Virus shedding and environmental deposition of novel A(H1N1) pandemic influenza virus

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Chapter 1 Introduction

As pandemic mitigation strategies have been developed over recent years it has become very clear that influenza transmission is one area that is poorly understood and hotly debated. Distinguishing the relative importance of the various modes of transmission (Box 1) is critical for the development of infection control precautions in health-care settings and in the home.

If contact transmission is dominant then hand hygiene becomes the most critical intervention. However, if respiratory droplet transmission is significant, surgical face masks that provide a barrier against droplets may be important, and the safe distance away from an infected person without a mask might be as close as 4 feet (ft), because droplets fall out of the air quickly and do not travel far. At present, opinions are sharply divided on the importance of bioaerosol transmission. 1,2 Tellier1 in particular, argues that the potential of short-range bioaerosol transmission has largely been ignored. At present, the UK recommends droplet precautions as opposed to bioaerosol precautions (surgical masks rather than respirators) for most forms of contact with patients with pandemic influenza,3 based on the current balance of limited evidence; however, this is contested by some frontline health-care workers who believe that these safeguards are inadequate, and there is little evidence with which to reassure them.

In parallel, the dynamics of viral shedding in relation to symptom onset and severity are important factors, highly relevant to estimates of the period of infectivity and to therapeutic management. In all previous research on influenza virus excretion, shedding has been determined by measurement of the quantity of virus recoverable from the patient’s nasopharynx, i.e. virus has been recovered by a deliberately performed invasive technique. These so called ‘viral shedding’ studies measure virus shed from infected cells; they do not actually measure virus that is deposited into the touched or respired environment; i.e. they do not define environmental contamination and the hazard posed to others. While such data are useful, if they could be linked to near-patient environmental sampling, estimates of the extent to which infectious virus is deposited on to surfaces and into the air in the patient’s immediate vicinity could be made.

Chapter 2 Methods

This multicentre, prospective, observational descriptive cohort study recruited subjects between 14 September 2009 and 25 January 2010, in accordance with the principles of the Declaration of Helsinki and UK regulatory requirements. It was approved by Leicestershire, Northamptonshire & Rutland Research Ethics Committee 1 (09/H0406/94).

Chapter 3 Results

One hundred and fifty subjects were screened between 14 September 2009 and 25 January 2010; 107 were ineligible, and 43 were enrolled and followed up. Reasons for exclusion at screening included: symptoms being present for too long (48%), influenza PCR test (as part of medical care)-negative (15%), declined to take part (9%). Pandemic H1N1 virus was detected in 19 subjects. The group of 24 pandemic-negative cases consisted of: RSV = 5 (all children); rhinovirus = 5; corona virus = 2; rhinovirus + corona virus = 1; NHS-pandemic H1N1 test-positive, study laboratory pandemic H1N1 test-negative = 2; unknown = 9. In the final analyses, one subject was excluded on the basis of having received pandemic H1N1 vaccine prior to enrolment, and three subjects were removed (all of whom tested negative for pandemic H1N1 according to the study laboratory); two because clinical (as part of medical care) and study pandemic H1N1 2009 PCR tests did not agree; and one because study documents were lost. Recruitment by group of the 39 remaining subjects was as follows: 9 AC, 12 AH, 15 CC and 3 CH (Figure 1).

FIGURE 1.

Participant flow diagram.

Of the remaining 39 subjects, 19 (49%) tested positive for pandemic H1N1 virus and 20 (51%) were negative. Follow-up of at least 8 days occurred in 16/19 positives and 12/20 negatives. The numbers enrolled, along with a demographic description of pandemic H1N1 cases, is shown in Table 1.

Symptoms

The most frequently reported symptoms in our subjects with pandemic H1N1 were: stuffy nose (100%), runny nose (100%), cough (100%), fatigue (95%) and sneezing (89%) (Table 3). Fever was reported on the day illness began in 13/19 (68%) cases, and was measured as high (≥ 38°C) during follow-up in 7/19 (37%) of cases.

TABLE 3 - Symptoms reported over the course of study follow-up in both patients with pandemic H1N1 influenza and others

The symptom of fever was not recorded on a daily basis, although an oral measurement of body temperature was.

	No. of patients, n (%)
	Pandemic H1N1 subjects (n = 19)	Others (n = 20)
Fever (on day of onset)a	13 (68)	12 (57)
Runny nose	19 (100)	20 (95)
Sore throat	12 (63)	17 (81)
Cough	19 (100)	21 (100)
Shortness of breath	14 (74)	20 (95)
Stuffy nose	19 (100)	18 (86)
Sneezing	17 (89)	17 (81)
Earache	3 (16)	8 (38)
Sinus tenderness	12 (63)	15 (71)
Diarrhoea	6 (32)	8 (38)
Vomiting	10 (53)	10 (48)
Fatigue	18 (95)	20 (95)
Headache	15 (79)	12 (57)
Myalgia	14 (74)	15 (71)

In general, symptom scores declined over time. URT and systemic symptoms peaked on day 2 of illness and LRT symptoms peaked on day 3. However, it should be noted that most subjects were recruited > 36 hours after illness onset, which may give misleading information on maximal symptom scores; there was only one patient with information available on day 1, and only five for day 2 (Figure 2a). Figure 2b shows mean symptom scores of subjects with pandemic H1N1 influenza as a function of the number of days since illness onset.

FIGURE 2.

Mean symptom scores of pandemic H1N1 cases over time. (a) Number of observations (subject data) available for each day. Day 1 is the day of symptom onset. (b) Mean symptom scores of subjects with pandemic H1N1 as a function of the number of days since symptoms started.

In a comparison of subjects who were positive for pandemic H1N1 infection with others recruited, no significant difference was seen in the average time from symptom onset to recruitment: positive cases (1.7 days), others (1.7 days) (p = 0.90). Visual inspection of plots showing mean symptom scores (broken down into categories) over time suggests that subjects who were negative for pandemic H1N1 infection had higher URT symptom scores and LRT symptoms that peaked 3 days after pandemic H1N1-positive subjects (Figure 3). However, no significant differences between these two groups were detected when comparing symptoms scores on the day of recruitment (URT p = 0.11, LRT p = 0.18 or systemic symptoms p = 0.20) or in the total mean symptom score over time (46.5 for subjects with pandemic H1N1 vs 52.3 for others, p = 0.54).

FIGURE 3.

Upper respiratory tract, lower respiratory tract and systemic symptom scores over time. First row: mean symptom scores for positive pandemic H1N1 cases (solid line) and negative cases (dashed line) as a function of the number of days since symptoms started. Second row: number of observations available for each day; pandemic H1N1 cases (solid line) and negative cases (dashed line). Day 1 is the day of symptom onset.

Antiviral drugs

Overall, 21/39 (54%) of enrolled subjects took an antiviral drug [either oseltamivir (20/21) or zanamivir (1/21)] and this occurred within 2 days of illness onset in 12/17 cases (71%) for which data are available. Of the pandemic H1N1-positive cases, 11/19 (58%) received an antiviral drug (all oseltamivir); hospital cases 7/8 (88%) and community cases 4/11 (36%). A total of 44% of pandemic H1N1 cases took oseltamivir within 48 hours, and the average time from symptom onset to treatment initiation in these subjects was 1.7 days (data on when treatment was begun for one patient is not available). The mean total symptom score on the first day of enrolment in the study was significantly higher for subjects with pandemic H1N1 who received antiviral drugs within 48 hours of symptom onset (mean score 16.6) than subjects with pandemic H1N1 who either did not take oseltamivir or did so after 48 hours of illness onset (8.6) (p = 0.018) (Figure 4a).

FIGURE 4.

Symptom scores over time for pandemic H1N1 cases who took antiviral drugs within 48 hours and those who did not. (a) Mean total symptom score as a function of the number of days since symptom onset for pandemic H1N1 subjects who received antivirals within 48 hours of symptoms onset (solid line) and those with pandemic H1N1 who did not (dashed line). (b) Number of observations available for each day.

Viral load

Subject viral loads were examined over time and in relation to symptom scores. Nasal swab viral loads, measured by PCR, varied widely across our pandemic H1N1-positive subjects, ranging from 0.9 × 10¹ to 1.7 × 10¹¹ copies/ml. Viral loads plotted over time are shown for four subjects from whom the most complete data were obtained (Figure 5a). All subject viral loads over time are shown in Figure 5b, which illustrates the heterogeneity of the data; for each individual trajectory, viral loads tend to decrease with time, but there is an apparent increase in the mean value, because individuals with high viral loads tend to shed for longer.

FIGURE 5.

Viral loads plotted over time. (a) Viral loads from selected subjects are shown. (b) All viral load data are shown. Subject trajectories are shown as dashed lines and the mean of these is shown as a solid line. (c) Number of observations available for each day. Day 1 is the day of symptom onset.

The mean peak viral loads of the four recruitment groups were 5.9 × 10⁵ for AH, 2.4 × 10⁵ for AC, 1.0 × 10⁷ for CH and 1.6 × 10⁶ for CC. No significant differences were detected between any of the groups, although there was a trend towards higher peak loads in children (Figure 6). The mean peak viral load of adults was 4.4 × 10⁵, and that of children was 2.2 × 10⁶, with no significant difference detected between them (p = 0.28).

FIGURE 6.

Peak viral loads of subjects categorised into their recruitment groups.

Neither total, URT or systemic symptom scores correlated with viral loads at different points in time. However, the LRT symptoms score on day 5 was significantly correlated (p = 0.049) (Figure 7).

FIGURE 7.

Scatter plots showing symptom scores and viral loads over time. (a) and (b) Total symptom score versus viral load as a function of the number of days since illness onset. (c) and (d) LRT score versus viral load as a function of the number of days since symptom onset.

Virus shedding

The duration of virus shedding measured by PCR had mean of 6.2 days and a range of 3–10 days. There was no difference between children (mean 6.1 days) and adults (mean 6.3 days) (p = 0.89). Based on the numbers involved, the power to detect a difference was 19% if adult shedding was 6 days and child shedding was 7 days. The duration of shedding of hospital cases (mean 6.8 days) was slightly longer than that of community cases (mean 5.7 days), although the difference was not significant (p = 0.33) (Figure 8).

FIGURE 8.

Distribution of the duration of virus shedding by PCR. The duration of viral shedding is defined as the time between symptom onset and the last day that a positive specimen was taken. Because patients were seldom recruited on the day symptoms began, an assumption has been made that they were shedding virus from the first day of symptoms to the last positive specimen.

No substantial correlation between the duration of shedding and symptom score on the day of recruitment was detected, with coefficients of correlation with URT symptoms of 6% (p = 0.8), with LRT symptoms 19% (p = 0.43) and with systemic symptoms 8% (p = 0.75).

A total of 12/19 cases (63%) were culture positive for pandemic H1N1. The mean duration of live virus shedding from these 12 cases was 4.7 days (range 3–8 days). However, because cases with no positive culture were excluded (durations too short to be observed or false-negative testing), this represents an upper bound for the duration of shedding. To obtain a lower bound for the duration, the calculation was repeated with the assumption that ‘negative’ patients do not shed live virus (duration of shedding = 0). This gives a mean duration of 2.9 days (range 0–8). The median value when all 19 subjects were included was 3 days, and 6/19 (31%) subjects shed live virus for at least 5 days from the onset of illness.

Figure 9 shows the distribution of live virus shedding for the 12 positive cases, and highlights the recruitment group to which each subject belongs. There was no significant correlation between the duration of the live virus shedding and total symptom score of these 12 cases on the day of recruitment [correlation coefficient –0.09, 95% confidence interval (CI) –0.63 to 0.51, p = 0.78] or the sum of total symptom scores during the whole follow-up (correlation coefficient –0.22, 95% CI –0.71 to 0.40, p = 0.48).

FIGURE 9.

Distribution of the duration of virus shedding by culture positivity (n = 12). The duration of viral shedding is defined as the time between symptom onset and the last day that a positive culture was obtained. Cultures were performed from the last day of nasal swab PCR positivity. If a culture was positive on any given day then it was assumed that previous days’ would also have been culture-positive.

The mean duration of shedding determined by both PCR and culture was not significantly different for subjects who received antivirals within 48 hours and those who received them after 48 hours or not at all [PCR: 6.4 days vs 5.9 days, p = 0.61; culture-positives: 4.6 days vs 4.8 days, p = 0.88). All culture results (assuming six have 0 days): 3.4 days vs 2.4 days, p = 0.43].

Box 2 summarises symptom and virus shedding findings.

BOX 2 - Symptom and virus shedding data summary

• Symptoms decline over time

• Initial symptom scores were similar in subjects positive or negative for pandemic H1N1 influenza

• Subjects with pandemic H1N1 had fewer URT symptoms and an earlier peak in LRT symptoms than other subjects

• Viral load was highly variable between subjects; children had higher peak viral loads than adults but this difference was not statistically significant

• No clear relationship was evident between symptom scores and viral load

• No clear distinction was shown in the duration of virus shedding between adults and children

• Mean duration of PCR-detectable virus shedding was 6.2 days (maximum was 10 days)

• Median duration of viable virus shedding was 3 days (maximum was 8 days)

• Total duration of virus shedding detectable by PCR or culture was unrelated to initial symptom severity

• No obvious relationship between shedding of viable virus and any particular symptom(s) was identified

Environmental deposition

Surfaces

In total, 414 community swabs (+ 52 sponges) and 45 hospital swabs (+ seven sponges) were taken, of which 397 swabs and 12 sponges were tested (not all swabs were tested because of sample processing rules, see Chapter 2, Laboratory methods). Pandemic H1N1 virus was detected by PCR on two occasions on surfaces from around one patient in the community (following discharge from hospital), giving a swab positivity rate of 0.5%. Quantitative PCR could only be performed on one sample because the amount of sample available in the other was insufficient. Live virus was recovered from one of these surfaces. The subject from around whom the swabs were taken was found to be shedding virus from the nose on the same day, although other household members were also unwell on these days; a 5-year-old was unwell with cough and fever on day 4, and a 2-year-old was unwell with cough and fever on day 10 (Table 4).

TABLE 4 - Details of surface swabs that were positive for pandemic H1N1 virus

N/A, not available.

Note, at the time swabs were taken other household members in this subject’s family were also unwell with symptoms of ILI.

	Specimen no.
	1	2
Subject ID	AH04	AH04
Surface (setting)	Kettle handle (home)	Bathroom tap (home)
Surface material	Plastic	Metal
Swab method	Cotton swab	Cotton swab
Number of days after symptoms began that swab was taken	4	10
Viral load from surface swab (copies/ml)	91,205	N/A
Viral load from nose on day swab collected (copies/ml)	902,703	N/A
Culture	Positive	Negative

Air

Air samples were collected from the immediate environment of five subjects (all of whom were rapid antigen test positive): three while in hospital and two in the community. Seventeen separate collections were undertaken, generating 51 samples (although one could not be processed because of insufficient sample volume). Air samples were positive from four out of five subjects. Eight out of 17 (47%) collections and 22/50 (44%) samples were positive for PCR. No samples were confirmed to contain live virus (Table 5).

TABLE 5 - Description of air particle samples collected

N/A, not available.

	Subject
	AH03				AH04			CC05				CC15				CH03
Subject setting (+ infected others)	Hospital bed in side room				Hospital bed in side room			Playing in bedroom				Playing in living room (6-year-old infected child also present)				Cot on neonatal unit (two infected neonates also present)
Room temperature (°C)	21.6				23.3			20.0				18.0				24.0
Room humidity (relative %)	50				50			64				60				40
No. of days after symptoms began that sample was taken	4				3			3				3				5
Viral load from nose on day sample collected (copies/ml)	238,091				10,625,714			699,723				178,923,317,453				24,208
Virus cultured in the nose on sampling day?	Yes				Yes			Yes				No				Yes
Duration of sampling (hours)	1	1	3	3	1	1	2	1	1	3	3	1	1	3	3	3	3
Approximate distance from subject (ft)	3	7	3	7	3	7	7	3	7	3	7	3	7	3	7	3	7
Pandemic H1N1 virus detected by PCR	+	–	+	–	+	–	–	–	–	–	–	+	+	+	+	+	+
Particle size virus detected in µm	< 1	–	< 1	–	–	–	–	–	–	–	–	< 1	< 1	N/A	< 1	< 1	< 1
	–	–	1–4	–	1–4	–	–	–	–	–	–	1–4	1–4	1–4	1–4	1–4	1–4
	–	–	> 4	–	–	–	–	–	–	–	–	> 4	> 4	> 4	> 4	> 4	> 4

Quantitative PCR demonstrated a range of values between 238 and 24,231 copies/ml; higher values were recorded in instances when more than one infected person was present in the sampling room. Samples collected over a 1-hour period generated 8/24 PCR-positives (33%), those over a 2-hour period zero out of three positives, and those over a 3-hour period 14/23 positives (61%). The risk ratio for a sample to be positive over a 3-hour period relative to a 1-hour period was 1.83 (95% CI 0.95 to 3.51, p = 0.082). Samples collected at a distance close to the subject (approximately 3 ft) generated 13/23 PCR-positives (57%), whereas those collected further away (at least 7 ft) generated 9/27 PCR-positives (33%). The risk ratio for a sample to be PCR positive at a distance of 3 ft versus ≥ 7 ft was 1.70 (95% CI 0.89 to 3.22, p = 0.15). Virus was detected in all particle sizes collected: particles < 1 µm gave 7/16 positives (44%); particles 1–4 µm gave 8/17 positives (47%) and particles > 4 µm gave 7/17 positives (41%). Among particles of size 1–4 µm and > 4 µm, the relative risk of obtaining a positive sample relative to particles of size < 1 µm was 1.08 (95% CI 0.51 to 2.28, p > 0.99) and 0.94 (95% CI 0.43 to 2.08, p > 0.99), respectively (Table 5).

Initially it appeared that 3 samples were culture positive for virus. To verify that the cultured virus in the air samples was the same as that from subject’s nose, PCR was carried out on the harvested virus to confirm the presence of pandemic H1N1. However, as well as the clear presence of pandemic H1N1 there was a signal that indicated the presence of another virus. Work was then undertaken to try and identify this virus though it is important to note the following: (1) there were no original samples left to reanalyse; (2) the signal was detected only in harvested, amplified virus; and (3) this signal was not seen in the air sample on which the initial PCR was done.

• PCR assays were performed (see Appendix 6, PCR protocol), which confirmed the contaminating virus to be influenza A, H1. Plaque assay on the harvested air sample virus was strongly positive (titre 30 × 10⁷ × 2.5/ml = 7.5 × 10⁸ plaque-forming units (pfu)/ml). (Note: pandemic H1N1 does not plaque in these cells.)

• Contamination with another influenza virus did not preclude there being live pandemic H1N1 virus in the cells as well. Therefore, an experiment was performed whereby diluted virus was cultured and an attempt made to quantify the amount of virus by PCR. If live pandemic H1N1 was present in the original sample, we postulated that extracted nucleic acid should be at higher concentration in the re-amplified aliquots. Harvested virus was diluted in 10-fold steps from neat to 10^–7. Each dilution was split into two aliquots: one frozen and the other inoculated into fresh MDCK (Madin–Darby Canine Kidney) cells. The MDCK cells were incubated for 48 hours before the virus was again harvested. Results indicate that there was no live pandemic H1N1 virus in these samples (at least by these methods). Three out of 11 dilutions were positive for pandemic H1N1 influenza prior to reamplification, but none of the dilutions was positive post re-amplification.

• Finally, in an attempt to determine conclusively the identity of the contaminating virus, samples of the matrix gene amplicons were sequenced. Results show that influenza A PR8 was the contaminating isolate (undoubtedly from the laboratory).

Findings from the environmental sampling are summarised in Box 3.

BOX 3 - Environmental sampling data summary

• Almost no fomite contamination was found (0.5% of all specimens taken)

• Five subjects had samples of the air around them taken and virus was detected by PCR from four of them; PCR positive specimens were equally well represented across all of the particle size ranges measured

• Although viable virus was recovered from three samples, we were unable to prove that this virus was pandemic H1N1, as opposed to a contaminant

Composite charts

In order to best demonstrate the information we have generated for each subject, charts integrating data from nasal swabs and environmental samples are shown below for selected patients (Figure 10). All patient charts are shown in Appendix 7.

FIGURE 10.

Composite charts for subjects. (a) AH04; (b) AH03; (c) CC15; (d) CH01. The ‘symptom’ bar shows the number of days for which symptoms were present. The ‘nasal swab’ bar shows the last day that a swab was either PCR-positive or culture-positive. The ‘surface’ and ‘air’ bars show days up to the time that a positive sample was obtained. Arrows show the days when oseltamivir was started and when follow-up ended. ‘Day 1’ is the day of illness onset.

Chapter 4 Discussion

This is the first study that has attempted to assess actual viral shedding from patients with influenza, by examining the near-patient environment for virus as opposed to simply taking respiratory specimens. Sampling virus, particularly live virus, in the environment is challenging; getting to the subject in time, executing optimal sampling while preserving virus viability and performing sensitive detection tests in the laboratory are all key factors that necessitate very extensive and complex logistic arrangements. An attempt to overcome this first problem was carried out by targeting recruitment in the community, as well as in hospital (when presentation is often delayed), enabling an approach to subjects early in their illness when virus shedding is usually at its highest. In addition, the use of a bioaerosol sampler, designed and validated by collaborators at NIOSH enabled us to sample air around infected subjects.

Subjects’ with pandemic H1N1 experienced a range of symptoms, but a mild illness was evident in the majority of cases, as has been reported elsewhere. 32 There were no significant differences with respect to symptom type or duration between those positive for pandemic H1N1 virus and those who were non-confirmed (negative). Although the non-confirmed cases included some individuals who were infected with other respiratory viruses, undoubtedly some were falsely negative on pandemic H1N1 virus testing.

Viral loads, in general, declined over time, although a lack of data hinders further interpretation. Only 5/19 subjects had data available at four or more time points. The wide range of results seen may in part be reflected by differences in sample quality. The peak viral load was found to be higher in children than adults, in line with other studies,12,15 although this was not significant. There was a significant association, however, found between viral load and LRT symptoms on day 5 of a subject’s illness, suggesting that persistent LRT symptoms might be a clinical marker for prolonged shedding. However, cautious interpretation of this result is necessary, given the lack of data.

Our findings on virus shedding, as conventionally described, are broadly in agreement with other published findings relating to pandemic H1N1 virus (Table 5). The median duration of virus shedding from the 19 infected cases was 6 days when detection was performed by PCR, and 3 days when detection was performed by a culture technique. Forty-four per cent of these subjects received oseltamivir within 2 days of illness onset. Fifty-eight per cent of subjects were recruited directly from the community, and these cases shed virus for a shorter period of time than the hospital cases (5.7 vs 6.8 days). Although this finding was not significant it accords, nevertheless, with data suggesting that hospitalised influenza cases shed virus for longer,9,10 with potential infection control implications for health-care institutions.

When comparing studies (Table 6), it should be borne in mind that differences in study populations may exist (children vs adults, hospital vs community cases), a variety of sampling methods are used and that the proportions of cases receiving antiviral drugs (particularly whether they received them within 48 hours) may differ. In a Vietnamese hospitalised cohort of 292 pandemic H1N1 cases, PCR detected virus in combined nose and throat swabs in the following proportion of patients: after 1 day of treatment 86% (165/192); day 2 59% (45/76); day 3 38% (27/72); day 4 25% (34/138); and day 5 14% (11/76). After 5 days of treatment, 7% (12/179) were still positive, although no positive cultures were obtained after day 5. 17 Laboratory findings from a study of 70 cases in Singapore gave a mean duration of viral shedding of 6 days, with shedding > 7 days in 37% of patients. The mean duration of positive culture results on six patients was 4 days. 13 Finally, in a Canadian study, 43 community patients with pandemic H1N1 had a nasopharyngeal specimen collected on day 8 of their illness: 74% were PCR-positive and 19% were culture-positive. 33

One subject from our study who demonstrated the shedding of live virus up to day 8 will be considered further. She was a 34-year-old woman, of South-Asian origin, who had no comorbidities, and did not take regular medicine. She spent one night in hospital on the first day of her illness and began taking oseltamivir on day 2 (the subject reported taking oseltamivir each day and, while there is no reason to suspect non-compliance, this cannot be excluded). Prominent symptoms early in her illness were fever, cough, sore throat and fatigue. The virus was sequenced across the HA gene during the period of time that it was shed, and no changes were detected. In addition, no common oseltamivir resistance mutations were detected. All of the other household family members subsequently developed symptoms of cough and fever; a 5-year-old daughter became unwell on day 4 of the mother’s illness, followed by a 2-year-old son on day 5 and her 30-year-old husband on day 6. Thus, a high secondary attack rate in this family was associated with high levels and prolonged shedding of virus, despite the index case being treated with oseltamivir.

It is interesting to note that no difference was found in the duration of viral shedding (PCR or culture) between those who took oseltamivir within 48 hours and those who did not, although our numbers are small (10 vs 8), and it is impossible to draw conclusions because a sample size of at least several hundred subjects would have been needed. Other studies have demonstrated a shortened duration or suppressed levels of shedding in association with oseltamivir when it is given early. 13,34,35 Subjects with pandemic H1N1 who did receive antiviral drugs had significantly higher initial symptom scores than those who did not, perhaps indicating that patients with more severe symptoms were more likely to access to early treatment. This difference might mask any effect of antiviral drugs on duration of shedding. In addition, it may explain why symptom scores were consistently lower among those who received no or late treatment than among those who received early treatment.

Our findings relating to the duration of live virus shedding have infection control implications. They suggest that over 30% of cases remain potentially infectious for at least 5 days and, given that live virus may persist in the environment for up to 48 hours,19 viable virus may be present for 7 days after an index case first develops symptoms. These data are consistent with other recent studies that suggest that pandemic H1N1 may be contagious for a longer period of time than seasonal flu. 13,33 This has clear implications for pandemic infection control and self-isolation guidelines.

However, despite finding that live virus shedding continued for over 4 days in most subjects, fomites contaminated with virus were found in only two instances, involving only one subject. Therefore, only 0.5% of all community fomites, and none of the hospital fomites, swabbed revealed virus, although on one occasion live virus was found. This instance occurred in a household where, at the time of taking the surface swab, a 5-year-old child was also experiencing her first day of symptoms, but the surface contamination was from a kettle handle and so is unlikely to have been directly handled by the secondary case. These findings are in contrast with those of Boone and Gerba,21 who detected influenza virus (by PCR) on over 50% of all swabs taken from a number of fomites in the home and in child-care centres. They also differ from the findings of a study that involved subjects who were experimentally infected with influenza virus. Swabs taken from fomites in subjects’ rooms (two subjects shared a room) revealed influenza (detected by PCR) in 9/48 swabs (19%), although no live virus was found (B Killingley, University of Nottingham, May 2010, personal communication). It is also likely that more than one individual contributed to virus deposition in Boone and Gerba’s study. 21 This contrasts with the circumstances of the current study, where only one individual was ill when the vast majority of swabs were taken. In addition, the homes used in Boone and Gerba’s study21 contained a symptomatic child 100% of the time compared with 79% of homes in the current study. It is also worth noting that no specific cleaning instructions were given during the follow-up of our subjects, so, for example, daily cleaning of hospital rooms would have continued, which may have contributed to the low positive swab rate. A more speculative suggestion would be that pandemic H1N1 is less stable in the environment than other influenza strains, and indeed there is some evidence to suggest that some influenza viruses may be more robust than others. In experimental conditions an avian virus survived for up to 6 days on some surfaces36 and unpublished observations (J Greatorex, HPA, May 2010, personal communication) suggest a laboratory-adapted PR8 (H1N1) virus is more hardy than seasonal wild-type strains. The finding of influenza RNA on fomites on its own does not prove that disease can be spread via the contact route – demonstration of live virus transmitted in an infectious dose would be required for this. Despite an isolated discovery of live virus, our findings overall suggest that the contact route of transmission for pandemic H1N1 may well play a more minor role in the transmission of influenza than hitherto suggested by experts, and by the current emphasis placed on hand hygiene as a means of interrupting transmission.

A noteworthy finding of this study is the demonstration of virus in particles collected from the air around subjects who have influenza; this has not previously been attempted in a community setting. Five subjects had samples of the air around them taken, and virus was detected by PCR from four of them. In two instances there were additional patients with pandemic H1N1 (children) present in the room as well as the study subject during air sampling, and it was these samples that revealed the most virus. All particle sizes collected contained virus detectable by PCR, including the < 1-µm and 1–4-µm fraction sizes, which are bioaerosols of a respirable size, i.e. they can reach the distal airways of the respiratory tract. 37 Sampling for a longer time period, and nearer to the subject, led to the detection of more virus as one might expect, although analyses did not reveal any statistical significance because numbers were small.

Unfortunately, we have been unable to conclusively demonstrate the presence of live pandemic H1N1 in any samples. Initial culture results indicated the presence of live virus in three samples from one subject (AH03) and PCR detected only pandemic H1N1 in the original samples. However, following amplification of the virus to permit further analysis, it appears that the sample became contaminated with a laboratory influenza strain. It was not possible to go back to the original sample (as none remained) or subsequently prove that the live virus detected was pandemic H1N1 as opposed to the contaminant.

There were no unusual room temperature or humidity readings recorded during sampling, but there are insufficient data to study the effects of these variables further.

It is unclear why it was not possible to culture live virus from specimens when most subjects had live virus detected on nasal swabs, although detecting live virus in samples is challenging and the techniques are still relatively new. Difficulties include the fragility of the virus particle (especially its susceptibility to desiccation) and the fact that sufficient virus needs to be collected to enable culture. Because the amount and concentration of virus being sampled in air is much lower than that from nasal swabs, detection is more difficult. The use of VTM during sample collection (as opposed to its addition afterwards) to help preserve virus has been cited as a necessity by some,38 and with sound reason. But, as has demonstrated in other unpublished laboratory work (B Killingley, University of Nottingham, May 2010, personal communication), this does not appear to be an absolute requirement with the samplers used.

Evidence backing up at least the potential for bioaerosol transmission of influenza infection has recently been reviewed;39 supporting evidence comes from the detection of influenza virus (by PCR) in the air around patients,26,27 the demonstration of bioaerosol transmission in animal models,40,41 and increasingly sophisticated mathematical modelling techniques, which suggest a role for bioaerosol spread. 42 Detecting the presence of influenza in the air is the first step in a chain of evidence needed to confirm that influenza viruses – emitted from an infected individual and existing as bioaerosols – can initiate infection in a person exposed to them. The other steps in this sequence are (1) confirming that live, i.e. infectious, virus is present and (2) confirming that sufficient live virus exists that can be inhaled by an individual to initiate infection. Couch et al. 43 conducted a series of experiments in 1966, culminating in a human-to-human transmission study attempting to follow this line of evidence for coxsackie virus, and came to the conclusion that bioaerosol transmission ‘unquestionably occurred’. Similar data on influenza are lacking and it remains that the human infectious dose of influenza in natural conditions is not known for any route. Alford et al. 25 showed that three times the TCID50 was needed to infect volunteers via bioaerosols; this compares to other studies showing that 127–320 TCID50 are needed to initiate infection by the intranasal route. 44 Using these data, attempts have been made to estimate the risk of infection attributable to the different routes of infection,45 but the outputs of such models are only ever as good as the input assumptions. However, if Alford et al. ’s25 supposition is true then even small quantities of viable virus expressed via bioaerosols might have significant infectious potential.

Detection of virus by PCR was seen from air samples collected at close range (3 ft) to subjects, well within the contact distance of an attending health-care worker suggesting that the theory of short range bioaerosol transmission advanced by Tellier39 cannot be dismissed. Although clearly based on extremely limited data, these finding are of sufficient importance to justify further efforts to reproduce them including further attempts to detect of live virus.

There are several limitations to this study. First, the numbers of subjects recruited was well below target. The study began recruiting just prior to the beginning of the second wave of the pandemic in England, but the overall number of people infected during the second wave was well below what had been predicted46 and seroconversions during the first wave were far higher than expected. 47 In addition a mild illness, including a high asymptomatic infection rate47 contributed to our difficulty. It is also evident that enrolling people early in the course of their illness is challenging. Over one-half of the volunteers we saw were ineligible because symptoms had been present for too long. A further problem was difficulty in identifying subjects as having influenza as opposed to other ARIs. It has been shown that the standard definition of ILI cannot be relied upon to distinguish pandemic H1N1 from other ARIs,48,49 and the low numbers of people with illness in the local population made the positive predictive value of even our modified definition of ILI low (48%). A near-patient rapid antigen test was used to help reveal influenza cases, but our original inclusion criteria that required a positive antigen test were modified because the sensitivity of the test in our hands (with a nasal swab) was low. Overall, 10/19 (53%) of our cases were antigen test-positive; the sensitivity in adults was 25% and in children 73%. These findings concur with a number of other reports about the low sensitivity of these tests to detect pandemic H1N1. 50–52 This resulted in a difficulty in reliably recruiting only subjects with pandemic H1N1, such that we followed up subjects who had other ARI. For technical and logistic reasons, the capacity to generate PCR results on samples quickly enough to limit this follow-up in most cases did not exist. The modest recruitment of pandemic H1N1 cases limits the study in several ways, including the generalisability of our findings and because of a lack of data the ability to address our primary aim – to correlate virus shedding on nose swabs with environmental samples.

Second, the sampling methods used require further consideration, as care is needed during interpretation of the results:

• Nasal swab Although a nasopharyngeal aspirate (NPA) is considered to be the best specimen for detecting influenza A viruses,53–55 this procedure causes more discomfort and is more difficult to perform, particularly in children. Indeed, studies attempting to collect daily NPA samples from subjects have reported problems with subjects’ tolerance and compliance with the procedure. 15 A nasal swab, however, has been shown to be an acceptable alternative that is not statistically less sensitive than a NPA,54–56 although suboptimal sampling (caused by interoperator variation in technique) can still occur.

• Fomite swabbing Despite adopting a similar swabbing technique to other comparable studies,22,23 and validating this in advance using experimentally deposited virus (B Killingley, University of Nottingham, May 2010, personal communication), virus was rarely isolated from fomites. Furthermore, the fomites sampled were similar, except that four of our nine chosen surfaces (bedside table, dining table, patient table and window sill) are not items that are actually picked up or grasped by the hand. Virus may well be transferred to, or settle on, such surfaces, but sampling was performed from only a small proportion of the surface area. Furthermore, many of these surfaces were made of wood, a material that does not support virus survival (J Greatorex, HPA, May 2010, personal communication). In future, consideration will be given to alternative sampling methods, for example using a sponge (wiping a surface may collect more material and can cover a larger surface area) and increasing our focus on ‘grasped’ items. We used and tested the sponge too infrequently during this study to draw any firm conclusions about its performance compared with a cotton swab.

Finally, all subjects from whom air samples were obtained tested positively on rapid antigen testing. This may have biased the group somewhat, as a positive rapid antigen test has been associated with higher viral loads in nasal samples. 50 On the other hand, our intention was to prove whether viable virus deposition on surfaces or in the air was possible in practice; so selection of these individuals was important. Also no measurements or estimates of room air flow patterns or ventilation were made when collecting samples. Such parameters are likely to have an influence on the ability to detect virus in the air.

Chapter 5 Conclusion

Despite limitations resulting in an inability to fully address the primary aims of the study, important observations have been made. Our findings show that live pandemic H1N1 virus can be found in the noses of over 30% of infected individuals for at least 5 days after symptoms begin. The evidence for the significance of both contact and bioaerosol routes of transmission, depends upon demonstrating that viable virus is deposited from an infected patient. This has been shown for touched surfaces, although the data suggest that contact transmission via fomites may be less important than hitherto emphasised. Transmission via bioaerosols at short range is not ruled out; virus was detected by PCR in aerosols, but we were unable to conclusively demonstrate the presence of live virus.

Acknowledgements

We extend our thanks to the individuals (and parents) who took part in this study and are grateful for the clinical and administrative assistance of the Department of Health (DH), the National Institute of Health Research, the Leicestershire, Northamptonshire & Rutland Research Ethics Committee 1, Division of Epidemiology and Public Health at Nottingham University, Trent Comprehensive Local Research network, Trent Local Children’s Research Network, East Midlands Primary Care Research Network, Nottingham University Hospitals NHS Trust, Nottingham City Primary Care Trust, Nottingham County PCT, Leicester University Hospitals NHS Trust, Sheffield Teaching Hospitals NHS Trust and to the HPA at Addenbrooke’s Hospital and the Health and Safety Laboratory for laboratory support. Particular thanks are extended to Sheila O’Malley, Penny Scardifield, Maria Benitez, Raquel Velos, Joanna Tyrrell, Pradhib Venkatesan, Harsha Varsani, Fayna Garcia, Paul Digard and Catherine Makison.

We are grateful to William Lindsley and Donald Beezhold at NIOSH for the loan of the air samplers and to the Centers for Disease Control and Prevention, Atlanta, for facilitating this.

BK is funded by a Medical Research Council clinical research training fellowship.

References

1. Tellier R. Review of aerosol transmission of influenza A virus. Emerg Infect Dis 2006;12:1657-62.
2. Brankston G, Gitterman L, Hirji Z, Lemieux C, Gardam M. Transmission of influenza A in human beings. Lancet Infect Dis 2007;7:257-65.
3. Department of Health . Influenza: A Summary of Guidance for Infection Control in Healthcare Settings 2009. www.dh.gov.uk/prod_consum_dh/groups/dh_digitalassets/@dh/@en/@ps/documents/digitalasset/dh_110899.pdf.
4. Boivin G, Goyette N, Hardy I, Aoki F, Wagner A, Trottier S. Rapid antiviral effect of inhaled zanamivir in the treatment of naturally occurring influenza in otherwise healthy adults. J Infect Dis 2000;181:1471-4.
5. Lau LL, Cowling BJ, Fang VJ, Chan KH, Lau EH, Lipsitch M, et al. Viral shedding and clinical illness in naturally acquired influenza virus infections. J Infect Dis 2010;201:1509-16.
6. Ng S, Cowling BJ, Fang VJ, Chan KH, Ip DK, Cheng CK, et al. Effects of oseltamivir treatment on duration of clinical illness and viral shedding and household transmission of influenza virus. Clin Infect Dis 2010;50:707-14.
7. Treanor JJ, Hayden FG, Vrooman PS, Barbarash R, Bettis R, Riff D, et al. Efficacy and safety of the oral neuraminidase inhibitor oseltamivir in treating acute influenza: a randomized controlled trial. US Oral Neuraminidase Study Group. JAMA 2000;283:1016-24.
8. Frank AL, Taber LH, Wells CR, Wells JM, Glezen WP, Paredes A. Patterns of shedding of myxoviruses and paramyxoviruses in children. J Infect Dis 1981;144:433-41.
9. Lee N, Chan PK, Hui DS, Rainer TH, Wong E, Choi KW, et al. Viral loads and duration of viral shedding in adult patients hospitalized with influenza. J Infect Dis 2009;200:492-500.
10. Leekha S, Zitterkopf NL, Espy MJ, Smith TF, Thompson RL, Sampathkumar P. Duration of influenza A virus shedding in hospitalized patients and implications for infection control. Infect Control Hosp Epidemiol 2007;28:1071-6.
11. Sato M, Hosoya M, Kato K, Suzuki H. Viral shedding in children with influenza virus infections treated with neuraminidase inhibitors. Pediatr Infect Dis J 2005;24:931-2.
12. Cao B, Li XW, Mao Y, Wang J, Lu HZ, Chen YS, et al. Clinical features of the initial cases of 2009 pandemic influenza A (H1N1) virus infection in China. N Engl J Med 2009;361:2507-17.
13. Ling LM, Chow AL, Lye DC, Tan AS, Krishnan P, Cui L, et al. Effects of early oseltamivir therapy on viral shedding in 2009 pandemic influenza A (H1N1) virus infection. Clin Infect Dis 2010;50:963-9.
14. Suess T, Buchholz U, Dupke S, Grunow R, Matthias An der H, Heider A, et al. Shedding and transmission of novel influenza virus A/H1N1 infection in households: Germany 2009. Am J Epidemiol 2010;71:1157-64.
15. To KK, Chan KH, Li IW, Tsang TY, Tse H, Chan JF, et al. Viral load in patients infected with pandemic H1N1 2009 influenza A virus. J Med Virol 2010;82:1-7.
16. van Doorn RH. Influenza Pandemic (H1N1) 2009 (68): Viet Nam, Virus Clearance 2009. www.promedmail.org/pls/apex/f?p=2400: 1001: 1739427750156980::NO:: F2400_P1001_BACK_PAGE,F2400_P1001_PUB_MAIL_ID: 1004,79587.
17. Reed SE. An investigation of the possible transmission of Rhinovirus colds through indirect contact. J Hyg 1975;75:249-58.
18. Hall CB, Douglas RG. Modes of transmission of respiratory syncytial virus. J Pediatr 1981;99:100-3.
19. Bean B, Moore BM, Sterner B, Peterson LR, Gerding DN, Balfour HH. Survival of influenza viruses on environmental surfaces. J Infect Dis 1982;146:47-51.
20. Thomas Y, Vogel G, Wunderli W, Suter P, Witschi M, Koch D, et al. Survival of influenza virus on banknotes. Appl Environ Microbiol 2008;74:3002-7.
21. Boone SA, Gerba CP. The occurrence of influenza A virus on household and day care center fomites. J Infect 2005;51:103-9.
22. Pappas DE, Hendley JO, Schwartz RH. Respiratory viral RNA on toys in pediatric office waiting rooms. Pediatr Infect Dis J 2010;29:102-4.
23. Loosli CG, Lemon HM, Robertson OH, Appel E. Experimental airborne influenza infection.I. Influence of humidity on survival of virus in air. Proc Soc Exp Biol Med 1943;53:205-6.
24. Wells WF, Brown HW. Recovery of influenza virus suspended in air. Science 1936;84:68-9.
25. Alford RH, Kasel JA, Gerone PJ, Knight V. Human influenza resulting from aerosol inhalation. Proc Soc Exp Biol Med 1966;122:800-4.
26. Blachere FM, Lindsley WG, Pearce TA, Anderson SE, Fisher M, Khakoo R, et al. Measurement of airborne influenza in a hospital emergency department. Clin Infect Dis 2009;48:438-40.
27. Lindsley WG, Blachere FM, Davis KA, Pearce TA, Fisher MA, Khakoo R, et al. Distribution of airborne influenza virus and respiratory syncytial virus in an urgent care medical clinic. Clin Infect Dis 2010;50:693-8.
28. McLean RL. The effect of ultraviolet radiation upon the transmission of epidemic influenza in long term hospitals. Am Rev Respir Dis 1961;83:36-8.
29. Moser MR, Bender TR, Margolis HS, Noble GR, Kendal AP, Ritter DG. An outbreak of influenza aboard a commercial airliner. Am J Epidemiol 1979;110:1-6.
30. Han K, Zhu X, He F, Liu L, Zhang L, Ma H, et al. Lack of airborne transmission during outbreak of pandemic (H1N1) 2009 among tour group members: China, June 2009. Emerg Infect Dis 2009;15:1578-81.
31. Nicas M, Nazaroff WW, Hubbard A. Toward understanding the risk of secondary airborne infection: emission of respirable pathogens. J Occup Environ Hyg 2005;2:143-54.
32. Writing Committee of the WHO Consultation on Clinical Aspects of Pandemic Influenza . Clinical Aspects of Pandemic 2009 Influenza A (H1N1) Virus Infection. N Engl J Med 2010;362:1708-19.
33. De Serres G, Rouleau I, Hamelin ME, Quach C, Skowronski D, Flamand L, et al. Contagious period for pandemic (H1N1) 2009. Emerg Infect Dis 2010;16:783-8.
34. Li IW, Hung IF, To KK, Chan KH, Wong SS, Chan JF, et al. The natural viral load profile of patients with pandemic 2009 influenza A(H1N1) and the effect of oseltamivir treatment. Chest 2010;137:759-68.
35. Schutten M, Lina CBB, Monto A, Osterhaus A, Nguyen Van-Tam J, Whitley RJ. Neuraminidase inhibitor (NAI) treatment in a 2009–H1N1 influenza A-infected patient population: The IRIS Study. Options for the Control of Influenza VII 2010.
36. Tiwari A, Patnayak DP, Chander Y, Parsad M, Goyal SM. Survival of two avian respiratory viruses on porous and nonporous surfaces. Avian Dis 2006;50:284-7.
37. Hinds WC. Aerosol technology. New York, NY: John Wiley and Sons; 1999.
38. Fabian P, McDevitt JJ, Houseman EA, Milton DK. Airborne influenza virus detection with four aerosol samplers using molecular and infectivity assays: considerations for a new infectious virus aerosol sampler. Indoor Air n.d.:433-41.
39. Tellier R. Aerosol transmission of influenza A virus: a review of new studies. J R Soc Interface 2009;6:S783-90.
40. Mubareka S, Lowen AC, Steel J, Coates AL, Garcia-Sastre A, Palese P. Transmission of influenza virus via aerosols and fomites in the guinea pig model. J Infect Dis 2009;199:858-65.
41. Munster VJ, de Wit E, van den Brand JM, Herfst S, Schrauwen EJ, Bestebroer TM, et al. Pathogenesis and transmission of swine-origin 2009 A (H1N1) influenza virus in ferrets. Science 2009;325:481-3.
42. Nicas M, Sun G. An integrated model of infection risk in a health-care environment. Risk Anal 2006;26:1085-96.
43. Couch RB, Cate TR, Douglas RG, Jr, Gerone PJ, Knight V. Effect of route of inoculation on experimental respiratory viral disease in volunteers and evidence for airborne transmission. Bacteriol Rev 1966;30:517-29.
44. Douglas R, Kilbourne ED. The influenza viruses and influenza. New York, NY: Academic Press; 1975.
45. Nicas M, Jones RM. Relative contributions of four exposure pathways to influenza infection risk. Risk Anal 2009;29:1292-303.
46. Colthart DG. Pandemic Influenza and Swine Flu 2009. www.parliament.uk/briefingpapers/commons/lib/research/briefings/snsc-05059.pdf.
47. Miller E, Hoschler K, Hardelid P, Stanford E, Andrews N, Zambon M. Incidence of 2009 pandemic influenza A H1N1 infection in England: a cross-sectional serological study. Lancet 2010;375:1100-8.
48. Chan M, Chen MI, Chow A, Lee CP, Tan AS, Lye DC, et al. Pandemic (H1N1) 2009: clinical and laboratory findings of the first fifty cases in Singapore. Ann Acad Med Singapore 2010;39:267-72.
49. Nguyen-Van-Tam J, Openshaw P, Hasim A, Gadd E, WS L, Semple M, et al. Risk factors for hospitalisation and poor outcome with pandemic A/H1N1 influenza: United Kingdom first wave (May–September 2009). Thorax 2010;65:645-51.
50. Evaluation of rapid influenza diagnostic tests for detection of novel influenza A (H1N1) virus: United States 2009. MMWR Morb Mortal Wkly Rep 2009;58:826-9.
51. Ginocchio CC, Zhang F, Manji R, Arora S, Bornfreund M, Falk L, et al. Evaluation of multiple test methods for the detection of the novel 2009 influenza A (H1N1) during the New York City outbreak. J Clin Virol 2009;45:191-5.
52. Vasoo S, Stevens J, Singh K. Rapid antigen tests for diagnosis of pandemic (Swine) influenza A/H1N1. Clin Infect Dis 2009;49:1090-3.
53. Ngaosuwankul N, Noisumdaeng P, Komolsiri P, Pooruk P, Chokephaibulkit K, Chotpitayasunondh T, et al. Influenza A viral loads in respiratory samples collected from patients infected with pandemic H1N1, seasonal H1N1 and H3N2 viruses. Virol J 2010;7.
54. Spyridaki IS, Christodoulou I, de Beer L, Hovland V, Kurowski M, Olszewska-Ziaber A, et al. Comparison of four nasal sampling methods for the detection of viral pathogens by RT-PCR-A GA(2)LEN project. J Virol Methods 2009;156:102-6.
55. Sung RY, Chan PK, Choi KC, Yeung AC, Li AM, Tang JW, et al. Comparative study of nasopharyngeal aspirate and nasal swab specimens for diagnosis of acute viral respiratory infection. J Clin Microbiol 2008;46:3073-6.
56. Heikkinen T, Salmi AA, Ruuskanen O. Comparative study of nasopharyngeal aspirate and nasal swab specimens for detection of influenza. BMJ 2001;322.

Appendix 1 Protocol, version 1.1, 8 October 2009

(PDF download)

Appendix 2 Consent forms

Appendix 3 Information sheets

Appendix 4 Eligibility checklist

Appendix 5 Recruitment leaflet

(PDF download)

Appendix 6 PCR protocol and culture protocol

Appendix 7 Composite subject charts

List of abbreviations

AC

adult in the community

AH

adult in hospital

ARI

acute respiratory infection

CC

child in the community

CH

child in hospital

CI

confidence interval

HA

haemaglutinin

HPA

Health Protection Agency

ILI

influenza-like illness

LRT

lower respiratory tract

MDCK

Madin–Darby Canine Kidney

NIHR

National Institute for Health Research

NIOSH

National Institute of Occupational Safety and Health

NPA

nasopharyngeal aspirate

PCR

polymerase chain reaction

PCT

Primary Care Trust

PPE

personal protective equipment

RSV

respiratory syncitial virus

SFM

serum-free medium

TCID

tissue culture infectious dose

URT

upper respiratory tract

VTM

viral transport medium