvanced yellow fever virus genome detection in point-of-care facilities and reference laboratoriesJ Clin Microbiol.201250124054406010.1128/JCM.01799-1223052311
  153. C TraiberP Coelho-AmaralVR RitterA WingeInfant meningoencephalitis caused by yellow fever vaccine virus transmitted via breastmilkJ Pediatr (Rio J).201187326927221461453
  154. M ChavesP RiccioL PatruccoJI RojasE CristianoLongitudinal myelitis associated with yellow fever vaccinationJ Neurovirol.200915434835010.1080/1355028090306280519579072
  155. L NoronhaC ZanlucaML AzevedoKG LuzCN SantosZika virus damages the human placental barrier and presents marked fetal neurotropismMem Inst Oswaldo Cruz.2016111528729310.1590/0074-0276016008527143490
  156. B RozeF NajioullahA SignateK ApetseY BrousteS GourgoudouZika virus detection in cerebrospinal fluid from two patients with encephalopathy, Martinique, February 2016Euro Surveill.201621161410.2807/1560-7917.ES.2016.21.16.30205
  157. S SwaminathanR SchlabergJ LewisKE HansonMR CouturierFatal Zika Virus Infection with Secondary Nonsexual TransmissionN Engl J Med.2016375191907190910.1056/NEJMc161061327681699
  158. G CalvetRS AguiarAS MeloSA SampaioI de FilippisA FabriDetection and sequencing of Zika virus from amniotic fluid of fetuses with microcephaly in Brazil: a case studyLancet Infect Dis.201616665366010.1016/S1473-3099(16)00095-526897108
  159. M Dupont-RouzeyrolA BironO O'ConnorE HuguonE DesclouxInfectious Zika viral particles in breastmilkLancet.201638710023105110.1016/S0140-6736(16)00624-326944028
  160. JS TownerPE RollinDG BauschA SanchezSM CraryM VincentRapid diagnosis of Ebola hemorrhagic fever by reverse transcription-PCR in an outbreak setting and assessment of patient viral load as a predictor of outcomeJ Virol.20047884330434110.1128/JVI.78.8.4330-4341.200415047846
  161. MS SowJF EtardS BaizeN MagassoubaO FayeP MsellatiNew Evidence of Long-lasting Persistence of Ebola Virus Genetic Material in Semen of SurvivorsJ Infect Dis.2016214101475147610.1093/infdis/jiw07827142204
  162. FM BaggiA TaybiA KurthM Van HerpA Di CaroR WolfelManagement of pregnant women infected with Ebola virus in a treatment centre in Guinea, June 2014Euro Surveill.201419491410.2807/1560-7917.ES2014.19.49.20983
  163. M JacobsA RodgerDJ BellS BhaganiI CropleyA FilipeLate Ebola virus relapse causing meningoencephalitis: a case reportLancet.20163881004349850310.1016/S0140-6736(16)30386-527209148
  164. AK RoweJ BertolliAS KhanR MukunuJJ Muyembe-TamfumD BresslerClinical, virologic, and immunologic follow-up of convalescent Ebola hemorrhagic fever patients and their household contacts, Kikwit, Democratic Republic of the Congo. Commission de Lutte contre les Epidemies a KikwitJ Infect Dis.1999179Suppl 1S28S3510.1086/5143189988162
  165. LF NgM WongS KohEE OoiKF TangHN LeongDetection of severe acute respiratory syndrome coronavirus in blood of infected patientsJ Clin Microbiol.200442134735010.1128/JCM.42.1.347-350.200414715775
  166. H WangY MaoL JuJ ZhangZ LiuX ZhouDetection and monitoring of SARS coronavirus in the plasma and peripheral blood lymphocytes of patients with severe acute respiratory syndromeClin Chem.20045071237124010.1373/clinchem.2004.03123715229153
  167. YD WangY LiGB XuXY DongXA YangZR FengDetection of antibodies against SARS-CoV in serum from SARS-infected donors with ELISA and Western blotClin Immunol.2004113214515010.1016/j.clim.2004.07.00315451470
  168. EC HungSS ChimPK ChanYK TongEK NgRW ChiuDetection of SARS coronavirus RNA in the cerebrospinal fluid of a patient with severe acute respiratory syndromeClin Chem.200349122108210910.1373/clinchem.2003.02543714633896
  169. ET IsakbaevaN KhetsurianiRS BeardA PeckD ErdmanSS MonroeSARS-associated coronavirus transmission, United StatesEmerg Infect Dis.200410222523110.3201/eid1002.03073415030687
  170. SY KimSJ ParkSY ChoRH ChaHG JeeG KimViral RNA in Blood as Indicator of Severe Outcome in Middle East Respiratory Syndrome Coronavirus InfectionEmerg Infect Dis.201622101813181610.3201/eid2210.16021827479636
  171. MN KimYJ KoMW SeongJS KimBM ShinH SungAnalytical and Clinical Validation of Six Commercial Middle East Respiratory Syndrome Coronavirus RNA Detection Kits Based on Real-Time Reverse-Transcription PCRAnn Lab Med.201636545045610.3343/alm.2016.36.5.45027374710
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