SARS-CoV-2 air and surface contamination on a COVID-19 ward and at home

SARS-CoV-2 has been detected both in air and on surfaces, but questions remain about the patient-specic and environmental factors affecting virus transmission to the environment. Additionally, more detailed information on viral ndings in air is needed. This cross-sectional study presents results from 259 air and 252 surface samples from the surroundings of 23 hospitalized and eight home-treated COVID-19 patients between July 2020 and March 2021 and compares the results between the measured environments and patient factors. The proportions of PCR-positive air and surface samples showed statistical similarity in hospital and in the home. In four cases, positive environmental samples were detected even after the patients had developed a neutralizing IgG response. SARS-CoV-2 RNA was detected in the following particle sizes: 0.65–4.7 µm, >7 µm, >10 µm, and <100 µm. Appropriate infection control against airborne and surface transmission routes is needed in both environments, even after antibody production has begun.


Introduction
Increasing scienti c evidence indicates the dominance of short-and long-term airborne transmission of SARS-CoV-2 1-6 , and discussion on precautions for hospital and home environments has been intense. In a study that aerosolized SARS-CoV-2 under laboratory conditions, aerosols' infectivity retained for up to 16 h 7 , while another study estimated the halflife in aerosols to be approximately 1.1 to 1.2 hours (95% CI 0.64 to 2.64) 8 . Outside of the laboratory, signs of viable SARS-CoV-2 in the air have been detected, and virus was recently also cultured from exhaled air [9][10][11] . In hospitals, PCRbased studies have detected SARS-CoV-2 RNA in room air [12][13][14] , as well as from air conditioning lters located over 50 m from the patient room 15 . However, these studies used long collection times or high ow rates, generating large sample volumes mainly from small spaces, and thus questions remain about the risk of infection during shorter meetings or in rooms with a larger air space, and whether the ndings would be similar in the home environment.
According to laboratory ndings, the stability of SARS-CoV-2 on surfaces varies depending on the surface type and environmental conditions 8,16−19 . However, its ability to sustain infectivity on surfaces in real environments is largely unknown 20 . SARS-CoV-2 RNA has been found, for example, on high-touch surfaces, oors, and toilets 13,14,21 , but no study has yet been able to culture SARS-CoV-2 from surfaces in real environments. The effect of age and neutralizing antibodies (NAbs) on the spread of SARS-CoV-2 has been speculated [22][23][24] , but there is a lack of clear evidence for the role of patient-related factors.
This study sought to increase knowledge of SARS-CoV-2 transmission in different environments by analyzing air, surface, and patient samples from a COVID-19 cohort ward in Helsinki University Hospital (HUS), Finland, and from patients' homes. The aims were to determine whether SARS-CoV-2 RNA or viable virus could be found in the home and hospital environments, and which patient-and environment-related factors affect the risk of environmental contamination. A team consisting of researchers from HUS, the University of Helsinki, the Finnish Meteorological Institute, and the Finnish Institute of Occupational Health was established to enable a multidisciplinary approach to the above research questions.

Patient characteristics
We performed 23 sample collections in HUS and 7 collections in patients' homes in the Uusimaa region, Finland, between July 2020 and March 2021. Collections included 31 index patients (1-2 per collection), 21 of whom were treated on a COVID-19 cohort ward in a large patient hall, one in a single-patient room, one in the intensive care unit (ICU), and eight patients treated in their homes (Fig. 1). Patient characteristics, including symptoms and laboratory results, are summarized in Table 1 (see Tables S1 and S2 for details of the patients and statistical tests used throughout the manuscript). Table 1 Characteristics of hospital-treated and home-treated index patients and statistical differences between the two groups.
Hospital (n = 23) Home a (n = 8) Total (N = 31) p Gender (% of males) 56. 5 (Table S4, results from home and hospital collections have been combined, as there was no statistically signi cant difference between the positivity rates for the collections).
SARS-CoV-2 RNA was found from particles in the size ranges 0.65-4.7 µm and >7 µm in Andersen collectors, >10 µm and <2.5 µm in Dekati samplers and <100µm in Button samplers (Tables 2 and S3). On-line particle concentrations measured with the eFilter on the COVID-19 ward were in the range of 534-6608 cm -3 (3380 ± 2320 cm -3 ), and no clear particle emission events were observed. During each Andersen collection, three Andersen samplers were used simultaneously.
The Button sampler is limited to particles smaller than 100 µm, but does not differentiate the sizes inside this range.
Similarly, the largest particle stages in Dekati (>10 µm) and Andersen samplers (>7 µm) do not limit the upper size range.  (Table 3). There was no statistically signi cant difference in the proportions of PCR-positive samples (p = 0.333) or collections (p = 1.000) between home and hospital. Only one sample, A77 (  (Table S3).

Effects of patient factors on environmental contamination
Positive air samples were found even when the index patient did not report any respiratory symptoms (2/3, 66.6%).
However, there was a statistically signi cant connection between low oxygen saturation (SpO2) levels and SARS-CoV-2 RNA ndings from surfaces, and a possible but nonsigni cant connection between low SpO2 levels and RNA ndings from the air (surface: p = 0.026, air: p = 0.098, Table S2). Toilet surfaces were PCR positive in 33.3% (3/9) of cases when the index patient had GI symptoms and 0% of cases (0/9) when the index patient did not report any GI symptoms (p =  (Table S6).
RNA copy numbers in saliva samples varied between 1.65 x 10 3 and 5.13 x 10 7 copies/ml (mean 3.55 x 10 6 copies/ml (SD 1.10 x 10 7 )). Age showed a trend of positive correlation with copy number, but it was not statistically signi cant

SARS-CoV-2 antibodies in serum samples
Serum samples were obtained from 21 hospital-treated patients (13 index patients and six other patients on the ward) and four home-treated patients (two index patients and two other patients). In total, ten serum samples were positive for IgG or NAbs. Antibodies were detected at the earliest on symptom day 3 (P13, positive with two IgG tests, NAb titer 80).
Of the antibody-positive patients, 9/10 were PCR positive from saliva and one (P16, symptom day 11, NAb titer 80) was also positive in viral culture. The index patients had NAbs against SARS-CoV-2 in ve of the collections, and in four of these, PCR-positive environmental samples were detected (active air samples in one (P49, NAb titer >640), deposition air samples in two (P41 and P43, NAb titers 40 and 10), and surface samples in two (P13 and P43, NAb titers 80 and 10)).

Environmental contamination and virus strain
The virus variant was determined in seven index patients during 2021, ve of whom were infected with alpha variant, one with an undetermined variant of concern (VoC), and one with a non-VoC strain. The remaining cases were considered as non-VoC, as no VoC strains had yet been detected in Finland at the time of the collection. The mean RNA copy number in saliva was 1.
Our ndings support discoveries that normal respiratory activities generate infective particles in the absence of aerosolgenerating procedures 2,6,25 . Most (83%, 15/18) of our positive samples were in particles smaller than 4.7 µm, which strengthens the recent nding that at least 85% of the viral load is emitted in aerosols smaller than 5 µm 6 .
SARS-CoV-2 was detected from air with a minimum collection period of 10 min (Andersen's impactor) and a minimum air volume of 72 l (Button sampler). With an average respiratory rate of 14/min and volume of 0.5 l/breath, this would mean exposure times of 40 min (Andersen) and 10 min (Button) for the examined virus variants (alpha and undetermined VoC (Andersen), as well as non-VoC (Button)). However, our results from respiratory activities demonstrated that 0.5-2 min of activity did not produce enough virus to be detected with PCR, even from a close distance of 10 cm. Current safety guidelines use 15 min exposure time regarding contact tracing 26 . Our results raise concern that shorter exposure time should be considered, at least for close contacts. Also, current virus variants such as the delta variant may further lower the exposure time needed for infection, as the estimated viral load in the presence of the delta variant is over 1000 times higher than the initial strain 27 .
Multiple positive air samples were collected from a large (655.25 m 3 ) mechanically ventilated hospital hall (Figure 1), even when there were only two patients. Overall, larger spaces are considered safer than small ones due to the larger air volume per person 28 . However, it seems that also larger indoor spaces may form a risk environment if occupied by an infected person for a prolonged time period as observed also in previous studies [29][30][31] . Higher viral loads have been associated with an increased probability of viral transmission 32 . We also found a higher number of patients in the room to be associated with higher numbers of positive surface samples.
Toilet surface samples were positive only when the index patient reported GI symptoms. Infectious SARS-CoV-2 has been recovered from urine and stool samples 33 , and ushing of the toilet and vomiting can generate aerosols, which will later deposit on the surfaces 34,35 . This risk should be noted and toilets should not be shared with non-COVID patients.
Other more frequently PCR-positive surfaces included highly-touched personal items, hospital equipment, and the oor, which is in line with previous ndings 13,14,21 . Even though RNA may persist on surfaces for some time, RNA ndings most likely result from contamination on the same day due to daily cleaning.
The relative importance of different infection routes remains somewhat unclear, although a recent animal study indicates that aerosol inoculation is a more e cient route and causes more severe pathology and higher viral loads 36 .
The recent evidence estimates surface transmission to be likely rare, generally less than 1 in 10 000, and the disease manifestation milder 37 . In this study, families that took protective measures (including isolation of the infected family member) and respiratory protection (surgical masks or FFP2 respirators) were able to prevent further infections even when PCR-positive samples were collected from both surfaces and air. However, in a household where all surfaces were cleaned many times a day but no respiratory protection was used, all family members became infected. This supports the importance of air hygiene and also encourages control of infection spread in homes. Infection control is even more important with VoC strains that feature a higher rate of household transmission 38 .
To better understand the infectivity and state of the infection compared to the environmental ndings, we collected saliva and serum samples from studied patients. SARS-CoV-2 was cultured from patients' saliva during symptom days 2-11. In contradiction to previous ndings 39  The effect of age on the generation of aerosols and thus the spread of SARS-CoV-2 has also been speculated 45 . We observed a strong trend for an older age being associated with a higher viral load and a larger number of positive surface samples, but con rming this would require further studies with a larger sample size. Possible reasons for the relationship between age and infectivity include reduced saliva production, differences in mucus viscosity and salivary immunoglobulins 46 , increased expression of the ACE2 receptors needed for cell entry of SARS-CoV-2 47 , thinning of the epithelium 48 , and impairment of the immune response with age 49 .
This study combined a large number of environmental samples and detailed patient data to more comprehensively understand environmental contamination and the effect of patient-dependent factors. The patient material was representative regarding symptoms and laboratory results for COVID-19 50 . In addition to the hospital environment, we collected samples from homes where patient symptoms are generally less severe, the time from the onset of symptoms is shorter, and air conditioning is different from that of a hospital.
Our study also has some limitations. We only conducted environmental sampling at a single time point. In the future, a longitudinal examination could enable more accurate examination of the effects of the course of disease for environmental contamination. In addition, we only measured the IgG and NAb response, but viral secretion from mucus membranes can continue if the IgA response is weak 51 . The IgA immune response should thus be examined further in upcoming studies. The qPCR results might include some uncertainty due to the differences in the texture and uidity of saliva and should be considered as estimates. As many samples were collected from a large patient hall, it is possible that some observed viruses might have originated from other than the index patient. However, most of the surface samples were from patient-speci c surfaces, and aerosols are known to concentrate near the source 52 , indicating that most of the positive samples are expected to be produced by the index patient. Particle size cutoffs in Andersen samplers might be slightly higher than estimated, as the amount of liquid used in the sampling was slightly smaller than recommended due to practical reasons. As seen also in earlier studies 53 (Table S1). In one case, the index patient was the same in two collections, and in two other collections, two index patients were included. All research personnel conducting the sampling followed aerosol safety protocols and precautions and no infections were detected. All procedures that involved human participants, including

Cell lines
Vero E6 cells (VE6) and their TMPRSS2-expressing clone VE6-TMPRSS2-10 (VE6T) 58 were grown as previously described 59 . To inhibit fungal growth, 0.205 µg/ml of amphotericin B (Fungizone, Thermo Scienti c) was added to the medium of the cells that were taken to the hospital for aerosol collections.
Sampling protocols for air sampling Seven different air collection methods were used. Details of the collections and samples are presented in Table S3 To evaluate the real-time particle number concentration during the hospital collections and to gather additional air samples, a Dekati eFilter was used in two collections. The eFilter monitors changes in real-time particle concentration by utilizing a small diffusion charger powered by an inner chargeable battery. The charge changes were automatically translated into a signal, which was recorded on a data card. When postprocessing the data, the raw charge signal was further converted to represent particle number concentrations using a conversion factor (411 cm^-3 fA^-1) provided by the manufacturer. A count median diameter (CMD) of 60 nm and a geometric standard deviation (GSD) of 1.5 were assumed 61,62 . In addition, the eFilter simultaneously collected samples on a 47-mm gelatin lter using an external pump.
After sample collection, the gelatin lter was transferred into 6 ml of MEM. The eFilter was tted with the same EPAdesigned inlets as the Andersen cascade impactors. The particle size cut point of the inlets was approximately 12 µm, with an air volume ow rate of 28.3 l/min. The duration of sample collection was 30 minutes at a similar distance from the patient as with the Andersen's cascade impactors.
Passive air samples were collected either directly on VE6 cells (2 collections) or VE6T cells (9 collections) grown on 100/20 MM (collection 22) or 35/10 MM (other collections) cell culture dishes or on empty 35/10 MM Petri dishes containing 1 ml of growth medium (10 collections). Open dishes were positioned at different proximities from the patient for 30-60 min, and the patient was encouraged to perform an aerosol-producing activity such as talking. The ability of SARS-CoV-2 to infect cells at room temperature was con rmed, and major differences in the culture sensitivity of these two collection methods were excluded under laboratory conditions (see supplementary methods for details).
Living cells were transported to the laboratory in a warm environment with heat accumulators warmed to 37°C. One plate was used as a negative control to ensure that the cells survived the transport. Other samples were transported with cold accumulators and handled during the same or next day.

Sampling protocols for surface sampling
Altogether, 252 surface samples in 26 collections for PCR testing were taken from surfaces in possible direct or indirect contact with the patient (Supplementary Table 3 Nasopharyngeal samples from consenting patients were taken and sent to HUSLAB for a fresh diagnostic PCR 63,64 . Serum samples from consenting patients were taken within a day from sampling and tested for SARS-CoV-2 IgG antibodies with two different tests 65 . Serum samples (dilutions 1:10 to 1:640) were studied with the microneutralization assay 66 . Blood lymphocyte and eosinophil counts, and plasma CRP from consenting patients were measured within a day of sampling, and plasma ferritin, ALP, ALT, D-dimer, and brinogen levels within three days. The respiration rate and SpO2 levels were measured during the same day (Table S1).
Since the rst cases caused by variants of concern (VoC) were detected in Finland at the end of December 2020, they were determined from all patients as a part of routine diagnostics. This information was used to compare the results between VoC strains (mainly alpha in Finland) and non-VoC strains. Virus strains of collections 1-22 (P1-P45) were considered as non-VoC, as they were collected before the rst cases were reported in Finland.
RNA extraction and PCR protocols for air, saliva, and culture medium samples Trizol (Invitrogen) was used to extract RNA from all saliva samples and from air and culture medium samples of collections and 58°C (N Charité)). The primer and probe concentrations of N Charité were according to the original publication, and those of N1 US CDC were 500 nM of both primers and 125 nM of probe (Table S7) Finally, RNA was quanti ed by spectrophotometry and the RNA copy number was calculated based on its concentration, length, and molecular weight. qPCR was performed with N Charité PCR by including a dilution series from 10 to 10 9 copies/reaction in triplicate.   Table S2.

Supplementary Files
This is a list of supplementary les associated with this preprint. Click to download. 20211020SupplementarymaterialsforSARSCoV2airandsurfacecontaminationonaCOVID19wardandathome.docx