Covid-19: Anal Swabs as the potentially optimal specimen for SARS-CoV-2 detection

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Since December 2019, an outbreak of atypical pneumonia caused by SARS coronavirus 2 (SARS-CoV-2) has led to a serious epidemic in China and other countries.

Phylogenetic analyses of the coronavirus genomes revealed that SARS-CoV-2 belongs to the Betacoronavirus genus, a class of positive-sense, ssRNA viruses that can cause respiratory, intestinal, liver and nervous system infections in animals and humans [1].

SARS-CoV-2 is composed of four structural proteins, known as the S (spike), E (envelope), M (membrane) and N (nucleocapsid) proteins, and possesses 82% identity to SARS-CoV and 50% identity to Middle East respiratory syndrome coronavirus (MERS-CoV) based on genome sequencing [2]. Moreover, it spreads by human-to-human transmission via droplets or direct contact, and infection has been estimated to have a mean incubation period of 6.4 days and a basic reproduction number of 2.24–3.58 [3].

Numerous retrospective studies have indicated that prevalent clinical manifestations of COVID-19 patients are fever, dry cough and dyspnea [4]; less common symptoms present as the production of sputum, headache and some gastrointestinal symptoms; moreover, an increasing number of patients with asymptomatic infection patients have been discovered [5–7].

According to the latest guidelines of the diagnosis and treatment of pneumonitis caused by 2019-nCoV (trial version 7) published by the National Health Commission of the People’s Republic of China [8], the diagnosis of COVID-19 must be confirmed by reverse transcriptase-PCR (RT-PCR) or gene sequencing.

At present, various biological samples of COVID-19 are used in the detection of SARS-CoV-2, and upper respiratory tract nasopharyngeal swabs are the most common sample type. However, growing evidence has revealed positive detection of nucleic acids in anal swabs of patients with COVID-19, although the positive rate is low [9,10].

A previous study showed a positive RT-PCR test on throat swabs of patients recovered from COVID-19 [11], leading to conflicts with current criteria [8].

Here, we reported clinically cured cases with only positive results in anal swabs, which conflicts with current criteria for releasing people from quarantine, and further investigated the clinical value of anal swabs for SARS-CoV-2 detection.

We propose anal swabs as the potentially optimal specimen for SARS-CoV-2 detection for evaluation of hospital discharge of COVID-19 patients.

Discussion
Since December 2019, the outbreak of COVID-2019 caused by SARA-CoV-2 has become a global health concern [15]. SARS-CoV-2 has quickly spread across China and all continents except Antarctica [16–19] and caused more than 3.5 million confirmed cases with 24,000 deaths (https://www.who.int/emergencies/diseases/novel-coronavirus-2019/situation-reports).

Thus far, the origin of the coronavirus remains unclear. The latest report discovered that the pangolin-CoV genome showed 91.02% nucleotide identity with the SARS-CoV genome, which suggested that pangolin species are a natural reservoir of SARS-CoV-2-like CoVs [20].

Molecular detection remains the gold standard for diagnosis. As a recommended method, RT-PCR is widely used to detect SARS-CoV-2. To date, throat, sputum and anal swabs have been considered applicable for RT-PCR detection. Evidence has shown that sample type plays a critical role in SARS-CoV-2 detection.

Viral RNA can be easily detected in nasopharyngeal, sputum and stool specimens [21], and the highest positivity rates were detected in sputum and bronchoalveolar lavage specimens [22]. However, the course of SARS-CoV-2 infection remains unclear.

As the cellular receptor for SARS-CoV-2, angiotensin-converting enzyme 2 (ACE2) is the key for SARS-CoV to enter target cells during the course of viral infection [23–25]. Expression of ACE2 protein in human organs showed that ACE2 is most abundantly expressed on the surface of alveolar epithelial cells and small intestine epithelial cells [26], which are involved in the progression of pneumonia [27]. Intriguingly, a connection may exist between the lungs and GI tract [28], and SARS-CoV-2 may be shed through multiple routes in the different phases of viral infection.

In this study, we found that SARS-CoV-2 detection was positive in anal swabs but negative in other sample types of a few cured patients, which challenges the current standards for discharge and termination of compulsory isolation for COVID-19 patients.

Similar to SARS-CoV and MERS-CoV patients [29,30], intestinal infection was observed in the later stages of infection, indicating that the clearance time of SARS-CoV-2 in the digestive tract was later than that in the respiratory tract. In particular, gastrointestinal symptoms were found in children with COVID-19 [31].

However, the burden of novel coronavirus infections is still underestimated; only approximately 1% of all confirmed SARS-CoV-2 cases involve children according to the current estimates [32], so more biological samples and methods (e.g., serologic detection) for SARS-CoV-2 infection in children must be studied.

Notably, live SARS-CoV-2 virus was isolated from fecal samples in three of 11 adult patients [33]. Therefore, anal swabs might be the optimal specimen for SARS-CoV-2 detection to evaluate hospital discharge of COVID-19 patients. Patients with positive stool results require further isolation until the virus is completely eliminated.

Based on the knowledge about this specific viral infection and considering the prolonged viral RNA detection in anal swabs [34] and detectable viral RNA in the blood cohort progressing to a severe symptom stage [35], we proposed the potential infection course of SARS-CoV-2 as follows (Figure 3): upper respiratory infection (mainly by respiratory droplets); lower respiratory infection (mainly presented as pulmonary infection); viremia formation; and transmission to other organs (including the GI tract) and colonization via ACE2. Therefore, different sample types should be chosen for SARS-CoV-2 detection in various infection phases. Fortunately, Sethuraman et al. devised a clinically useful timeline of diagnostic markers for the detection of COVID-19 [36].

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Figure 3.
Potential infection course of SARS-CoV-2 and the different specimens for SARS-CoV-2 detection.
First, SARS-CoV-2 infects the upper respiratory system mainly by respiratory droplets (asymptomatic or fever, dry cough, fatigue, myalgia and dyspnoea; high positive RT-PCR results in throat swabs). Subsequently, it infects the lower respiratory tract and massively replicates (mainly presented as pulmonary infection; high positive results of RT-PCR in throat swabs and sputum). Furthermore, virus is released into blood, leading to the formation of viremia (low copy number detected in blood by RT-PCR). Finally, it is transmitted to other organs, including the GI tract, and colonizes via ACE2 (higher positive detection rate in anal swabs).
ACE2: Angiotensin-converting enzyme 2; GI : Gastrointestinal; RT-PCR: Reverse transcriptase-PCR.

In summary, we found that SARS-CoV-2 detection was positive in anal swabs but negative in other sample types of several cured patients. Our findings greatly contribute to a comprehensive understanding of COVID-19. Although the study was limited to a small number of patients, and further longitudinal studies on a larger cohort would help to understand the prognosis of the disease.


Coronavirus disease 2019 (COVID-19) is caused by severe acute respiratory syndrome coronavirus-2 (SARS-CoV-2), can lead to severe or critical disease, and is a global pandemic [1,2]. The clinical manifestations of COVID-19 range from asymptomatic infection, to mild, severe or critical respiratory tract infections, gastrointestinal and neurological symptoms, and death [3–7]. SARS-CoV-2 infection is estimated to be responsible for approximately 20% of severe cases and approximately 5% of fatal cases among infected individuals [2].

For this deadly infection, much attention should be paid to decreasing mortality in severe cases with effective antiviral therapies. It is therefore of great significance to precisely determine the kinetics of virus shedding and the sites of viral replication.

The presence of viral RNA has been reported in a broad range of sample types, including but not limited to respiratory, stool, urine, and blood samples [3,7]. Viral RNA detection not only provides major evidence for clinical diagnosis but also reflects the virus replication sites, and the viral RNA load is a useful parameter to identify the status of viral replication and clearance [7–9].

Hence, viral load quantification in patients has been used to monitor disease progression. The correlations between viral RNA load and clinical symptoms and laboratory test results have provided clues to predict disease severity. For example, the viral RNA levels in nasopharyngeal aspirates and blood were correlated with death in SARS-CoV-infected patients [10].

In COVID-19 patients, the viral loads in sputum and blood were found to be related to prognosis [7,11]. However, most recent studies involved few severe cases and employed a single sample type. Whether the viral load in samples collected from different anatomical sites will predict clinical outcome in severe patients still needs to be thoroughly investigated.

In a previous report, two COVID-19 cohorts suffering from severe infections were recruited for a clinical trial (LOTUS) to determine the antiviral efficacy of lopinavir-ritonavir [8]. In this study, we longitudinally quantified the viral load in consecutive throat swab (TS) and anal swab (AS) samples collected from the LOTUS cohorts to evaluate the viral loads in specimens collected from different anatomical sites and their association with clinical outcomes in severe COVID-19 patients. Our findings suggest that viral replication in extrapulmonary sites and viral RNA load are highly correlated with adverse outcome of COVID-19 patients.

Discussion
In this study, we analysed the viral RNA positive rate and viral loads in consecutively collected paired TS and AS samples from hospitalized severe COVID-19 patients. We found that viral RNA could be detected in TS and AS samples, but the rates of positivity were different (TS 33.4%, AS 20.9%).

The mean viral loads were also different between groups (TS 1.0×106 copies/ml, AS 2.3×105 copies/ml). The time from symptom onset to positive viral RNA detection in AS samples was significantly lower in non-survivors than in survivors (median number of days of 14 vs 19). The virus positive rate and the viral load in AS in week 2 after symptom onset were significantly higher in non-survivors than in survivors.

Several groups have reported the detection rate of SARS-CoV-2 in different samples from COVID-19 patients with different disease severities. However, the data vary greatly between studies [11,12,15]. Other studies reported that the average viral RNA positive rates in TS and faecal samples from COVID-19 patients were 18.2%–62.5% and 17.0%–26.7%, respectively [12,16].

These disparate values may be attributed to the inconsistent disease severity, sampling time, sample number and type used to evaluate the viral positive detection rates across studies. The TS viral load between weeks showed no significant difference PSO in our study, which was consistent with Zheng et al.’s study [7]. But Wölfel et al.’s considered there were significant differences of TS viral load between weeks PSO [14]. The difference between our findings and those of previous reports may related to the enrolled cases with different disease severities.

The viral RNA detections in enteric samples were similar to that in SARS patients, in which the virus was isolated from stool samples, and a high viral RNA prevalence was found in the stool samples [17,18]. Human organoid culture experiments have shown that replication of SARS-CoV-2 in the gut is higher than that in the lungs [19,20].

The expression of N protein was visualized in the cytoplasm of gastric, duodenal, and rectal glandular epithelial cells, which further confirmed the regional replication of SARS-CoV-2 [21]. The presence of viral RNA in different anatomical sites indicates the location of replication and/or the transmission route. It is well known that the respiratory tract is the initial replication site of SARS-CoV-2.

The detection of viral RNA in anal samples might be the result from transmission of virus from the respiratory tract to the intestinal tract by swallowing, the replication of virus within extrapulmonary organs, or the increased intestinal permeability during disease progression. However, we found no correlation of the viral load in AS with intestinal infection symptoms in our study, though diarrohea and vomiting were reported in the COVID-19 patients [15].

To obtain proof of active virus replication in the absence of histopathology, we also analysed viral subgenomic mRNAs in clinical samples [14]. It showed that subgenomic mRNAs were detectable in both viral RNA positive TS (35.6%) and AS (13.9%) samples (Supplementary Figure 1).

Our viral shedding data also indicate the important role of the gut during disease progression. Collectively, these findings emphasize that enteric viral replication and transmission in individuals are important predictors of disease severity. Enteric samples should be routinely collected for virally testing for COVID-19 diagnosis, as they are for SARS diagnosis.

The viral load reflects the dynamic interplay between viral replication and virus clearance by host immune activities [18]. The examination of viral load in SARS and Middle East Respiratory Syndrome (MERS) patients has been used to predict disease progression [10,22].

In SARS, a high viral load in respiratory, stool and blood samples was related to death [10]. In MERS, the viral loads in the severe group were higher than those in the mild group, while the viral shedding time and intensity were closely related to SARS [22,23]. The viral load in the respiratory tract was reported positively linked to lung disease severity in COVID-19 patients, indicating that it is a predictor of disease severity [24].

In our data, the TS viral load was higher than the AS viral load. However, we found no correlations of TS viral load with death in our study. Such finding was consistent with Fajnzylber et al’s report, in which they recruited severe COVID-19 patients [25]. The disparities might be related to the disease severity of recruited patients, sampling time and sample number.

Only the AS viral load was significantly higher in non-survivors than in survivors, and the difference became significant at week 2 PSO, which may indicate that the second week during disease progression is a critical point for determining COVID-19 outcome. The presence or absence of an extrapulmonary infection at week 2 indicates whether a patient’s immune system has been effective in preventing the spread of the virus, thereby determining the patient’s chance of survival.

The viral RNA detections and viral load in consecutively collected paired samples from patients showed that TS was positive in high viral RNA concentrations, followed by AS from survivors and non-survivors. The respiratory tract being the primary replication site of SARS-CoV-2 was supported by the high TS viral load. Viral RNA could be detected earlier in TS than AS, and TS had higher viral loads than AS in both survivors and non-survivors.

The expression of angiotensin-converting enzyme 2 (ACE2), the receptor for SARS-CoV-2, is much higher in the small intestine than in the lungs [26–28]. It is hypothesized that in some patients, the virus travels to the intestine after the initial respiratory system infection and actively replicates [29]; viral RNA “spillover” into the blood would thus predict adverse outcomes. Monitoring enteric and blood samples would be a specific way to monitor disease progression.

There are some limitations of our study. One is that there was a lack of samples from the first 5 days, and as such, we could not provide a detailed characterization of viral load kinetics in the early stage. The second is that although the AS and TS were collected at the same time every 4–7 days until the patients were discharged or died, we did not continue to monitor the patients after discharge. However, it would be not affected the conclusions in this study.

The third is that we analysed the dynamics of viral RNA positivity and viral load with samples taken from patients who received antivirals, antibiotics, corticosteroids and other treatments, which could have affected the patterns. However, our findings can help identify those patients with severe COVID-19 who are likely to experience disease progression.

In conclusion, based on the analysis of a relative large amount of samples collected from severe COVID-19 patients, we found that a high viral RNA positivity rate in AS, a high viral load in AS, and early positive detection in AS can predispose COVID-19 patients to adverse outcomes. Early administration of effective antiviral drugs is critical for treating COVID-19. The presence of viral replication in extrapulmonary sites predisposes to adverse outcomes and should thus be monitored carefully.

reference link: https://www.tandfonline.com/doi/full/10.1080/22221751.2020.1858700


Reference

  1. Papers of special note have been highlighted as: • of interest; •• of considerable interest1. Weiss SR, Leibowitz JL. Coronavirus pathogenesis. Adv. Virus Res. 81, 85–164 (2011). [PMC free article] [PubMed] [Google Scholar]
  2. Lu R, Zhao X, Li J. et al. Genomic characterization and epidemiology of 2019 novel coronavirus: implications for virus origins and receptor binding. Lancet 395(10224), 565–574 (2020). [PMC free article] [PubMed] [Google Scholar]
  3. Lai CC, Shih TP, Ko WC, Tang HJ, Hsueh PR. et al. Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) and coronavirus disease-2019 (COVID-19): the epidemic and the challenges. Int. J. Antimicrob. Agents. 55(3), 105924 (2020). [PMC free article] [PubMed] [Google Scholar]
  4.  Rodriguea-Morales AJ, Cardona-Ospina JA, Gutiérrez-Ocampo E. et al. Clinical, laboratory and imaging features of COVID-19: a systematic review and meta-analysis. Travel Med. Infect. Dis. 34, 101623 (2020). [PMC free article] [PubMed] [Google Scholar]
  5. Wong J, Abdul Aziz ABZ, Chaw L. et al. High proportion of asymptomatic and presymptomatic COVID-19 infections in travelers and returning residents to Brunei. J. Travel Med. (2020) (Epub ahead of print). [PMC free article] [PubMed] [Google Scholar]
  6. Wang L, Xu X, Ruan J. et al. Quadruple therapy for asymptomatic COVID-19 infection patients. Expert Rev. Anti Infect. Ther. (2020) (Epub ahead of print). [PMC free article] [PubMed] [Google Scholar]
  7. Corman VM, Rabenau HF, Adams O. et al. SARS-CoV-2 asymptomatic and symptomatic patients and risk for transfusion transmission. Transfusion (2020) (Epub ahead of print). [PMC free article] [PubMed] [Google Scholar]
  8. National Health Commission of the People’s Republic of China. New coronavirus pneumonia prevention and control program (7nd ed.) (in Chinese). 4 March 2020. National Health Commission of the People’s Republic of China. http://www.nhc.gov.cn/xcs/zhengcwj/202003/46c9294a7dfe4cef80dc7f5912eb1989/files/ce3e6945832a438eaae415350a8ce964.pdf
  9. Wu J, Liu J, Li S. et al. Detection and analysis of nucleic acid in various biological samples of COVID-19 patients. Travel Med. Infect. Dis. (2020) (Epub ahead of print). [PMC free article] [PubMed] [Google Scholar]
  10. Zhang W, Du RH, Li B. et al. Molecular and serological investigation of 2019-nCoV infected patients: implication of multiple shedding routes. Emerg. Microbes Infect. 9(1), 386–389 (2020). [PMC free article] [PubMed] [Google Scholar]
  11. Lan L, Xu D, Ye G. et al. Positive RT-PCR Test Results in Patients Recovered From COVID-19. JAMA 323(15), 1502–1503 (2020). [PMC free article] [PubMed] [Google Scholar]•• Describes reverse transcriptase–polymerase chain reaction (RT-PCR) test results in four health professionals discharged from hospitalization or quarantine after two negative RT-PCR test results and resolution of clinical COVID-19 infection, which proposed that more attention should be paid to the follow-up of recovered patients.
  12.  Guiot J, Demarche S, Henket M. et al. Methodology for Sputum Induction and Laboratory Processing. J. Vis. Exp. (130), 56612 (2017). [PMC free article] [PubMed] [Google Scholar]
  13. Lippi G, Simundic AM, Plebani M. Potential preanalytical and analytical vulnerabilities in the laboratory diagnosis of coronavirus disease 2019 (COVID-19). Clin. Chem. Lab Med. (2020) (Epub ahead of print). [PubMed] [Google Scholar]
  14. Buonsenso D, Parri N, De Rose C, Valentini P. Gemelli-pediatric COVID-19 team. Toward a clinically based classification of disease severity for paediatric COVID-19. Lancet Infect. Dis. (2020) (Epub ahead of print). [PMC free article] [PubMed] [Google Scholar]
  15. Zhu N, Zhang D, Wang W. et al. A novel coronavirus from patients with pneumonia in China, 2019. N. Engl. J. Med. 382(8), 727–733 (2020). [PMC free article] [PubMed] [Google Scholar]
  16. Deng Z, Yuxing HU, Yang P. et al. Diagnosis and treatment of an acute severe pneumonia patient with COVID-19: case report. J. Med. Virol. (2020) (Epub ahead of print). [PubMed] [Google Scholar]
  17. Li Q, Guan X, Wu P. et al. Early transmission dynamics in Wuhan, China, of Novel Coronavirus-Infected Pneumonia. N. Engl. J. Med. 382(13), 1199–1207 (2020). [PMC free article] [PubMed] [Google Scholar]
  18. Huang C, Wang Y, Li X. et al. Clinical features of patients infected with 2019 novel coronavirus in Wuhan, China. Lancet 395(10223), 497–506 (2020). [PMC free article] [PubMed] [Google Scholar]
  19. Holshue ML, DeBolt C, Lindquist S. et al. First case of 2019 novel coronavirus in the United States. N. Engl. J. Med. 382(10), 929–936 (2020). [PMC free article] [PubMed] [Google Scholar]
  20. Zhang T, Wu Q, Zhang Z. Probable pangolin origin of SARS-CoV-2 associated with the COVID-19 outbreak. Curr. Biol. 30(7), 1346–1351 (2020). [PMC free article] [PubMed] [Google Scholar]
  21. an W, Lu Y, Guo Y. et al. Viral kinetics and antibody responses in patients with COVID-19. medRxiv (2020) (Epub ahead of print). [Google Scholar]
  22. Yang Y, Yang M, Shen C. et al. Evaluating the accuracy of different respiratory specimens in the laboratory diagnosis and monitoring the viral shedding of 2019-nCoV infections. medRxiv. (2020) (Epub ahead of print). [Google Scholar]
  23. Walls AC, Park YJ, Tortorici MA, Wall A, McGuire AT, Veesler D. Structure, function, and antigenicity of the SARS-CoV-2 spike glycoprotein. Cell 181 (2), 281–292 (2020). [PMC free article] [PubMed] [Google Scholar]
  24. Letko M, Marzi A, Munster V. Functional assessment of cell entry and receptor usage for SARS-CoV-2 and other lineage B betacoronaviruses. Nat. Microbiol. 5(4), 562–569 (2020). [PMC free article] [PubMed] [Google Scholar]
  25. Hoffmann M, Kleine-Weber H, Schroeder S. et al. SARS-CoV-2 cell entry depends on ACE2 and TMPRSS2 and is blocked by a clinically proven protease inhibitor. Cell 181(2), 271–280 (2020). [PMC free article] [PubMed] [Google Scholar]
  26. Hamming I, Timens W, Bulthuis MLC, Lely AT, Navis GJ, Goor Hvan. Tissue distribution of ACE2 protein, the functional receptor for SARS coronavirus. A first step in understanding SARS pathogenesis. J. Pathol. 203(2), 631–637 (2004). [PMC free article] [PubMed] [Google Scholar]
  27. Xu Z, Shi L, Wang Y. et al. Pathological findings of COVID-19 associated with acute respiratory distress syndrome. Lancet Respir. Med. 8(4), 420–422 (2020). [PMC free article] [PubMed] [Google Scholar]
  28. Gao QY, Chen YX, Fang JX. 2019 novel coronavirus infection and gastrointestinal tract. J. Dig. Dis. 21(3), 125–126 (2020). [PMC free article] [PubMed] [Google Scholar]
  29. Lee SH. The SARS epidemic in Hong Kong. J. Epidemiol. Community Health 57(9), 652–654 (2003). [PMC free article] [PubMed] [Google Scholar]
  30.  Leung WK, To KF, Chan PK. et al. Enteric involvement of severe acute respiratory syndrome-associated coronavirus infection. Gastroenterology 125(4), 1011–1017 (2003). [PMC free article] [PubMed] [Google Scholar]
  31. Parri N, Lenge M, Buonsenso D. Coronavirus Infection in Pediatric Emergency Departments (CONFIDENCE) Research Group. Children with Covid-19 in pediatric emergency departments in Italy. N. Engl. J. Med. (2020) (Epub ahead of print]. [PMC free article] [PubMed] [Google Scholar]
  32. Buonsenso D, Zampino G, Valentini P. Novel coronavirus disease 2019 infection in children: the dark side of a worldwide outbreak. Front. Pediatr. 8, 215 (2020). [PMC free article] [PubMed] [Google Scholar]
  33. Yao HP, Lu XY, Chen Q. et al. Patient-derived mutations impact pathogenicity of SARS-CoV-2. medRxiv (2020) (Epub ahead of print). [Google Scholar]•• Live SARS-CoV-2 virus was isolated from faecal samples in three of 11 adult patients in this research, implying infective faecal of COVID-19 patients.
  34. Hu Y, Shen L, Yao Y, Xu Z, Zhou J, Zhou H. A report of three COVID-19 cases with prolonged viral RNA detection in anal swabs. Clin. Microbiol. Infect. 26(6), 786–787 (2020). [PMC free article] [PubMed] [Google Scholar]
  35. Chen W, Lan Y, Yuan X. et al. Detectable 2019-nCoV viral RNA in blood is a strong indicator for the further clinical severity. Emerg. Microbes Infect. 9(1), 469–473 (2020). [PMC free article] [PubMed] [Google Scholar]
  36. Sethuraman N, Jeremiah SS, Ryo A. Interpreting diagnostic tests for SARS-CoV-2. JAMA (2020) (Epub ahead of print). [PubMed] [Google Scholar]• Describes how to interpret two types of diagnostic tests commonly in use for SARS-CoV-2 infections- RT-PCR and IgM and IgG ELISA – and how the results may vary over time. A clinically useful timeline of diagnostic markers for detection of COVID-19 was devised.

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