Form 3
Annual Report
LaMer, Ehime University
Date (28, 02, 2018)
To Director of LaMer
Principle Investigator: Ana Catarina Sousa
Affiliation: CNRS LabEx DRIIHM, France
Position: Researcher
Name in print:

Include the report on the result of the project/meeting in a separate sheet.
1. Project / Meeting title
Environmental risk factors for the development and progression of pulmonary diseases
2. Members of project / meeting
Name

Affiliation
CNRS-INEE-ECCOREV(Unité FR3098); OHMi
Estarreja-OHM Bassin
Minier de Provence;
Reasercher

Position
Researcher

Contribution part
Sampling, Chemical analysis,
Results interpretation; paper
writing

M. Ramiro
Pastorinho

Faculty of Health
Sciences, University ofBeira Interior, Portugal

Invited AssistantProfessor

Supervisor for the Project in
portugalRumi Tanoue

CMES, Ehime university

Researcher

Chemical analysis, Resultsinterpretation

LaMer Faculty
member inchargeTatsuya Kunisue

Professor

Supervisor for the projectat CMESPI
Ana Sousa

3. CONTENTS
3.1. Introduction
Over the last century, a shift from an outdoor to an indoor lifestyle occurred and presently wespend 80-90% of our lives inside buildings [1]. This artificial habitat exhibits uniquecharacteristics (e.g. insulation, humidity) that promote the accumulation of not only biologicalagents (e.g. bacteria, fungi) but also chemical contaminants and therefore it represents a prime
interface between such agents and humans [1]. However, humans are not evolutionary prepared
to deal with such an artificial habitat [2] and scientific evidence suggests that the increasing
incidence of NCDs in western societies may be a consequence of the ubiquitous exposure toenvironmental contaminants [3]. Respiratory diseases constitute a paradigmatic case study, as
they are deeply responsive to environmental contaminants. Furthermore, the economic burdenof respiratory diseases is overwhelming, exceeding 380 billion € in Europe, with asthma andchronic obstructive pulmonary disease (COPD) representing the greatest portion with over 200
billion € [4]. Recent studies have suggested that the indoor environment is an important sourceof human exposure to not only toxic chemical contaminants but also microbial communities [57] and that such exposures deleteriously affect human respiratory health [8, 9]. Furthermore,
the microbial community structure can be altered by the use of household products such as
antimicrobial agents. In our ongoing microRESPIRA project we have identified the presence of awide variety of fungi and bacteria including antibiotic resistant bacteria in house dust and indoor
air samples [10]. Our preliminary results were the first to disclose the presence of antibiotic
resistant bacteria in house dust samples. Given the interest of our dataset and in order to provide
for the first compelling evidence of the modulation of indoor microbiome by household products
and their joint effect in the respiratory status of COPD patients we focused this LaMer project on
the evaluation of antimicrobials in dust samples and matched urine samples from patients with
COPD.
3.2. Procedure
3.2.1. Sampling
Dust and urine samples from patients with COPD and respective controls were collected in
Estarreja, Portugal between January 2017 and April 2017 under the framework of RESPIRA
project. Dust samples were collected from the vacuum cleaner bag that was delivered to thevolunteers and that was used for 60 days. The samples were then sieved trough a 63µm sieveand preserved at room temperature in dark conditions until chemical analysis. Urine samples
were collected directly into sterile containers and at the laboratory of Beira Interior University,
several aliquots of 1.5ml were prepared and immediately frozen. Two aliquots of each sample
were transported to CMES in cool conditions and preserved at -20oC until analysis.
13.2.2. Chemical analysis – urine samples
Levels of triclosan, triclorocarban and parabens (Methyl, Ethyl, Propyl, Butyl) were quantified inurine samples. At first, the extraction protocol was optimized. Two different extraction
approaches were tested, liquid/liquid extraction and solid phase extraction (SPE).
a) Liquid/liquid extraction: 500 µL of hydrolyzed urine samples (see 3.2.3.) were extracted twicewith ethyl acetate, spiked with ultrapure water, and the organic phase concentrated under gentle
nitrogen flux. The target compounds were then re-dissolved with methanol/water (1:1, v/v) andthe final solution was sonicated, centrifuged and filtered using a 0.2 µm cellulose membrane
syringe filter (Satorius stedim biotech, Minisart RC4, 0.2 µm RC). The sample was preserved inamber LC glass vials at 40C until injection into the LC-MS/MS.
b) Solid phase extraction: After hydrolysis, cold methanol, ultrapure water and ammonia
solution (5% NH4OH) were added to the urine sample. The sample was then loaded into a preconditioned OASIS MAX cartridge (MTBE, Methanol and ultrapure water). Prior to elution withformic acid:MTBE:methanol = 0.2: 3: 7 (v/v/v) the cartridge was washed with 5% NH4OH andmethanol and afterwards dried for 15 min. The eluted target compounds were evaporated todryness under nitrogen flux and re-dissolved with methanol and ultra-pure water. The sample
was preserved in amber LC glass vials at 40C until injection into the LC-MS/MS.
3.2.3. Urine Hydrolysis
Urine samples were thawed at room temperature. 500 µL of the thawed urine sample was added
to a 10 mL glass vial, to which 50 µL of Internal Standards (ISs) Mixture and 125 µL of 1.0 mol/Lammonium acetate buffer (pH = 4.7) containing β-glucuronidase/aryl-sulfatase was added. The
solution was vortexed and incubated at 37oC under gentle agitation for 16hours.
The ISs mixture contained 100 ng/mL of Triclosan-13C6, 20 ng/mL of Triclocarban-13C6, 20 ng/mL
of Methyl paraben-13C6 and 20 ng/mL of Butyl paraben-13C6). The activity of β-glucuronidase/arylsulfatase was 290 units per mL of urine. The solution was prepared freshly every week by adding
4.7 ml of 1.0 mol/L ammonium acetate; 5.3 mL of 1.0 mol/L acetic acid and 50 µL of βglucuronidase/aryl-sulfatase solution (116,000 units/mL). The crude mixture of βglucuronidase/sulfatase from Helix pomatia (Type HP-2, aqueous solution, 116,000 units/mL
glucuronidase and 1020 units/mL sulfatase) was purchased from Sigma-Aldrich (St. Louis, MO,USA); (G7017).
3.2.4. Chemical analysis – dust samples
Levels of triclosan, triclorocarban and parabens (Methyl, Ethyl, Propyl, Butyl) were quantified indust samples. Since the results obtained for SPE with OASIS MAX Cartridge were better (see Table1) all the dust analysis were performed using this technique. In brief, ISs mixture was added to2

30mg of dust sample. The target compound were extracted twice by ultra-sonication withacetonitrile and methanol. After two centrifugations, the supernatants were combined andafterwards diluted with ultra-pure water. The sample was then loaded into a pre-conditioned
OASIS MAX cartridge (MTBE, Methanol and ultrapure water). Prior to elution with formic
acid:MTBE:methanol = 0.2: 3: 7 (v/v/v), the cartridge was washed with 5% NH4OH and methanol
and afterwards dried for 15 min. The eluted target compounds were evaporated to dryness
under nitrogen flux and re-dissolved with methanol and ultra-pure water. The sample waspreserved in amber LC glass vials at 40C until injection into the LC-MS/MS.
3.4. Results
3.4.1. Protocol optimization
In order to compare liquid/liquid extraction and solid phase extraction protocols 3 samples ofurine plus one procedural blank were tested, furthermore recovery tests with three samples ofurine spiked with PPCPs native working solution (100ppb) were also carried out. The obtained
results disclose higher absolute recoveries and lower coefficient of variation for antimicrobials
when using SPE with Oasis MAX cartridge (Table 1) and thus for the analysis of patients’ urine
and dust samples this protocol was used.
Table 1. Comparison of the absolute recoveries obtained using different extraction protocols for urine
samples and IS corrected recoveries when using solid phase extraction with Oasis Max Cartridge.
CompoundAbsolute recovery rate (%, %CV)
LL extraction

Methyl paraben
Ethyl paraben
Propyl paraben
Butyl paraben
Triclocarban
Triclosan

8.2 (0.38)
5.9 (1.4)
28 (1.6)47 (0.88)
54 (6.7)
9.8 (6.2)

SPE (type of cartridge)
Oasis MCX
Oasis MAX
4.4 (4.2)
28 (4.9)3.9 (3.1)
25 (4.1)
21 (2.1)
36 (3.3)
34 (1.6)
49 (3.6)
3.3 (7.3)
31 (21)7.8 (4.7)
78 (3.1)

IS corrected recovery rate(%, %CV)97 (2.0)
89 (3.9)79 (1.7)
110 (0.58)
100 (2.5)
100 (2.4)

3.4.2. Urine and dust samples
Table 2 depicts the levels of parabens (Methyl, Ethyl, Propyl, Butyl) and antimicrobials (triclosanand triclorocarban) in human urine and dust samples. Overall, the concentrations in dust
samples are one to two orders of magnitude higher that the concentrations in human urine.
Triclosan was detected in 56% of urine samples and in all the dust samples, triclocarban was
bellow detection limit (0.25 ng/mL) in all urine samples and was detected in 82% of the dust
samples. Amongst parabens, methyl paraben exhibited the highest concentrations in both typeof samples. Due to the limited number of dust samples available, it was not possible to find any

3

association between dust and urine levels. Interestingly, the highest level reported in dust for
triclosan (1200 ng/g) corresponded to the house of the patient with the highest triclosan
concentration in urine (140 ng/mL). Furthermore in this house, bacteria resistant to ampicillin
were also detected in dust, with over 78% of the bacteria detected being resistant to thisantibiotic. Such results, although preliminary suggest that the use of antimicrobials might beassociated with the presence of resistant bacteria and thus deserve to be further studied.
Table 2. Summary of the concentrations of parabens (methyl, ethyl, propyl, butyl) and antimicrobials(triclosan and triclorocarban) in urine (ng/mL) and dust (ng/g).
Parabens

Urine(n=43)% detection
minmax
medianmean
stdev
Dust (n=11) % detection
minmax
medianmean
stdev

Antimicrobials

MethylEthyl

PropylButyl

Triclocarban

Triclosan

72
<MDL
1600
6.7
63
240
91
<MDL
490
170
220
160

84
<MDL
160
2.3
12
28
91
<MDL
420
36
85
120

44
<MDL
130
<MDL
6.8
22
91
<MDL
370
77
120
130

23
<MDL
5.1
<MDL
0.15
0.77
100
0.73
46
12
17
16

0.0
<MDL
<MDL
<MDL
<MDL

56
<MDL
430
0.35
15
67
100
23
1200
105.0
275.9
362.0

82
<MDL
160
7.2
29
46

<MDL: below the method detection limits
3.5. Future perspectives
We are planning to include the triclosan and triclocarban dust results in the paper that is being
prepared on the house dust samples bacterial community. We expect to have all the data
concerning bacterial community characterization by the end of April 2018 and to submit the
paper by July 2018.
3.6. Achievements
3.6.1. List of papers published:
The impact factor™ (IF) and quartile values corresponds to Thomson Reuters metrichttps://jcr.incites.thomsonreuters.com. (*) Corresponding authorSousa ACA*. Tanabe S. Pastorinho MR (2017) Organotins: Sources and Impacts on
Health and Environment. Reference Module in Earth Systems and Environmental
Sciences. Elsevier. http://dx.doi.org/10.1016/B978-0-12-409548-9.09986-3. This paper
was also published in the Encyclopedia of the Anthropocene (see 3.6.2).
4



Coelho SD. Maricoto T. Pastorinho MR. Itai T. Isobe T. Kunisue T. Tanabe S. Sousa ACA*.
Nogueira AJA (2017) Cadmium intake in women from Aveiro University. Portugal – a
duplicate diet study. Journal of Geochemical Exploration 183(B): 187-190;
http://dx.doi.org/10.1016/j.gexplo.2017.02.003; 5 year IF 2016: 3.024. Q2. 62.5%

Coelho SD. Maricoto T. Tanabe S. Nogueira AJA. Sousa ACA* (2017) Dietary Habits of a
Portuguese Academic Community - A Food Frequency Questionnaire Approach. Journalof Nutrition and Diabetes Research 1 (1)3.6.2. Book chapters:Sousa ACA. Tanabe S. Pastorinho MR (2018) Organotins: Sources and Impacts on
Health and Environment In Encyclopedia of the Anthropocene. A2 – Dellasala DA.
Goldstein MI (Eds). (Oxford: Elsevier). pp. 133-139.
https://www.sciencedirect.com/science/article/pii/B9780128096659099869

3.6.3. Oral Communications by InvitationSousa ACA (2018) “Environmental contaminants and endocrine disruption: The story ofobesogens”. 20th European Congress of Endocrinology. Symposium 4: “Environmental
Effects on Endocrine Function”. S4.2. Scheduled for 20 May 2018. Barcelona. Spain

3.6.4. Oral Communications
Sousa ACA. Silva T. Amaro R. Miranda S. Marques A. Valente C. Pastorinho MR. Pereira
CC. Teixeira JP. Henriques I. and the RESPIRA Group (2017) microRESPIRA:
microenvironmental risk factors for the progression of pulmonary diseases. 5º Colloquede Restitution do OHM- Estarreja. 3 November 2017. Aveiro. Portugal. P.7

References:
1. Sousa. A.C.A.. S. Takahashi. and S. Tanabe. Organic contaminants in house dust. Current
Organic Chemistry. 2014. 18(17): p. 2181.
2. Maziak. W.. The asthma epidemic and our artificial habitats. BMC Pulmonary Medicine. 2005.
5(1): p. 5.
3. WHO. Global status report on noncommunicable diseases 2014. 2014. World HealthOrganization: Geneve. p. 280.
4. European Lung Association. European Lung White Book. 2013.
5. Prussin. A.J.. E.B. Garcia. and L.C. Marr. Total Concentrations of Virus and Bacteria in Indoor
and Outdoor Air. Environmental Science & Technology Letters. 2015.
6. Kettleson. E.M.. et al.. Key determinants of the fungal and bacterial microbiomes in homes.
Environmental Research. 2015. 138(0): p. 130-135.
7. Barberán. A.. et al.. The ecology of microscopic life in household dust. Proceedings of the Royal
Society of London B: Biological Sciences. 2015. 282(1814).
8. Sousa. A.C.A.. et al.. Characterization of Fungal Communities in House Dust Samples Collected
From Central Portugal—A Preliminary Survey. Journal of Toxicology and Environmental Health.
Part A. 2014. 77(14-16): p. 972-982.
9. Sharpe. R.A.. et al.. Indoor fungal diversity and asthma: A meta-analysis and systematic
review of risk factors. Journal of Allergy and Clinical Immunology. 2015. 135(1): p. 110-122.
10.Silva. T.. Microbial community in houses from patients with chronic respiratory diseases in
Estarreja. in Biology Department. 2017. University of Aveiro: Aveiro.
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