唾液によるCoVID-19の診断価値

最もSARS-CoV-2のウイルスを検出するために、2020年4月時点で、痰が最も感度が高いのですが、次に鼻腔で咽頭はこれらに劣るとされていました。

CoVID-19は鼻汁が殆ど出ない疾患で、鼻腔に綿棒を入れる検査は辛いものです。

SARS-CoV-2は肺の分泌腺を破壊することで、サーファクタントが出なくなり、呼吸障害が進行します。

唾液腺にも同様にACE2に発現していて、唾液にSARS-CoV-2が分泌されるために、鼻腔検査よりも優れている可能性が出てきました。

また唾液でRNAが陽性の人は、喋ることで他人へ感染させるリスクも高く、行動制限の指標としての実用的価値も高いと思われます。

（※　管理者注　2020/04/28記載）

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進行中のCOVID-19のパンデミックを制御するには、迅速で正確なSARS-CoV-2診断テストが不可欠です。

COVID-19診断の現在のゴールドスタンダードは、鼻咽頭スワブからのSARS-CoV-2のリアルタイムRT-PCR検出です。

ただし、感度が低く、医療従事者への曝露リスク、および綿棒と個人用保護具の世界的な不足のため、新しい診断アプローチの検証が必要です。唾液は、以前の研究でSARS-CoV-2診断の有望な候補です。これは、

（1）収集の侵襲が最小限であり、確実に自己投与できること、および

（2）唾液は、ヒトコロナウイルスを含む他の呼吸器病原体の検出において鼻咽頭スワブに匹敵する感度を示したこと、を示します。

SARS-CoV-2検出での唾液の使用を検証するために、確認されたCOVID-19患者からの鼻咽頭および唾液サンプルと、COVID-19病棟の医療従事者からの自己収集サンプルをテストしました。

患者が一致する上咽頭と唾液のサンプルからのSARS-CoV-2検出を比較すると、唾液が感染の過程全体で検出感度と一貫性に優れていることがわかりました。

さらに、唾液の自己サンプルコレクションの変動が少ないことを報告します。

併せて私たちの調査結果は、唾液が上咽頭スワブの実行可能でより敏感な代替物であり、正確な大規模SARS-CoV-2テストのために自宅での自己管理型のサンプル収集を可能にする可能性があることを示しています。

＜イントロダクション＞

SAV-CoV-2、COVID-19パンデミックを引き起こす新規コロナウイルスを制御するための取り組みは、正確で迅速な診断テストに依存しています。これらの検査は、

（1）効果的な自己隔離を促進し、ハイリスクグループ内の感染を減らすために、軽度で無症候性の感染に敏感でなければなりません。

（2）疾患の進行を確実に監視し、一貫して臨床上の決定を支援する。

（3）社会的離隔措置を安全に緩和できる場合など、地域および国の公衆衛生政策を通知するための拡張性を提供する。

ただし、現在のSARS-CoV-2テスト戦略は、リアルタイムRT-PCRで広く推奨されているサンプルタイプとして鼻咽頭スワブに依存しているため、これらの基準を満たさないことがよくあります。

鼻咽頭スワブは呼吸器ウイルスの診断によく使用されますが、初期感染ではSARS-CoV-2検出の感度が比較的低く、連続したテスト中では一貫性がありません。

さらに、鼻咽頭スワブを採取すると、手技の侵襲性のために患者に不快感を与え、繰り返し検査のコンプライアンスが制限され、患者がくしゃみや咳をしたり、ウイルス粒子を排出したりする可能性があるため、医療従事者にかなりのリスクをもたらします。

医療従事者のための綿棒と個人用保護具の不足が広がっており、鼻咽頭綿棒の自己採取は困難であり、ウイルス検出の感度が低いため、この手順は大規模な検査にも役立ちません。

COVID-19のパンデミックが低所得国で激化するにつれて、これらの課題はさらに悪化します。

制限があるため、より信頼性が高く、リソースをあまり必要としないサンプル収集方法、理想的には自宅での自己収集に対応する方法が緊急に必要です。

唾液の採取は非侵襲的で自己管理が容易であるため、唾液の採取は鼻咽頭スワブの魅力的な代替手段です。

2つの季節性ヒトコロナウイルスを含む呼吸器病原体のRT-PCR検出に対する鼻咽頭と唾液の一致の分析は、2つのサンプルタイプ間で同等の診断感度を示唆しています。

予備調査結果は、

（1）SARS-CoV-2がCOVID-19患者の唾液から検出できること、および

（2）自己採取した唾液サンプルが、医療従事者が軽度そして無症状のCOVID-19症例から採取した鼻咽頭スワブに匹敵するSARS-CoV-2検出感度を持つことを示しています。

しかし批判的には、鼻咽頭スワブに関する唾液中のSARS-CoV-2検出の感度の厳密な評価は、COVID-19感染の経過中に入院患者から行われていません。

この研究では、COVID-19入院患者と無症候性の医療従事者からCOVID-19曝露のリスクが中程度から高いリスクで収集された鼻咽頭スワブと唾液のペアでSARS-CoV-2検出を評価しました。

私たちの結果は、SARS-CoV-2検出に唾液を使用する方が、鼻咽頭スワブを使用するよりも感度が高く、一貫していることを示しています。

全体として、COVID-19テストの要求を軽減するためには、唾液を信頼できるサンプルタイプと見なす必要があることを示しています。

＜結果＞

＜入院患者の鼻咽頭スワブより唾液から検出されたSARS-CoV-2力価が高い。＞

SARS-CoV-2診断に鼻咽頭スワブを使用するという米国のCDC勧告と同様に唾液が機能するかどうかを判断するために、COVID-19入院研究参加者44人から臨床サンプルを収集しました（表1）。

このコホートは、2020年4月5日の時点で、19人（43％）が集中治療を必要とし、10人（23％）が人工呼吸器換気を必要とし、2人（5％）が死亡した、重症のCOVID-19患者の範囲を表しています。

CDC SARS-CoV-2 RT-PCRアッセイでは、このコホートから121例の自己採取した唾液または医療従事者が投与した鼻咽頭スワブをテストしました。

米国のCDCの「N1」と「N2」のプライマープローブセット（拡張データ図1）の間に強い一致が見られ、「N1」セットのみを使用してウイルス力価（ウイルスコピー/ mL）が計算されました。

テストしたすべての陽性サンプル（鼻咽頭:n = 46、唾液:n=37）から、唾液からの幾何平均ウイルス力価が鼻咽頭スワブよりも約5倍高いことがわかりました（p <0.05、Mann-Whitney検定;図1a）。

患者の一致する鼻咽頭と唾液のサンプルのみに分析を限定すると（各サンプルタイプについてn = 38）、唾液からのSARS-CoV-2力価は鼻咽頭スワブよりも有意に高いことがわかりました（p = 0.0001、Wilcoxon検定;図1b）。

さらに、8つのサンプル（21％）では鼻咽頭スワブから検出せず、唾液からのみSARS-CoV-2を検出しました。3つのサンプル(8％)では鼻咽頭スワブのみからSARS-CoV-2を検出しました 。(1c)

全体的に、病院の入院患者の鼻咽頭スワブよりも唾液のSARS-CoV-2力価が高いことがわかりました。

＜入院患者の唾液を検査する際の一時的なSARS-CoV-2変動＞

鼻咽頭スワブからの一時的なSARS-CoV-2診断テストは変数すると報告されており、入院患者の縦鼻咽頭と唾液のサンプルをテストして、どのサンプルタイプがより一貫した結果となるかを判断しました。

複数の上咽頭スワブを使用した22人の参加者と複数の唾液サンプルを使用した12人の参加者から、SARS-CoV-2力価は一般的に、発症日以降、両方のサンプルで次第に低下することがわかりました（図2a）。

私たちの上咽頭スワブの結果は、SARS-CoV-2力価が変動するという以前の報告と一致しています：

参加者の上咽頭スワブがSARS-CoV-2について陰性であり、次の収集時に陽性結果が続いた5例を見つけました（5/33例。繰り返し33％;図2b）。

ただし、12人の患者からの長期的な唾液採取では、サンプルが陰性であり、その後陽性の結果が得られた事例はありませんでした。

真の陰性の検査結果は臨床医が患者の改善を追跡し、退院に関する決定を下すために重要であるため、私たちのデータは、唾液がSARS-CoV-2力価の時間変化を監視する上咽頭スワブよりも一貫したサンプルタイプであることを示唆しています。

＜唾液を使用する医療従事者からのより一貫した自己サンプリング＞

無症状のSARS-CoV-2感染を検出するための唾液の検証は、遠隔患者の診断と医療従事者の監視の両方に変革をもたらす可能性があります。

これを調査するために、無症状の医療従事者98人を調査に登録し、平均2.9日ごと(範囲= 1〜8日、表2)に唾液および/または鼻咽頭スワブを収集しました。

これまでに、US CDCの「N1」および「N2」テストの両方を使用して鼻咽頭スワブで陰性であり、症状を報告しなかった2人の医療従事者の唾液からSARS-CoV-2を検出しました。

これらの個人の1人の唾液は、2日後の繰り返し検査で、一致する鼻咽頭スワブとともに陽性と再び検査されました。

無症候性の医療従事者の唾液からのウイルス力価は、症候性の入院患者から通常検出される値よりも低く（図3a）、症状の欠如を説明できる可能性があります。

私たちの限られたデータは、唾液が無症候性または発症前の感染を検出するための感度が高い可能性があります。ただし、確認するにはより大きなサンプルサイズが必要です。

鼻咽頭スワブサンプリングの不整合は、偽陰性の潜在的な問題の1つである可能性があるため（図2）、適切なサンプル収集のために内部コントロールをモニタリングすることにより、ヒトRNase Pは、代替の評価手法を提供します。

人間のRNase P検出は、入院患者コホートと医療従事者コホートの両方で唾液からの方が優れていましたが（図3b）、これだけではウイルス検出が良好であるとは限りません。

さらに重要なことに、人間のRNase P検出は、入院患者から収集された鼻咽頭スワブ（p = 0.0001、分散のF検定）および医療従事者から自己収集された（p = 0.0002;図3b）の方が変動が大きいことがわかりました。

私たちの結果は、無症候性または発症前のSARS-CoV-2感染のスクリーニングには、唾液が鼻咽頭スワブに代わる適切な、そしておそらくより感度の高い代替手段である可能性があることを示唆しています。

＜議論＞

私たちの研究は、唾液がSARS-CoV-2検出のための鼻咽頭スワブの実行可能で好ましい代替手段であることを示しています。唾液からのSARS-CoV-2検出の感度は、入院初期の鼻咽頭スワブよりも優れているとは言えないが同等であり、長期の入院および回復時により一貫していることがわかりました。

さらに、鼻咽頭スワブが検出できないにもかかわらず、2人の無症候性医療従事者の唾液からSARS-CoV-2が検出された場合、唾液も軽度または無症状の感染を特定するための実行可能な代替手段になる可能性があります。

更なる検証により、唾液サンプリングの広範な実装は、公衆衛生の取り組みにとって変革となる可能性があります。唾液の自己採取は、医療従事者と患者の直接的な相互作用の必要性、いくつかの主要なテストボトルネックの原因である綿棒と個人用防護具の問題、および院内感染リスクを軽減します。

SARS-CoV-2のウイルス量は軽度と重症の場合で異なるため、本研究の制限はCOVID-19入院患者に主に焦点を当てており、その多くは重症の疾患です。

病院環境での唾液の有効性を、感染過程の初期でより厳密に比較するには、より多くのデータが必要ですが、最近の2つの研究の結果は、無症候性の個人と外来患者の両方からSARS-CoV-2を検出できる可能性を裏付けています。

COVID-19患者の唾液から感染性ウイルスが検出されたため、無症候性感染のダイナミクスを理解する上で、ウイルスゲノムのコピーと感染前ウイルスの唾液中の感染性ウイルス粒子との関係を確認することが重要な役割を果たします。

無症候性の個人でのSARS-CoV-2検出の有望な結果に基づいて、唾液SARS-CoV-2検出アッセイはすでに米国食品医薬品局の緊急使用許可を通じて承認を得ています。

しかし、増大するテストの需要を満たすために、認定された臨床検査室でSARS-CoV-2診断用の唾液を即座に検証および実装する必要性があります。

＜採取方法＞

入院患者の場合：

上咽頭および唾液のサンプルは、臨床経過を通じて3日ごとに取得されました。

鼻咽頭サンプルは、BDユニバーサルウイルス輸送（UVT）システムを使用して登録看護師が採取しました。

柔軟なミニチップ綿棒を後鼻咽頭に到達するまで患者の鼻孔に通し、数秒間そのままにして分泌物を吸収させ、回転させながらゆっくりと取り出しました。

綿棒を無菌ウイルス輸送培地（総容量3mL）に入れ、しっかりと密封した。唾液サンプルは患者が自己採取した。覚醒時に、患者は、サンプルが収集されるまで、食物、水、および歯磨きを避けるように求められました。患者は、液体が約3分の1になるまで（気泡を除く）、無菌尿カップに繰り返し吐き出すように求められ、その後、しっかりと閉じました。すべてのサンプルは室温で保管され、サンプル収集後5時間以内にイェール公衆衛生学校の研究所に輸送されました。

医療従事者の場合：

医療従事者は、自己投与された鼻咽頭スワブと唾液のサンプルを2週間にわたって3日ごとに収集するように求められました。サンプルは、研究所に輸送されるまで+ 4°Cで保管されました。

＜SARS-CoV-2検出方法＞

研究室に到着したら、製造元のプロトコルに従ってMagMAX Viral / Pathogen Nucleic Acid Isolationキット（ThermoFisher Scientific）を使用して、鼻咽頭スワブから 75μlの溶出バッファーに入れ、300μlのウイルス輸送培地または300μlの唾液全体から総核酸を抽出しました。

SARS-CoV-2 RNA検出には、2019-nCoV_N1および2019-nCoV_N2およびヒトのUS CDCリアルタイムRT-PCRプライマー/プローブセットを使用して、前述のように5μlのRNAテンプレートをテストしました。

抽出コントロールとしてのRNase P（RP）を使用しました。 N1とN2の両方のプライマープローブセットが<38 CTで検出された場合、サンプルはSARS-CoV-2陽性と分類されました。以前に生成したRNA転写産物の10倍希釈標準曲線を使用してウイルスコピーを定量化しました。 N1とN2の結果は同等であったため（拡張データ図1）、すべてのウイルスコピーはN1プライマープローブセットを用いて計算された。

Saliva is more sensitive for SARS-CoV-2 detection in COVID-19 patients than nasopharyngeal swabs

View ORCID ProfileAnne Louise Wyllie, John Fournier, Arnau Casanovas-Massana, Melissa Campbell, Maria Tokuyama, Pavithra Vijayakumar, Bertie Geng, M. Catherine Muenker, Adam J. Moore, Chantal B. F. Vogels, Mary E. Petrone, Isabel M. Ott, Peiwen Lu, Alice Lu-Culligan, Jonathan Klein, Arvind Venkataraman, Rebecca Earnest, Michael Simonov, Rupak Datta, Ryan Handoko, Nida Naushad, Lorenzo R. Sewanan, Jordan Valdez, Elizabeth B. White, Sarah Lapidus, Chaney C. Kalinich, Xiaodong Jiang, Daniel J. Kim, Eriko Kudo, Melissa Linehan, Tianyang Mao, Miyu Moriyama, Ji Eun Oh, Annsea Park, Julio Silva, Eric Song, Takehiro Takahashi, Manabu Taura, Orr-El Weizman, Patrick Wong, Yexin Yang, Santos Bermejo, Camila Odio, Saad B. Omer, Charles S. Dela Cruz, Shelli Farhadian, Richard A. Martinello, Akiko Iwasaki, Nathan D. Grubaugh, Albert I. Ko

doi: https://doi.org/10.1101/2020.04.16.20067835

https://www.medrxiv.org/content/10.1101/2020.04.16.20067835v1

Abstract

Rapid and accurate SARS-CoV-2 diagnostic testing is essential for controlling the ongoing COVID-19 pandemic.

The current gold standard for COVID-19 diagnosis is real-time RT-PCR detection of SARS-CoV-2 from nasopharyngeal swabs.

Low sensitivity, exposure risks to healthcare workers, and global shortages of swabs and personal protective equipment, however, necessitate the validation of new diagnostic approaches. Saliva is a promising candidate for SARS-CoV-2 diagnostics because (1) collection is minimally invasive and can reliably be self-administered and (2) saliva has exhibited comparable sensitivity to nasopharyngeal swabs in detection of other respiratory pathogens, including endemic human coronaviruses, in previous studies.

To validate the use of saliva for SARS-CoV-2 detection, we tested nasopharyngeal and saliva samples from confirmed COVID-19 patients and self-collected samples from healthcare workers on COVID-19 wards.

When we compared SARS-CoV-2 detection from patient-matched nasopharyngeal and saliva samples, we found that saliva yielded greater detection sensitivity and consistency throughout the course of infection.

Furthermore, we report less variability in self-sample collection of saliva.

Taken together, our findings demonstrate that saliva is a viable and more sensitive alternative to nasopharyngeal swabs and could enable at-home self-administered sample collection for accurate large-scale SARS-CoV-2 testing.

Introduction

Efforts to control SARS-CoV-2, the novel coronavirus causing COVID-19 pandemic, depend on accurate and rapid diagnostic testing. These tests must be ( 1 ) sensitive to mild and asymptomatic infections to promote effective self isolation and reduce transmission within high risk groups1 ; ( 2) consistent to reliably monitor disease progression and aid clinical decisions2 ; and ( 3) scalable to inform local and national public health policies, such as when social distancing measures can be safely relaxed.

However, current SARS-CoV-2 testing strategies often fail to meet these criteria, in part because of their reliance on nasopharyngeal swabs as the widely recommended sample type for real-time RT-PCR.

Although nasopharyngeal swabs are commonly used in respiratory virus diagnostics, they show relatively poor sensitivity for SARS-CoV-2 detection in early infection and are inconsistent during serial testing2 –6.

Moreover, collecting nasopharyngeal swabs causes discomfort to patients due to the procedure’s invasiveness, limiting compliance for repeat testing, and presents a considerable risk to healthcare workers, because it can induce patients to sneeze or cough, expelling virus particles 7 .

The procedure is also not conducive to large-scale testing, because there are widespread shortages of swabs and personal protective equipment for healthcare workers 8 , and self-collection of nasopharyngeal swabs is difficult and less sensitive for virus detection 9 .

These challenges will be further exacerbated as the COVID-19 pandemic intensifies in low income countries.

Given the limitations, a more reliable and less resource-intensive sample collection method, ideally one that accommodates self-collection in the home, is urgently needed.

Saliva sampling is an appealing alternative to nasopharyngeal swab, since collecting saliva is non-invasive and easy to self-administer.

An analysis of nasopharyngeal and saliva concordance for RT-PCR detection of respiratory pathogens, including two seasonal human coronaviruses, suggests comparable diagnostic sensitivity between the two sample types.

Preliminary findings indicate that ( 1 ) SARS-CoV-2 can be detected from the saliva of COVID-19 patients and ( 2 ) self-collected saliva samples have comparable SARS-CoV-2 detection sensitivity to nasopharyngeal swabs collected by healthcare workers from mild and subclinical COVID-19 cases.

Critically, however, no rigorous evaluation of the sensitivity of SARS-CoV-2 detection in saliva with respect to nasopharyngeal swabs has been conducted from inpatients during the course of COVID-19 infection.

In this study, we evaluated SARS-CoV-2 detection in paired nasopharyngeal swabs and saliva samples collected from COVID-19 inpatients and asymptomatic healthcare workers at moderate-to-high risk of COVID-19 exposure.

Our results indicate that using saliva for SARS-CoV-2 detection is more sensitive and consistent than using nasopharyngeal swabs.

Overall, we demonstrate that saliva should be considered as a reliable sample type to alleviate COVID-19 testing demands.

Results

Higher SARS-CoV-2 titers detected from saliva than nasopharyngeal swabs from inpatients

To determine if saliva performs as well as the U.S. CDC recommendation of using nasopharyngeal swabs for SARS-CoV-2 diagnostics, we collected clinical samples from 44 COVID-19 inpatient study participants ( Table 1 ).

This cohort represents a range of COVID-19 patients with severe disease, with 19 (43%) requiring intensive care, 10 (23%) requiring mechanical ventilation, and 2 (5%) deceased as of April 5th, 2020. Using the U.S. CDC SARS-CoV-2 RT-PCR assay, we tested 121 self-collected saliva or healthcare worker-administered nasopharyngeal swabs from this cohort.

We found strong concordance between the U.S. CDC “N1” and “N2” primer-probe sets ( Extended Data Fig. 1 ), and thus calculated virus titers (virus copies/mL) using only the “N1” set.

From all positive samples tested ( n = 46 nasopharyngeal, 37 saliva), we found that the geometric mean virus titers from saliva were about 5⨉ higher than nasopharyngeal swabs ( p < 0.05, Mann-Whitney test; Fig. 1a ).

When limiting our analysis to only patient-matched nasopharyngeal and saliva samples ( n = 38 for each sample type), we found that SARS-CoV-2 titers from saliva were significantly higher than nasopharyngeal swabs ( p = 0.0001, Wilcoxon test; Fig. 1b ).

Moreover, we detected SARS-CoV-2 from the saliva but not the nasopharyngeal swabs from eight matching samples (21%), while we only detected SARS-CoV-2 from nasopharyngeal swabs and not saliva from three matched samples (8%; Fig. 1c ).

Overall, we found higher SARS-CoV-2 titers from saliva than nasopharyngeal swabs from hospital inpatients.

Table 1. COVID-19 inpatient cohort characteristics

Figure 1. SARS-CoV-2 titers are higher in the saliva than nasopharyngeal swabs from hospital inpatients.

( a ) All positive nasopharyngeal swabs ( n = 46) and saliva samples ( n = 39) were compared by a Mann-Whitney test ( p < 0.05). Bars represent the median and 95% CI. Our assay detection limits for SARS-CoV-2 using the US CDC “N1” assay is at cycle threshold 38, which corresponds to 5,610 virus copies/mL of sample (shown as dotted line and grey area).

( b ) Patient matched samples ( n = 38), represented by the connecting lines, were compared by a Wilcoxon test test ( p < 0.05).

( c ) Patient matched samples ( n = 38) are also represented on a scatter plot. All of the data used to generate this figure, including the raw cycle thresholds, can be found in Supplementary Data 1 . Extended Data Fig. 1 shows the correlation between US CDC assay “N1” and “N2” results.

Less temporal SARS-CoV-2 variability when testing saliva from inpatients

As temporal SARS-CoV-2 diagnostic testing from nasopharyngeal swabs is reported to be variable2 ,3, we tested longitudinal nasopharyngeal and saliva samples from inpatients to determine which sample type provided more consistent results.

From 22 participants with multiple nasopharyngeal swabs and 12 participants with multiple saliva samples, we found that SARS-CoV-2 titers generally decreased in both sample types following the reported date of symptom onset ( Fig. 2a ).

Our nasopharyngeal swab results are consistent with previous reports of variable SARS-CoV-2 titers and results2 ,3: we found 5 instances where a participant’s nasopharyngeal swab was negative for SARS-CoV-2 followed by a positive result during the next collection (5/33 repeats, 33%; Fig. 2b) .

In longitudinal saliva collections from 12 patients, however, there were no instances in which a sample tested negative and was later followed by a positive result.

As true negative test results are important for clinicians to track patient improvements and for decisions regarding discharges, our data suggests that saliva is a more consistent sample type than nasopharyngeal swabs for monitoring temporal changes in SARS-CoV-2 titers.

Figure 2: SARS-CoV-2 detection is less variable between repeat sample collections with saliva.

(a)Longitudinal SARS-CoV-2 titers from saliva or nasopharyngeal swabs are shown as days since symptom onset.

Each circle represents a separate sample, which are connected to additional samples from the same patient by a dashed line.

Our assay detection limits for SARS-CoV-2 using the US CDC “N1” assay is at cycle threshold 38, which corresponds to 5,610 virus copies/mL of sample (shown as dotted line and grey area).

( b ) The data are also shown by sampling moment (sequential collection time) to highlight the differences in virus titers between collection points.

All of the data used to generate this figure, including the raw cycle thresholds, can be found in Supplementary Data 1 .

More consistent self-sampling from healthcare workers using saliva

Validating saliva for the detection of subclinical SARS-CoV-2 infections could prove transformative for both remote patient diagnostics and healthcare worker surveillance.

To investigate this, we enrolled 98 asymptomatic healthcare workers into our study and collected saliva and/or nasopharyngeal swabs on average every 2.9 days (range = 1-8 days, Table 2 ).

To date, we have detected SARS-CoV-2 in saliva from two healthcare workers who were negative by nasopharyngeal swabs using both the US CDC “N1” and “N2” tests and did not report any symptoms.

The saliva from one of these individuals again tested positive alongside a matching negative nasopharyngeal swab upon repeat testing 2 days later.

Virus titers from asymptomatic healthcare workers’ saliva are lower than what we typically detect from symptomatic inpatients ( Fig. 3a ), which likely supports the lack of symptoms.

Our limited data supports that saliva may be more sensitive for detecting asymptomatic or pre-symptomatic infections; however, a larger sample size is needed to confirm.

As nasopharyngeal swab sampling inconsistency may be one of the potential issues for false negatives ( Fig. 2 ), monitoring an internal control for proper sample collection, human RNase P, may provide an alternative evaluation technique.

While human RNase P detection was better from saliva in both the inpatient and healthcare worker cohorts ( Fig. 3b) , this alone may not indicate better virus detection.

More importantly, we found that human RNase P detection was more variable from nasopharyngeal swabs collected from inpatients ( p = 0.0001, F test for variances) and self-collected from healthcare workers ( p = 0.0002; Fig. 3b ).

Our results suggest that saliva may also be an appropriate, and perhaps more sensitive, alternative to nasopharyngeal swabs for screening asymptomatic or pre-symptomatic SARS-CoV-2 infections.

Table 2. Healthcare worker cohort

Figure 3. Saliva is an alternative for SARS-CoV-2 screening from healthcare workers and asymptomatic cases.

( a ) SARS-CoV-2 titers measured from the saliva of healthcare workers and inpatients. Our assay detection limits for SARS-CoV-2 using the US CDC “N1” assay is at cycle threshold 38, which corresponds to 5,610 virus copies/mL of sample (shown as dotted line and grey area).

( b ) RT-PCR cycle thresholds (Ct) values for human RNase P, and internal control for sample collection, from either inpatients (left panel) or health care workers (right panel) were compared by variances using the F test ( p = 0.0001 for inpatients; p = 0.0002 for healthcare workers).

All of the data used to generate this figure, including the raw cycle thresholds, can be found in Supplementary Data 1 .

Discussion

Our study demonstrates that saliva is a viable and preferable alternative to nasopharyngeal swabs for SARS-CoV-2 detection. We found that the sensitivity of SARS-CoV-2 detection from saliva is comparable, if not superior to nasopharyngeal swabs in early hospitalization and is more consistent during extended hospitalization and recovery.

Moreover, the detection of SARS-CoV-2 from the saliva of two asymptomatic healthcare workers despite negative matched nasopharyngeal swabs suggests that saliva may also be a viable alternative for identifying mild or subclinical infections.

With further validation, widespread implementation of saliva sampling could be transformative for public health efforts: saliva self-collection negates the need for direct healthcare worker-patient interaction, a source of several major testing bottlenecks and overall nosocomial infection risk, and alleviates supply demands on swabs and personal protective equipment.

As SARS-CoV-2 viral loads differ between mild and severe cases, a limitation of our study is the primary focus on COVID-19 inpatients, many with severe disease.

While more data are required to more rigorously compare the efficacy of saliva in the hospital setting to earlier in the course of infection, findings from two recent studies support its potential for detecting SARS-CoV-2 from both asymptomatic individuals and outpatients.

As infectious virus has been detected from the saliva of COVID-19 patients, ascertaining the relationship between virus genome copies and infectious virus particles in the saliva of pre-symptomatic individuals will play a key role in understanding the dynamics of asymptomatic transmission.

Stemming from the promising results for SARS-CoV-2 detection in asymptomatic individuals, a saliva SARS-CoV-2 detection assay has already gained approval through the U.S. Food and Drug Administration emergency use authorization.

To meet the growing testing demands, however, our findings support the need for immediate validation and implementation of saliva for SARS-CoV-2 diagnostics in certified clinical laboratories.

Methods

Ethics

All study participants were enrolled and sampled in accordance to the Yale University HIC-approved protocol #2000027690.

Demographics, clinical data and samples were only collected after the study participant had acknowledged that they had understood the study protocol and signed the informed consent.

All participant information and samples were collected in association with study identifiers.

Participant enrollment

Inpatients

Patients admitted to Yale New Haven Hospital (a 1541-bed tertiary care medical center in New Haven, CT, USA), who tested positive for SARS-CoV-2 by nasopharyngeal and/or oropharyngeal swab (CDC approved assay) were invited to enroll in the research study.

Exclusion criteria were age under 18 years, non-English speaking and clinical, radiological or laboratory evidence for a non-infectious cause of fever or respiratory symptoms or a microbiologically-confirmed infectious source (e.g. gastrointestinal, urinary, cardiovascular) other than respiratory tract for symptoms and no suspicion for COVID-19 infection.

Healthcare workers

Asymptomatic healthcare workers (e.g., without fever or respiratory symptoms) with occupational exposure to patients with COVID-19 were invited to enroll in the study. Study participation enabled active surveillance to ensure early detection following exposure and to further protect other healthcare workers and patients.

Sample collection

Inpatients

Nasopharyngeal and saliva samples were obtained every three days throughout their clinical course.

Nasopharyngeal samples were taken by registered nurses using the BD universal viral transport (UVT) system.

The flexible, mini-tip swab was passed through the patient's nostril until the posterior nasopharynx was reached, left in place for several seconds to absorb secretions then slowly removed while rotating.

The swab was placed in the sterile viral transport media (total volume 3 mL) and sealed securely.

Saliva samples were self-collected by the patient.

Upon waking, patients were asked to avoid food, water and brushing of teeth until the sample was collected. Patients were asked to repeatedly spit into a sterile urine cup until roughly a third full of liquid (excluding bubbles), before securely closing it.

All samples were stored at room temperature and transported to the research lab at the Yale School of Public Health within 5 hours of sample collection.

Healthcare workers

Healthcare workers were asked to collect a self-administered nasopharyngeal swab and a saliva sample every three days for a period of 2 weeks.

Samples were stored at +4°C until being transported to the research lab.

SARS-CoV-2 detection

On arrival at the research lab, total nucleic acid was extracted from 300 μl of viral transport media from the nasopharyngeal swab or 300 μl of whole saliva using the MagMAX Viral/Pathogen Nucleic Acid Isolation kit (ThermoFisher Scientific) following the manufacturer's protocol and eluted into 75 μl of elution buffer.

For SARS-CoV-2 RNA detection, 5 μl of RNA template was tested as previously described2 1,22, using the US CDC real-time RT-PCR primer/probe sets for 2019-nCoV_N1 and 2019-nCoV_N2 and the human RNase P (RP) as an extraction control. Samples were classified as positive for SARS-CoV-2 when both N1 and N2 primer-probe sets were detected <38 CT .

Virus copies were quantified using a 10-fold dilution standard curve of RNA transcripts that we previously generated.

As results from N1 and N2 were comparable ( Extended Data Fig. 1 ), all virus copies are shown as calculated using the N1 primer-probe set.

Statistical analysis

Statistical analyses were conducted in GraphPad Prism 8.0.0 as described in the Results.

Acknowledgments

We gratefully acknowledge the study participants for their time and commitment to the study. We thank all members of the clinical team at Yale-New Haven Hospital for their dedication and work which made this study possible.

We also thank S. Taylor and P. Jack for technical discussions.

Funding

The study was partially funded by the Yale Institute for Global Health. The corresponding authors had full access to all data in the study and had final responsibility for the decision to submit for publication.

Extended data

Extended Data Fig. 1. Concordance between SARS-CoV-2 detection using US CDC “N1” and “N2” primer and probe sets. Ct = RT-PCR cycle threshold. Dotted line and grey areas indicate the limits of detection.

References

1. Kimball, A. et al. Asymptomatic and Presymptomatic SARS-CoV-2 Infections in Residents of a Long-Term Care Skilled Nursing Facility - King County, Washington, March 2020.

MMWR Morb. Mortal. Wkly. Rep. 69 , 377–381 (2020).

2. Wölfel, R. et al. Virological assessment of hospitalized patients with COVID-2019.

Nature (2020) doi: 10.1038/s41586-020-2196-x .

3. Zou, L. et al. SARS-CoV-2 Viral Load in Upper Respiratory Specimens of Infected Patients.

N. Engl. J. Med. 382 , 1177–1179 (2020).

4. Zhao, J. et al. Antibody responses to SARS-CoV-2 in patients of novel coronavirus disease 2019.

Clin. Infect. Dis. (2020) doi: 10.1093/cid/ciaa344 .

5. Xie, X. et al. Chest CT for Typical 2019-nCoV Pneumonia: Relationship to Negative RT-PCR Testing.

Radiology 200343 (2020).

6. Wang, W. et al. Detection of SARS-CoV-2 in Different Types of Clinical Specimens.

JAMA (2020) doi: 10.1001/jama.2020.3786 .

7. To, K. K.-W. et al. Temporal profiles of viral load in posterior oropharyngeal saliva samples and serum antibody responses during infection by SARS-CoV-2: an observational cohort study.

Lancet Infect. Dis. (2020) doi: 10.1016/S1473-3099(20)30196-1 .

8. CDC. Interim Infection Prevention and Control Recommendations for Patients with Suspected or Confirmed Coronavirus Disease 2019 (COVID-19) in Healthcare Settings. Centers for Disease Control and Prevention https://www.cdc.gov/coronavirus/2019-ncov/hcp/infection-control-recommendations.h tml?CDC_AA_refVal=https%3A%2F%2Fwww.cdc.gov%2Fcoronavirus%2F2019-ncov %2Finfection-control%2Fcontrol-recommendations.html (2020).

9. Dhiman, N. et al. Effectiveness of patient-collected swabs for influenza testing.

Mayo Clin. Proc. 87 , 548–554 (2012).

10. Kim, Y.-G. et al. Comparison between Saliva and Nasopharyngeal Swab Specimens for Detection of Respiratory Viruses by Multiplex Reverse Transcription-PCR.

J. Clin. Microbiol. 55 , 226–233 (2017).

11. Wyllie, A. L. et al. Molecular surveillance of nasopharyngeal carriage of Streptococcus pneumoniae in children vaccinated with conjugated polysaccharide pneumococcal vaccines.

Sci. Rep. 6 , 23809 (2016).

12. To, K. K.-W. et al. Consistent Detection of 2019 Novel Coronavirus in Saliva.

Clin. Infect. Dis. (2020) doi: 10.1093/cid/ciaa149 .

13. Kojima, N. et al. Self-Collected Oral Fluid and Nasal Swabs Demonstrate Comparable Sensitivity to Clinician Collected Nasopharyngeal Swabs for Covid-19 Detection.

medRxiv 2020.04.11.20062372 (2020).

14. Tran, K., Cimon, K., Severn, M., Pessoa-Silva, C. L. & Conly, J. Aerosol generating procedures and risk of transmission of acute respiratory infections to healthcare workers: a systematic review.

PLoS One 7 , e35797 (2012).

15. Judson, S. D. & Munster, V. J. Nosocomial Transmission of Emerging Viruses via Aerosol-Generating Medical Procedures.

Viruses 11 , (2019).

16. Wang, D. et al. Clinical Characteristics of 138 Hospitalized Patients With 2019 Novel Coronavirus-Infected Pneumonia in Wuhan, China.

JAMA (2020) doi: 10.1001/jama.2020.1585 .

17. Liu, Y. et al. Viral dynamics in mild and severe cases of COVID-19.

Lancet Infect. Dis. (2020) doi: 10.1016/S1473-3099(20)30232-2 .

18. U.S. Food & Drug Administration. Accelerated Emergency Use Authorization (EUA) Summary SARS-CoV-2 Assay (Rutgers Clinical Genomics Laboratory).

https://www.fda.gov/media/136875/download .

19. Lauer, S. A. et al. The Incubation Period of Coronavirus Disease 2019 (COVID-19) From Publicly Reported Confirmed Cases: Estimation and Application.

Ann. Intern. Med.(2020) doi: 10.7326/M20-0504 .

20. He, X. et al. Temporal dynamics in viral shedding and transmissibility of COVID-19.

medRxiv (2020).

21. Vogels, C. B. F. et al. Analytical sensitivity and efficiency comparisons of SARS-COV-2 qRT-PCR assays.

Infectious Diseases (except HIV/AIDS) (2020) doi: 10.1101/2020.03.30.20048108 .

23. CDC. Coronavirus Disease 2019 (COVID-19).

Centers for Disease Control and Prevention

https://www.cdc.gov/coronavirus/2019-ncov/lab/rt-pcr-panel-primer-probes.html (2020).

CoVID-19の流行は最初に2019年12月下旬に報告され、現在、世界的な大流行を引き起こしています。

唾液は、主に口腔の唾液腺から生成され、SARS-CoV-2のRNAが陽性であると報告されています。

肺に加えて、唾液腺と舌はACE2の発現によりSARS-CoV-2の肺外の宿主細胞となる可能性があります。

CoVID-19がWHOの主張どおりに広まる主な方法は、感染性唾液飛沫の密接な接触または短距離伝播ですが、長距離唾液エアロゾルの伝播は、歯科診療などのエアロゾル生成手順により、室内空間内の環境に大きく依存します。

これまでのところ、SARS-CoV-2が長期間にわたって空気の流れに存在できるという直接の証拠は見つかりませんでした。（※　管理者注　空気感染しないという意味か？）

したがって、感染性の唾液の飛沫の形成を防ぎ、室内の空気を完全に消毒し、唾液の飛沫の取得をブロックすると、SARS-CoV-2の伝播が遅くなる可能性があります。

このレビューでは、SARS-CoV-2の唾液の診断値、おそらく口腔組織への直接浸潤、および唾液滴によるSARS-CoV-2の接触感染を要約し、CoVID-19の流行抑制に寄与すると予想しています。

Int J Oral Sci. 2020 Apr 17;12(1):11.

doi: 10.1038/s41368-020-0080-z.

Saliva: Potential Diagnostic Value and Transmission of 2019-nCoV

Ruoshi Xu , Bomiao Cui , Xiaobo Duan , Ping Zhang , Xuedong Zhou , Quan Yuan

Affiliations expand

PMID: 32300101 PMCID: PMC7162686 DOI: 10.1038/s41368-020-0080-z

https://pubmed.ncbi.nlm.nih.gov/32300101/

Abstract

2019-nCoV epidemic was firstly reported at late December of 2019 and has caused a global outbreak of COVID-19 now.

Saliva, a biofluid largely generated from salivary glands in oral cavity, has been reported 2019-nCoV nucleic acid positive. Besides lungs, salivary glands and tongue are possibly another hosts of 2019-nCoV due to expression of ACE2. Close contact or short-range transmission of infectious saliva droplets is a primary mode for 2019-nCoV to disseminate as claimed by WHO, while long-distance saliva aerosol transmission is highly environment dependent within indoor space with aerosol-generating procedures such as dental practice. So far, no direct evidence has been found that 2019-nCoV is vital in air flow for long time. Therefore, to prevent formation of infectious saliva droplets, to thoroughly disinfect indoor air and to block acquisition of saliva droplets could slow down 2019-nCoV dissemination. This review summarizes diagnostic value of saliva for 2019-nCoV, possibly direct invasion into oral tissues, and close contact transmission of 2019-nCoV by saliva droplets, expecting to contribute to 2019-nCoV epidemic control.

Introduction

An outbreak of coronavirus disease (COVID-19) is emerging and rapidly spreading worldwide. 1

A public health emergency of international concern (PHEIC) was declared over COVID-19, which is the sixth time WHO has declared a PHEIC since the International Health Regulations took effect in 2005 (http://www.euro.who.int/en/health-topics/health-emergencies/international-health-regulations/news/news/2020/2/2019-ncov-outbreak-is-an-emergency-of-international-concern). 2

This new strain of disease was firstly reported in the late December of 2019 and has not been previously identified in human. The novel coronavirus isolated by researchers afterward was named as 2019 novel coronavirus (2019-nCoV). 2

Coronaviruses are enveloped RNA viruses, and two strains of them—severe acute respiratory syndrome coronavirus (SARS-CoV) and Middle East respiratory syndrome coronavirus (MERS-CoV)—are zoonotic in origin and known to cause fatal respiratory diseases as 2019-nCoV.

Due to wide distribution and genomes recombination of coronaviruses, 2019-nCoV is the successive but novel coronavirus and shown to have a higher rate of infection. 3–5

Early diagnosis of coronavirus and effective prevention of transmission are core tasks in control of 2019-nCoV epidemic.

WHO has claimed that 2019-nCoV spreads primarily through saliva droplets or discharge from the nose (https://www.who.int/health-topics/coronavirus#tab=tab_1).

Saliva is secreted 90% from major salivary glands and 10% from minor salivary glands within pH from 6 to 7. (6,7)

Whole saliva is a bio-mixture, which physiologically contains crevicular fluid, desquamated oral epithelial cells, and microorganisms, and may contain blood, respiratory secretions, gastric acid from reflux, and food debris in pathological occasions. 8

Around 99% of saliva is water and the rest 1% contains a large group of components for the purpose of digesting, taste, buffering, balance of remineralization, and anti-microorganisms. 9

Oral cavity is an entrance and an outlet of body, and saliva is supposed to play a role in early diagnosis and close contact transmission in infectious diseases.

Here, we summarize the reports associated with saliva and 2019-nCoV.

Diagnostic value of saliva for 2019-nCoV

The officially pathogen detection is the confirmation of 2019-nCoV nucleic acid from throat swabs. 10

Throat swabs are relatively invasive, induce coughing and cause bleeding occasionally, which may increase risks of healthcare workers infection.

Saliva stands at the entry of respiratory system and was also found 2019-nCoV nucleic acid positive. 11–14

With the nature of noninvasion and less hazard to healthcare workers, saliva specimen collection has the advantages of being more acceptable for patients and more secured for healthcare workers for diagnosis of coronavirus.

Till now, three approaches have been reported to collect saliva—coughing out, saliva swabs, and directly from salivary gland duct.

In two studies on coughed out saliva, 11 cases out of 12 (91.67%) and 20 cases out of 23 (86.96%) COVID-19 patients were 2019-nCoV RNA positive in saliva, respectively.

In one study of saliva swabs, half of 15 (50%) COVID-19 patients were 2019-nCoV RNA positive in saliva.

In one study of saliva directly from salivary gland duct, four cases of 31 (12.90%) COVID-19 patients were 2019-nCoV RNA positive in saliva, three of which were critically ill.

Early diagnosis of 2019-nCoV is still difficult, diagnostic value of saliva specimens for 2019-nCoV nucleic acid examination remains limited but promising, which we should still be cautious but expected about.

Deep throat saliva

A study from To et al. showed that deep throat saliva has high diagnosis rate of 2019-nCoV. 11

Twelve positive patients were confirmed based on epidemiological history, clinical criteria, and laboratory detection of 2019-nCoV in nasopharyngeal or sputum specimens, and saliva were collected by coughing out a few days after hospitalization. 11

Using real-time reverse transcription-quantitative polymerase chain reaction by testing the S gene of 2019-nCoV, 11 saliva specimens were positive for 2019-nCoV out of 12 patients (91.67%).11

Those 33 patients who are negative for laboratory test of 2019-nCoV were all negative in saliva examination.

In addition, six patients offer serial saliva, and five out of them showed a declining trend of virus as hospitalization is going on. 11

Live virus was detected in three patients of the above six patients by viral culture. 11

Another study from the same group used self-collected saliva from deep throat by COVID-19 patients, tested 2019-nCoV RNA, and analyzed temporal profile of 2019-nCoV load. 12

From this study, saliva mixed with nasopharyngeal and bronchopulmonary secretions from deep throat was collected by coughing out in the morning. 12

Among 23 COVID-19 patients included for this study, 20 cases of their saliva showed detectable 2019-nCoV RNA. 12

In the temporal profile of viral load, saliva reached the peak of viral load during the first week of symptom onset and then declined. 12

This group also detected 2019-nCoV RNA of saliva after treatment. 12

Even if using antibodies against 2019-nCoV, viral RNA could still be detected for 20 days or even longer in deep throat saliva specimens of one third of included patients, suggesting the viral RNA could stay a long period of time instead of dying out after antibody application. 12

One patient with complete symptom resolved was found 2019-nCoV RNA positive again after 2 days of negative results, suggesting that low levels of 2019-nCoV RNA could still be excreted in saliva even after clinical recovery. 12

More precisely, whether 2019-nCoV RNA detected in saliva after complete symptom resolved means infectious or shedding virus needs further studies to confirm.

Saliva in oral cavity

Oral swabs are probably applicable in early detection. 13

By harvesting oral swabs and testing RNA among 15 COVID-19 patients, Zhang et al. found that half of them (50%) were 2019-nCoV RNA positive in oral swabs, four (26.7%) had positive anal swabs, six (40%) had positive blood test, and three (20%) were serum positive. 13

Dynamic viral RNA presence in saliva compared with anal swabs were analyzed among 16 patients. Among all swab positive together, most of the positive result was from oral swabs at early stage, while more positive came from anal swabs at late stage of COVID-19, suggesting that oral swabs may indicate early infection of 2019-nCoV but cannot be used as a discharge criteria. 13

Salivary gland

To rule out contamination of respiratory secretion, Chen et al. collected saliva directly from the opening of salivary gland and found 2019-nCoV nucleic acid, suggesting that salivary glands were 2019-nCoV infected. 14

Thirteen cases who were nucleic acid positive by oropharyngeal swab among 31 COVID-19 patients were included, and four of them (12.90%) were positive in saliva. 14

Three cases of these four were critically ill patents in need of ventilator support, suggesting 2019-nCoV nucleic acid positive in salivary-gland-originated saliva as an indicator of severity of COVID-19.

Possible direct invasions into oral tissues

In the cycle of infection for most virus, the first step is to attach to the surface and recognize cell surface receptor of the host cell for invasion.15,16

With similar external subdomain of receptor-binding domain (RBD), 2019-nCoV spike share same host-cell receptor—angiotensin-converting enzyme II (ACE2)—with SARS-CoV spike, but in a higher affinity than SARS-CoV spike.(17–21)

In another word, cells expressing cell surface receptor ACE2 are susceptible to 2019-nCoV, similar to SARS-CoV. ACE2 was found expressed in lungs, esophagus, ileum, colon, cholangio of liver, and bladder. 22–25

Consistently, bronchoalveolar-lavage fluid,2 nasopharyngeal swabs,26 stool,27,28 and blood17 of COVID-19 patients were RT-PCR-positive for 2019-nCoV.

Several studies have shown that salivary gland and tongue express ACE2 receptor, suggesting oral cavity as host for 2019-nCoV to invade.

Expression of ACE2 in oral tissues

Xu et al. analyzed public bulk RNA-seq from paracarcinoma normal tissues and found expression of ACE2 in oral buccal and gingiva tissue.29

This group also analyzed data of single-cell RNA-seq from patients’ oral tissue and found that ACE2 were highly enriched in epithelial cells of tongue, and also in epithelial cells, T cells, B cells, and fibroblasts of oral mucosa. 29

Saliva is generated in salivary glands and flow through ducts into oral cavity.

Liu et al. analyzed rhesus macaques and found ACE2 were also expressed in epithelial cells lining on minor salivary gland ducts,30 which could be found in sinonasal cavity, oral cavity, pharynx, larynx, trachea, and lungs, amounting to 800–1000 individuals in total and contributing nearly 1% of saliva a day. 31

This group also set up animal models by inoculating functional pseudovirus intranasally, and found that ACE2+ epithelial cells of minor salivary gland ducts are targeted host cells as early as 48 h after infection.30

Besides above evidence from animal study, Chen et al. analyzed data from GTEx, HPA, FANTOM5, and consensus datasets, and revealed the expression of ACE2 receptor in human granular cells in salivary glands.14 ACE2+ cell in salivary glands could possibly be the target cells of 2019-nCoV and generate infectious saliva in sustained way theoretically.

Expression of furin on tongue

Furin has been implicated in virus infection by cleaving viral envelope glycoproteins and enhancing infection with host cells. 32

A furin-like cleavage site in the Spike protein of 2019-nCoV has been identified. 33,34

Furin is highly expressed in lung tissue, possibly providing a gain-of-function to infectivity of 2019-nCoV. 33,35,36

Furin expression was detected by immunostaining in human tongue epithelia, and significantly upregulated when squamous cell carcinoma (SCC) occured. 37

Combined with high expression of ACE2, tongue has high risk of coronavirus infection among oral cavity and SSC even increases the risk once exposed to coronavirus. While it suggests that cells expressing furin have lower restriction for virus entry theoretically, it should still be cautious whether the furin-like cleavage site plays a big role in 2019-nCoV infection. 36

Transmission of saliva 2019-nCoV

2019-nCoV transmission occurred within indoor space. 26

As noted that 2019-nCoV RNA is detected in saliva, whether 2019-nCoV in saliva could be disseminated by long-distance aerosol transmission is concerned by public.

WHO has claimed that droplets generated by an infected people by coughing, sneezing, or talking in close contact is the main routine of 2019-nCoV transmission besides touching contaminated surfaces without washing hands (https://www.who.int/news-room/q-a-detail/q-a-coronaviruses#).

WHO has updated the definition of close contact—any person within 1 m with a confirmed case at their symptomatic period, starting from 4 days before symptom onset. 38

However, airborne transmission could also be set up, especially within the same indoor space and aerosol-generating procedure is implemented.

Size of saliva droplets

Whether droplets can travel long and far along air flow is largely determined by their size. 39

Most communicable respiratory infections are transmitted via large droplets within short distance or by contacting contaminated surfaces. 40,41

Large droplets (diameter > 60 μm) tend to quickly settle form the air, so the risk of pathogen transmission is limited to individuals in close proximity to the saliva droplet source. 39

Small droplets (diameter ≤ 60 μm) may get involved in short-range transmission (distance between individuals less than 1 m).

Small droplets are likely to evaporate into droplet nuclei (diameter < 10 μm) in favorable environment, then become potential for long-distance aerosol transmission. 42

Generation of saliva droplets by a person

Saliva droplets are generated when breathing, talking, coughing, or sneezing and formed as particles in a mixture of moisture and droplet nuclei of microorganisms. 43

The amount, distance, and size of saliva droplets varies among people, suggesting the infectious strength and transmission path of saliva droplets differ when same pathogen was contracted. 44

Three thousand saliva droplet nuclei could be generated by one cough, which nearly equals to the amount produced during a 5-min talk. 43

Around 40,000 saliva droplets reaching several meters in air can be generated by one sneeze. 43,45

One normal exhalation can generate saliva droplets reaching the distance of 1 m in air. 43

Large saliva droplets with more mass tends to fall ballistically to the ground and small saliva droplets travel like a cloud over longer distance by air flow. 39,43,45

Environment-dependent saliva aerosols transmission

Aerosols are suspension of particles in air, liquid, or solid, within size from 0.001 to above 100 μm.39 Infectious aerosols contain pathogens. 39

Long-distance aerosol transmission is determined by sufficiently small infectious droplets, being almost indefinitely airborne and transmitted at a long distance (distance between individuals more than 1 m). 39

Aerosol transmission is well accepted in infection of tuberculosis, measles, and chickenpox, and other infectious agents may behave as airborne transmission in a favorable environment or opportunistically, such as SARS-CoV, influenza virus, and adenovirus. 40,46

Opportunistically airborne transmission is a mode that infectious agents not only have transmission routines by contacting and droplets but also can reach distant susceptible hosts under restricted conditions by fine-particle aerosols in favorable environments. 40

It is possible when aerosol-generating procedure is implemented, such as dental practice, that 2019-nCoV could possibly spread in airborne transmission. 38,47

Whether saliva droplets can become truly long-distance aerosol transmission is determined by how long the saliva droplets can reside in the air (physical decay), how long the pathogen in saliva droplets remain infectious (biological decay), and whether theses infectious saliva droplets can be acquired by another person (acquisition). 40

In terms of physical decay, saliva droplets evaporate fast into reduced mass in dry air, tending to stay longer along with air flow. 39

The composition of droplet nuclei determines its terminal size. 39

For droplets with slow biological decay, temperature differences and opened door set up droplets exchange along with air flow. 48

Biological decay is determined by dehydration, exposure to ultraviolet and chemicals. 39

Only hardy organism such as M. tuberculosis can survive long in air to form long-distance transmission. 49

A recent review summarized that coronavirus stay vital on surfaces of metal, glass, or plastic for up to 9 days, but no solid evidence has been found how long in air. 50

The coronavirus on inanimate surfaces could be efficiently inactivated by 0.1% sodium hypochlorite, 62–71% ethanol, or 0.5% hydrogen peroxide within 1 min as summarized in the literature. 50

So far, no solid evidence to consistently support that 2019-nCoV in saliva droplets can keep vital along air flow for very long time. Liu et al. collected 35 aerosol samples from three areas in two hospitals of Wuhan, and tested 2019-nCoV RNA by droplet digital polymerase chain reaction. 51

They found that patient area had low or even undetectable aerosol 2019-nCoV RNA, but deposition aerosol were tested positive, suggesting that not much vital virus in air flow but tend to deposit to the floor, which is similar to movements of large saliva droplets as noted previously. 51

In medical staff area, airborne 2019-nCoV RNA concentration was decreased after patients reduced and sanitization rigorously implemented. 51

In public area, accumulation of crowds increased airborne 2019-nCoV RNA concentration from undateable level.51

Acquisition of infectious saliva aerosols

For acquisition of infectious saliva droplets by a susceptible host, infectious saliva droplets could land in month, eyes, or be inhaled into lungs directly. 26,52

A case report shows that 2019-nCoV infection occurred in a fever clinic when a susceptible person wore an N95 mask covering mouth and nose without eyes protected, suggesting a transmission to eyes.53

It is also reported that SARS-CoV is predominantly transmitted by contacting eye, mouth, or nose.54

Respiratory virus could lead to respiratory infections of another person through inducing ocular complications.55

Exposed mucous membranes increased risk of virus transmission by a SARS-CoV study, and close exposure to an infected person increases the chance of infection.55

A previous study confirmed that infection of SARS-CoV was reduced to a certain degree by wearing surgical masks of susceptible healthcare workers.56

Comparison of saliva 2019-nCoV and SARS-CoV

2019-nCoV, which is also named as SARS-CoV-2,57 shares about 79% nucleotide sequence similarity with SARS-CoV.5,17,58–61 SARS-CoV have a higher mortality rate, while 2019-nCoV spreads much faster.12 The similarities and differences of saliva are summarized as follows in terms of diagnosis value of saliva, direct invasion to oral tissues, and saliva droplet transmission between SARS-CoV and 2019-nCoV, hopefully explaining the faster transmission speed of 2019-nCoV (Table

(Table1).

Table 1

Comparison of 2019-nCoV and SARS-CoV in terms of saliva

For the diagnostic value of saliva in coronavirus infection, high expression level of SARS-CoV RNA was detected in saliva specimens from 17 SARS patients, four of which had not yet lung lesion, suggesting value of early diagnosis of saliva, similarly to 2019-nCoV.62

A previous animal study on early events of SARS-CoV infection showed that SARS-CoV was detected in oral swabs before blood test turned positive on second day after viral challenge through nasal cavity.30,63

The viral load profile in saliva of 2019-nCoV nearly peaks at the time of symptom onset, while SARS-CoV peaks at around 10 days after symptoms.64–67

The high viral load of 2019-nCoV suggests it can be transmitted even if symptom is mild or less obvious.

Older age was associated with higher SARS-CoV in saliva, and a high initial SARS-CoV load was associated with death.　68,69

2019-nCoV RNA could be detected in saliva for 20 days or even longer, and the prolonged detection of viral RNA also exist in SARS-CoV and MERS-CoV infection.64–67

Despite 2019-nCoV and SARS-CoV share ACE2 receptor on host cells, which are found in salivary gland and tongue tissues,23–25,29,53

2019-nCoV is likely more infectious than SARS-CoV possibly due to lower RBD-ACE2 binding free energy and more flexible RBD of 2019-nCoV than that of SARS-CoV. 70

Compared with SARS-CoV, a furin-like cleavage site is peculiar in the S protein of 2019-nCoV, which could theoretically be cleaved by furin expressed in tongue tissues.33,34

For saliva droplets as transmission routine in coronavirus infection, a retrospective cohort study of SARS-CoV transmission reported that students who were in the same cubicle with the SARS patient contracted SARS-CoV, telling us that proximity to SARS patients increases chances of SARS-CoV infection. 52

Virus-laden droplets were found in a study as a routine of transmission in SARS epidemics. 71,72

van Doremalen et al. evaluated the stability of 2019-nCoV and SARS-CoV in aerosils using a Bayesian regression model, and found that 2019-nCoV remained viable ins aerosols throughout 3-h experiment duration, which is similar to SARS-CoV. 73

SARS-CoV dissemination belongs to opportunistically airborne transmission. 40,74

2019-nCoV also possibly belong to the same transmission type, dissemination of virus occurs if individuals are exposed to a high concentration of infectious aerosols in comparatively sealed space for long time.75

Prospective

It seems that the diagnostic value of saliva depends on how saliva specimens are collected. Saliva from deep throat (91.67 and 86.96% corresponding to two studies), from oral cavity (50%), or from salivary glands (12.90%) indicates a diagnostic tendency of decreased positive rate of 2019-nCoV RNA among COVID-19 patients. 11–14

For clinical application in need of high positive rate of virus detection, saliva from deep throat has the highest positive rate, which may stand for early diagnosis of COVID-19. Saliva directly from saliva glands ducts is associated with severe COVID-19 and possibly could be a predictive and noninvasive test for severed patients.

Whether 2019-nCoV RNA in saliva equals to infectious saliva or a condition of shedding vital virus is still lacking evidence.

Even if diagnosis by saliva is noninvasive and less hazardous compared with throat swabs, comprehensive diagnosis should be supported by complete information of symptoms, epidemiological history, and analysis of multiple clinical examinations.

Besides lungs, oral tissue is possible to be directly invaded theoretically due to expression of ACE2 receptor and furin enzyme. 14,29,30,37

About half of the victims reported symptoms of dry mouth and amblygeustia. 14

These symptoms probably came from dysfunction of tongue expressing ACE2 and furin, and salivary gland expression ACE2.

However, there is no histopathological evidence to support the direct invasion of 2019-nCoV to oral tissue so far.

While it suggests that cells expressing ACE2 and furin have lower restriction for virus entry theoretically, the molecular mechanism of 2019-nCoV infection is not yet fully unfolded and we should still be cautious about and not overstating the current virus-invade-host theory.

Saliva is a common and transient medium for virus transmission.

Among saliva droplets with different sizes generated by breathing, talking, and sneezing, large droplets easily fall onto the floor and only set up short-distance transmission.39

Saliva could form aerosols and reach a distant host along air flow when in a favorable environment. 39

So far, no solid evidence supports that SARS-nCoV or 2019-nCoV can survive in air outdoors for long time to set up long-distance aerosol transmission.

Therefore, wearing masks to prevent formation of infectious saliva droplets projecting to the air, thorough disinfection of indoor air to block dissemination of infectious saliva droplets, and keep a distance with people not to acquire infectious saliva droplets could slow down 2019-nCoV epidemic to a certain degree (Fig.1).

References

1. 1. Wang C, Horby PW, Hayden FG, Gao GF. A novel coronavirus outbreak of global health concern. Lancet. 2020;395:470–473. doi: 10.1016/S0140-6736(20)30185-9. - DOI - PMC - PubMed

   2. Zhu N, et al. A novel coronavirus from patients with pneumonia in China, 2019. N. Engl. J. Med. 2020;382:727–733. doi: 10.1056/NEJMoa2001017. - DOI - PMC - PubMed

   3. Cui J, Li F, Shi ZL. Origin and evolution of pathogenic coronaviruses. Nat. Rev. Microbiol. 2019;17:181–192. doi: 10.1038/s41579-018-0118-9. - DOI - PMC - PubMed

   4. Wong G, et al. MERS, SARS, and Ebola: the role of super-spreaders in infectious disease. Cell Host Microbe. 2015;18:398–401. doi: 10.1016/j.chom.2015.09.013. - DOI - PMC - PubMed

   5. Bai Y., Nie X. & Wen C. Epidemic Prediction of 2019-nCoV in Hubei Province and Comparison With SARS in Guangdong Province. https://ssrn.com/abstract=3531427 (2020).

   6. Spielmann N, Wong DT. Saliva: diagnostics and therapeutic perspectives. Oral. Dis. 2011;17:345–354. doi: 10.1111/j.1601-0825.2010.01773.x. - DOI - PMC - PubMed

   7. Navazesh M, Kumar SK, University of Southern California School of Dentistry. Measuring salivary flow: challenges and opportunities. J. Am. Dent. Assoc. 2008;139(Suppl):35S–40S. doi: 10.14219/jada.archive.2008.0353. - DOI - PubMed

   8. Kaufman E, Lamster IB. The diagnostic applications of saliva-a review. Crit. Rev. Oral. Biol. Med. 2002;13:197–212. doi: 10.1177/154411130201300209. - DOI - PubMed

   9. Zhang CZ, et al. Saliva in the diagnosis of diseases. Int J. Oral. Sci. 2016;8:133–137. doi: 10.1038/ijos.2016.38. - DOI - PMC - PubMed

   10. General Office of the National Health Commission, General Office of the Nation Administration of Traditional Chinese Medicine. Notice on issuing the pneumonia diagnosis and treatment plan for the new coronavirus infection (The 7th Trial Edition). (2020).

   11. To, K. K. et al. Consistent detection of 2019 novel coronavirus in saliva. Clin. Infect. Dis.10.1093/cid/ciaa149 (2020). - PMC - PubMed

   12. To, K. K.-W. et al. Temporal profiles of viral load in posterior oropharyngeal saliva samples and serum antibody responses during infection by SARS-CoV-2: an observational cohort study. Lancet Infect. Dis.10.1016/s1473-3099(20)30196-1 (2020). - PMC - PubMed

   13. Zhang W, et al. Molecular and serological investigation of 2019-nCoV infected patients: implication of multiple shedding routes. Emerg. Microbes Infect. 2020;9:386–389. doi: 10.1080/22221751.2020.1729071. - DOI - PMC - PubMed

   14. Chen, L. et al. Detection of 2019-nCoV in Saliva and Characterization of Oral Symptoms in COVID-19 Patients. https://ssrn.com/abstract=3557140 (2020).

   15. Mostafa, A., Abdelwhab, E. M., Mettenleiter, T. C. & Pleschka, S. Zoonotic potential of influenza A viruses: a comprehensive overview. Viruses10, (2018). 10.3390/v10090497. - PMC - PubMed

   16. Maginnis MS. Virus-receptor interactions: the key to cellular invasion. J. Mol. Biol. 2018;430:2590–2611. doi: 10.1016/j.jmb.2018.06.024. - DOI - PMC - PubMed

   17. Zhou P, et al. A pneumonia outbreak associated with a new coronavirus of probable bat origin. Nature. 2020;579:270–273. doi: 10.1038/s41586-020-2012-7. - DOI - PMC - PubMed

   18. Li W, et al. Angiotensin-converting enzyme 2 is a functional receptor for the SARS coronavirus. Nature. 2003;426:450–454. doi: 10.1038/nature02145. - DOI - PMC - PubMed

   19. Hofmann H, et al. Human coronavirus NL63 employs the severe acute respiratory syndrome coronavirus receptor for cellular entry. Proc. Natl Acad. Sci. USA. 2005;102:7988–7993. doi: 10.1073/pnas.0409465102. - DOI - PMC - PubMed

   20. Xu X, et al. Evolution of the novel coronavirus from the ongoing Wuhan outbreak and modeling of its spike protein for risk of human transmission. Sci. China Life Sci. 2020;63:457–460. doi: 10.1007/s11427-020-1637-5. - DOI - PMC - PubMed

   21. Wrapp D, et al. Cryo-EM structure of the 2019-nCoV spike in the prefusion conformation. Science. 2020;367:1260–1263. doi: 10.1126/science.abb2507. - DOI - PMC - PubMed

   22. Zhang, H. et al. The Digestive System is a Potential Route of 2019-nCov Infection: A Bioinformatics Analysis Based on Single-cell Transcriptomes. 10.1101/2020.01.30.927806 (2020).

   23. Zhao, Y. et al. Single-Cell RNA Expression Profiling of ACE2, the Putative Receptor of Wuhan 2019-nCov. 10.1101/2020.01.26.919985 (2020).

   24. Zou, X. et al. Single-cell RNA-seq data analysis on the receptor ACE2 expression reveals the potential risk of different human organs vulnerable to 2019-nCoV infection. Front. Med. 10.1007/s11684-020-0754-0 (2020). - PubMed

   25. Chai, X. et al. Specific ACE2 Expression in Cholangiocytes may Cause Liver Damage After 2019-nCoV Infection. 10.1101/2020.02.03.931766 (2020).

   26. Chan JF, et al. A familial cluster of pneumonia associated with the 2019 novel coronavirus indicating person-to-person transmission: a study of a family cluster. Lancet. 2020;395:514–523. doi: 10.1016/S0140-6736(20)30154-9. - DOI - PMC - PubMed

   27. Holshue ML, et al. First case of 2019 novel coronavirus in the United States. N. Engl. J. Med. 2020;382:929–936. doi: 10.1056/NEJMoa2001191. - DOI - PMC - PubMed

   28. Liang, W. et al. Diarrhoea may be underestimated: a missing link in 2019 novel coronavirus. Gut. 10.1136/gutjnl-2020-320832 (2020). - PubMed

   29. Xu H, et al. High expression of ACE2 receptor of 2019-nCoV on the epithelial cells of oral mucosa. Int J. Oral. Sci. 2020;12:8. doi: 10.1038/s41368-020-0074-x. - DOI - PMC - PubMed

   30. Liu L, et al. Epithelial cells lining salivary gland ducts are early target cells of severe acute respiratory syndrome coronavirus infection in the upper respiratory tracts of rhesus macaques. J. Virol. 2011;85:4025–4030. doi: 10.1128/JVI.02292-10. - DOI - PMC - PubMed

   31. Kessler AT, Bhatt AA. Review of the major and minor salivary glands, part 1: anatomy, infectious, and inflammatory processes. J. Clin. Imaging Sci. 2018;8:47. doi: 10.4103/jcis.JCIS_45_18. - DOI - PMC - PubMed

   32. Izaguirre, G. The proteolytic regulation of virus cell entry by furin and other proprotein convertases. Viruses11, 10.3390/v11090837 (2019). - PMC - PubMed

   33. Coutard B, et al. The spike glycoprotein of the new coronavirus 2019-nCoV contains a furin-like cleavage site absent in CoV of the same clade. Antivir. Res. 2020;176:104742. doi: 10.1016/j.antiviral.2020.104742. - DOI - PMC - PubMed

   34. Li, X. D. et al. A Furin Cleavage Site was Discovered in the S Protein of the Wuhan 2019 Novel Coronavirus. www.chinaxiv.org/abs/202002.00004 (2020).

   35. Bassi DE, Zhang J, Renner C, Klein-Szanto AJ. Targeting proprotein convertases in furin-rich lung cancer cells results in decreased in vitro and in vivo growth. Mol. Carcinog. 2017;56:1182–1188. doi: 10.1002/mc.22550. - DOI - PMC - PubMed

   36. Mallapaty S. Why does the coronavirus spread so easily between people? Nature. 2020;579:183. doi: 10.1038/d41586-020-00660-x. - DOI - PubMed

   37. Lopez de Cicco R, Watson JC, Bassi DE, Litwin S, Klein-Szanto AJ. Simultaneous expression of furin and vascular endothelial growth factor in human oral tongue squamous cell carcinoma progression. Clin. Cancer Res. 2004;10:4480–4488. doi: 10.1158/1078-0432.CCR-03-0670. - DOI - PubMed

   38. WHO. The First Few X Cases and Contact Investigation Protocol for 2019-Novel Coronavirus Infection. https://www.who.com.au/2019-nCoV/FFXprotocol/2020 (2020).

   39. Tang JW, Li Y, Eames I, Chan PK, Ridgway GL. Factors involved in the aerosol transmission of infection and control of ventilation in healthcare premises. J. Hosp. Infect. 2006;64:100–114. doi: 10.1016/j.jhin.2006.05.022. - DOI - PMC - PubMed

   40. Roy CJ, Milton DK. Airborne transmission of communicable infection-the elusive pathway. N. Engl. J. Med. 2004;350:1710–1712. doi: 10.1056/NEJMp048051. - DOI - PubMed

   41. Chapin CV. The importance of contact infection. Am. J. Public Hyg. 1910;20:742–759. - PMC - PubMed

   42. Fennelly KP, et al. Cough-generated aerosols of Mycobacterium tuberculosis: a new method to study infectiousness. Am. J. Respir. Crit. Care Med. 2004;169:604–609. doi: 10.1164/rccm.200308-1101OC. - DOI - PubMed

   43. Kohn WG, et al. Guidelines for infection control in dental health-care settings-2003. MMWR Recomm. Rep. 2003;52:1–61. - PubMed

   44. Edwards DA, et al. Inhaling to mitigate exhaled bioaerosols. Proc. Natl Acad. Sci. USA. 2004;101:17383–17388. doi: 10.1073/pnas.0408159101. - DOI - PMC - PubMed

   45. Cole EC, Cook CE. Characterization of infectious aerosols in health care facilities: an aid to effective engineering controls and preventive strategies. Am. J. Infect. Control. 1998;26:453–464. doi: 10.1016/S0196-6553(98)70046-X. - DOI - PMC - PubMed

   46. Sehulster L, Chinn RY. Guidelines for environmental infection control in health-care facilities. Recommendations of CDC and the Healthcare Infection Control Practices Advisory Committee (HICPAC) MMWR Recomm. Rep. 2003;52:1–42. - PubMed

   47. Peng X, et al. Transmission routes of 2019-nCoV and controls in dental practice. Int J. Oral Sci. 2020;12:9. doi: 10.1038/s41368-020-0075-9. - DOI - PMC - PubMed

   48. Tang JW, et al. Door-opening motion can potentially lead to a transient breakdown in negative-pressure isolation conditions: the importance of vorticity and buoyancy airflows. J. Hosp. Infect. 2005;61:283–286. doi: 10.1016/j.jhin.2005.05.017. - DOI - PMC - PubMed

   49. Fitzgerald, D. W., Sterling, T. R. & Haas, D. W. Mycobacterium tuberculosis. In: Mandell, G. L., Bennett, J. E. & Dolin, R. (Eds). Mandell, Douglas, and Bennett’s principles and practice of infectious diseases. Seventh edition (Philadelphia, PA, Churchill Livingstone Elsevier, 2010).

   50. Kampf G, Todt D, Pfaender S, Steinmann E. Persistence of coronaviruses on inanimate surfaces and their inactivation with biocidal agents. J. Hosp. Infect. 2020;104:246–251. doi: 10.1016/j.jhin.2020.01.022. - DOI - PMC - PubMed

   51. Liu, Y. et al. Aerodynamic Characteristics and RNA Concentration of SARS-CoV-2 Aerosol in Wuhan Hospitals during COVID-19 Outbreak. 10.1101/2020.03.08.982637 (2020).

   52. Wong TW, et al. Cluster of SARS among medical students exposed to single patient, Hong Kong. Emerg. Infect. Dis. 2004;10:269–276. doi: 10.3201/eid1002.030452. - DOI - PMC - PubMed

   53. Lu CW, Liu XF, Jia ZF. 2019-nCoV transmission through the ocular surface must not be ignored. Lancet. 2020;395:e39. doi: 10.1016/S0140-6736(20)30313-5. - DOI - PMC - PubMed

   54. Peiris JS, Yuen KY, Osterhaus AD, Stohr K. The severe acute respiratory syndrome. N. Engl. J. Med. 2003;349:2431–2441. doi: 10.1056/NEJMra032498. - DOI - PubMed

   55. Belser JA, Rota PA, Tumpey TM. Ocular tropism of respiratory viruses. Microbiol Mol. Biol. Rev. 2013;77:144–156. doi: 10.1128/MMBR.00058-12. - DOI - PMC - PubMed

   56. Seto WH, et al. Effectiveness of precautions against droplets and contact in prevention of nosocomial transmission of severe acute respiratory syndrome (SARS) Lancet. 2003;361:1519–1520. doi: 10.1016/S0140-6736(03)13168-6. - DOI - PMC - PubMed

   57. Gorbalenya, A. E. et al. Severe acute respiratory syndrome-related coronavirus: the species and its viruses—a statement of the Coronavirus Study Group. Preprint at https://www.biorxiv.org/content/, 10.1101/2020.02.07.937862v1 (2020).

   58. Chen N, et al. Epidemiological and clinical characteristics of 99 cases of 2019 novel coronavirus pneumonia in Wuhan, China: a descriptive study. Lancet. 2020;395:507–513. doi: 10.1016/S0140-6736(20)30211-7. - DOI - PMC - PubMed

   59. Li, Q. et al. Early transmission dynamics in Wuhan, China, of novel coronavirus-infected pneumonia. N. Engl. J. Med. 10.1056/NEJMoa2001316 (2020). - PMC - PubMed

   60. de Wit E, van Doremalen N, Falzarano D, Munster VJ. SARS and MERS: recent insights into emerging coronaviruses. Nat. Rev. Microbiol. 2016;14:523–534. doi: 10.1038/nrmicro.2016.81. - DOI - PMC - PubMed

   61. Al-Tawfiq JA, Zumla A, Memish ZA. Coronaviruses: severe acute respiratory syndrome coronavirus and Middle East respiratory syndrome coronavirus in travelers. Curr. Opin. Infect. Dis. 2014;27:411–417. doi: 10.1097/QCO.0000000000000089. - DOI - PubMed

   62. Wang WK, et al. Detection of SARS-associated coronavirus in throat wash and saliva in early diagnosis. Emerg. Infect. Dis. 2004;10:1213–1219. doi: 10.3201/eid1007.031113. - DOI - PMC - PubMed

   63. Qin C, et al. An animal model of SARS produced by infection of Macaca mulatta with SARS coronavirus. J. Pathol. 2005;206:251–259. doi: 10.1002/path.1769. - DOI - PMC - PubMed

   64. Perlman S, Netland J. Coronaviruses post-SARS: update on replication and pathogenesis. Nat. Rev. Microbiol. 2009;7:439–450. doi: 10.1038/nrmicro2147. - DOI - PMC - PubMed

   65. Huang C, et al. Clinical features of patients infected with 2019 novel coronavirus in Wuhan, China. Lancet. 2020;395:497–506. doi: 10.1016/S0140-6736(20)30183-5. - DOI - PMC - PubMed

   66. Peiris JS, et al. Clinical progression and viral load in a community outbreak of coronavirus-associated SARS pneumonia: a prospective study. Lancet. 2003;361:1767–1772. doi: 10.1016/S0140-6736(03)13412-5. - DOI - PMC - PubMed

   67. Oh MD, et al. Viral load kinetics of MERS coronavirus infection. N. Engl. J. Med. 2016;375:1303–1305. doi: 10.1056/NEJMc1511695. - DOI - PubMed

   68. Chu CM, et al. Initial viral load and the outcomes of SARS. CMAJ. 2004;171:1349–1352. doi: 10.1503/cmaj.1040398. - DOI - PMC - PubMed

   69. Chen WJ, et al. Nasopharyngeal shedding of severe acute respiratory syndrome-associated coronavirus is associated with genetic polymorphisms. Clin. Infect. Dis. 2006;42:1561–1569. doi: 10.1086/503843. - DOI - PMC - PubMed

   70. He, J., Tao, H., Yan, Y., Huang, S.-Y. & Xiao, Y. Molecular Mechanism of Evolution and Human Infection With the Novel Coronavirus (2019-nCoV). 10.1101/2020.02.17.952903 (2020).

   71. Yu IT, et al. Evidence of airborne transmission of the severe acute respiratory syndrome virus. N. Engl. J. Med. 2004;350:1731–1739. doi: 10.1056/NEJMoa032867. - DOI - PubMed

   72. Li Y, Duan S, Yu IT, Wong TW. Multi-zone modeling of probable SARS virus transmission by airflow between flats in Block E, Amoy Gardens. Indoor Air. 2005;15:96–111. doi: 10.1111/j.1600-0668.2004.00318.x. - DOI - PubMed

   73. van Doremalen, N. et al. Aerosol and surface stability of SARS-CoV-2 as compared with SARS-CoV-1. N. Engl. J. Med. 10.1056/NEJMc2004973 (2020). - PMC - PubMed

   74. WHO. Infection Prevention and Control of Epidemic- and Pandemic-prone Acute Respiratory Infections in Health Care (World Health Organization, 2014).

   75. Su J. [Aerosol transmission risk and comprehensive prevention and control strategy in dental treatment] Zhonghua Kou Qiang Yi Xue Za Zhi. 2020;55:E006. - PubMed