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M. González, R. Vieira, P. Nunes, T. Rosado, S. Martins, A. Candeias, A. Pereira and A. T. Caldeira, Fluorescence In Situ Hybridization: a potentially useful technique for detection of microorganisms on mortars, e-conservation Journal 2, 2014, pp. 44-52
Available online 21 May 2014



Fluorescence In Situ Hybridization: a potentially useful technique for detection of microorganisms on mortarss 

Marina González, Ricardo Vieira, Patrícia Nunes, Tânia Rosado, Sérgio Martins, António Candeias, António Pereira and Ana Teresa Caldeira




Abstract
This paper discusses the possibilities of applying Fluorescence In Situ Hybridization (FISH) to detect microorganisms on mortars, as this analytical technique has been used in different fields for the detection and identification of individual microbial cells in situ. FISH technique was applied for microbial detection on test and real mortars inoculated with fungal suspensions of S. cerevisae 396 and Nectria sp. A universal eukaryotic probe (EUK516) labelled with fluorescent dye (Cy3) was tested with different cell fixation procedures (4% (w / v) paraformaldehyde or 50% (v / v) ethanol in PBS). Positive results were obtained with FISH detection of Nectria on testing/artificial as well as authentic/historical mortars, which confirms successful application of FISH technique to a new  on mortars.


1. Introduction

Microbial activity plays an important role in the deterioration of built heritage. Although the influence of microorganisms on deterioration processes is undisputable, the role of individual microbial species that form the communities is not yet fully understood. The development of new microbial detection and identification techniques is crucial for furthering our knowledge of microbial influence on heritage deterioration and designing appropriate preservation strategies.

Detection and identification of microbial communities present on artwork can be achieved using various complementary methods, with new approaches being continuously developed. The traditional culture-based techniques are time-consuming and are limited by the microorganisms’ ability to grow under standard laboratory cultivation conditions [1, 2]. To overcome this drawback, culture–independent techniques based on molecular approaches, that are more sensitive and need smaller quantities of sample than those previously mentioned, have been applied. The use of molecular techniques based on expensive Polymerase Chain Reaction (PCR) present an important limitation, i.e. the impossibility of studying the microorganisms in situ [1, 2]. Nevertheless, a “non-PCR”-based technique is available that combines the precision of molecular techniques with providing information on the number and spatial distribution of microorganisms: Fluorescence In Situ Hybridization (FISH) [3]. As well as being inexpensive and informative, this analytical technique is also very powerful, rapid and straightforward [4]. Surprisingly, only a few studies in the field of cultural heritage conservation and restoration have exploited the potentials of using FISH method [5-8], despite it being the most commonly applied non-PCR-based method in other fields [1, 2, 9]. FISH allows the detection of microorganisms by a fluorescently labelled oligonucleotide target probe that hybridizes specifically to its complementary target sequence within the cell [2]. The selection of rRNA probes enables phylogenetic specificity to be varied from universal to subspecies level [9].

This study applied FISH technique for direct detection of microorganisms on mortars, using a rRNA probe, as a complementary technique used for characterization of the microbiological community. FISH was applied to a series of test and real mortars, which were inoculated with cell suspensions of Saccharomyces cerevisiae 396 (yeast) and Nectria sp. (filamentous fungi), from a laboratory collection. Nectria sp. was selected because its involvement in deterioration of mortars of historical Alentejo buildings (Évora, Portugal) has been previously confirmed. On the other hand, S. cerevisiae, although not biodeteriogenic, has been selected, because its detection by FISH in suspension using EUK516 labelled with a fluorophore has been used successfully in the past [10].

This preliminary study was undertaken to evaluate the applicability of this technique, using specific probes, as a simple, rapid and efficient tool to identify biodeteriogenic agents in mortars.

 
2. Material and Methods

FISH technique was performed on a series of test and real mortars, which were inoculated with fungal suspensions. The FISH results were compared to data obtained from techniques that evaluate presence of microorganisms, their proliferation and metabolic activity.

2.1. Sample Preparation

2.1.1. Mortars
Both test and real mortars were used to investigate the applicability of FISH technique for the detection of microorganisms. Real mortars were collected from damaged frescoes of the abandoned church of Santo Aleixo (Montemor-o-Novo, Évora, Portugal). Mortars microfragments (50 mg), were sampled with sterile scalpels and microtubes using micro-invasive methods and then sterilized.

2.1.2. Preparation of Microbial Suspension
Two fungal suspensions were prepared using Saccharomyces cerevisiae 396 (SC396) and Nectria sp. microorganisms belonging to laboratory collection (HERCULES-Biotech Laboratory,  University of Évora). The strains were grown in Malt Extract Agar (MEA) slants at 28°C for 1 day (yeast) or 5 days (filamentous fungi). Cells were harvested from the agar surface and then suspended in NaCl 0.85% (w/v) aqueous solution.

2.1.3. Inoculation of Mortars
Mortars (1.5 g) were inoculated with 1.0 mL of each fungal suspension at room temperature for 24 h.

2.2. Detection and Evaluation of Microbial Contamination

2.2.1. Fluorescence In Situ Hybridization
Each sample (0.1 g of mortars) was washed once with 0.5 mL phosphate buffered saline solution (PBS; 130 mM NaCl, 8 mM NaH2PO4, 2.7 mM KCl, 1.5 mM KH2PO4, pH 7.2) and fixed for 4 h with 1 mL of fresh fixation buffer solution (4% w/v paraformaldehyde in PBS at 4ºC or 1:1 (v/v) PBS/ethanol at 20ºC). To avoid cellular aggregation the fixatives were added drop by drop. Following centrifuging for 5 min at 4500 g, the supernatant was discarded. Cells fixed with paraformaldehyde were washed three times and those fixed with ethanol were washed once. For hybridization,   80 μl of hybridization buffer (0.9 M NaCl, 20 mM Tris–HCl, 0.1% SDS) and 1 μl (120 ng/ μl) of the probe EUK516 (5’-ACCAGACTTGCCCTCC-3´) rRNA labelled with indocarbocianine (Cy3) were added. Then, samples were incubated at 46ºC for 2 h and centrifuged for 5 min at 4500 g. The supernatant was discarded and the mortars were washed with 100 μl of pre-warmed hybridization buffer for 30 min at 46ºC. The samples were then centrifuged  5 min at 4500 g and the supernatant discarded. The mortars were spotted onto microscope slides and observed with a Leica DM 2500M darkfield microscope. Image caption was carried out with a Leica DFC290HD camera.

2.2.2. Biological Activity Assay
The colorimetric 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay was used to estimate the cell viability in mortars as previously described by Rosado et al [11]. It is based on the ability of living cells to reduce metabolically the pale yellow MTT to a purple formazan salt. Mortar samples (0.1 g) were incubated with 0.5 mL of MTT solution (0.5 mg/mL in PBS) for 4 h at room temperature in the dark. After that, the mixture was centrifuged for 10 min at 8000 g and the supernatant discarded. The resulting purple formazan crystals were dissolved in 1 mL of DMSO/ ethanol (1:1) and absorbance intensity at 570 nm was then measured with a spectrophotometer (Hitachi, U-3010).

2.2.3. DNA Extraction and Amplification
Metagenomic DNA was directly extracted from the mortar microfragment (0.1 g) collected with NucleoSpin740945 DNA Extraction kit (Macherey-Nagel, Düren, Germany). The extracted DNA was used as template for PCR amplification. A partial sequence of 18S rDNA gene was amplified using primer pair NS1 (5’-GTAGTCATATGCTTGTCTC-3’) / GCfung (5’-CGCCCGCCGCGCCCCGCGCCCGGCCCGCCGCCCCCGCCCCATTCCCCGTTACCCGTTG-3’) [12, 13].

PCR reactions were carried out in a Robocycler (MJ Mini Bio-Rad). The PCR mixture (25 µL) contained reaction buffer 10x supplied with MgCl2 25 mM, dNTPs 2 mM, primer sets 0.4 µM, Taq DNA polymerase 5 U and 1 µL of the extracted DNA. The PCR program consisted of an initial denaturising step for 1 min at 95ºC, followed by 40 cycles with a denaturising step at 94°C for 1 min, an annealing step at 50°C for 1 min, an extension step at 72°C for 2 min and a final elongation cycle at 72°C for 6 min. The PCR products, the molecular-weight size marker Generuler 100 bp (Fermentas, Lithuania) and a control sample were run on a 1.2% agarose gel containing ethidium bromide 10 mg/ mL, at 90 V and at room temperature. The amplification products were visualized under UV light (Bio-Rad system).

2.2.4. Microbiological Proliferation Assessment
Microanalysis of the selected fragments of mortars was performed using a HITACHI 3700N scanning electron microscope (SEM) coupled with a Bruker AXS X-ray energy dispersive (EDX). The accelerating voltage was 18–20 kV. This technique allowed the observation of mortars’ microstructure and morphology, as well as microbial contaminations and elemental composition (point analysis and 2D mapping).


3. Results and Discussion

Positive results were obtained from FISH experiments using an universal probe EUK516-Cy3 on test sterile mortars inoculated with Nectria and SC396 using both paraformaldehyde (4%) and ethanol (50%) as fixing agents. FISH results of a sample inoculated with Nectria, shown in Figure 1, reveal that it is possible to detect the microorganism on a mortar sample. Hybridization of SC396 was  not observed. A control test was carried out by adding NaCl 0.85% (w/v) aqueous solution to a mortar sample instead of a microbial suspension. The results for the paraformaldehyde control reveal that paraformaldehyde residues remained detectable but did not interfere with microorganism detection. However, as no differences in fluorescence intensity were observed for paraformaldehyde and ethanol, the latter is initially preferred to avoid interferences of residue detection.

Once it was confirmed that FISH technique could be successfully applied for detection of microorganisms (at least filamentous fungi in the first instance) on test mortars, the FISH technique was then applied to samples of real mortars. Both fixing agents, 50% ethanol and 4% paraformaldehyde, were tested in order to evaluate them, to develop a protocol for detection of microbial communities on real mortars. In situ hybridization signals were not obtained for SC396 in real mortars. However, positive results were obtained for Nectria with the universal probe EUK516-Cy3 (Figure 2) which confirmed good cell permeability conditions of Nectria with the fixing agents and revealed the possibility of applying FISH technique to real mortar samples. Complementary techniques were also used for the following purposes: i) to evaluate microbial contamination in mortars inoculated with SC396 and to confirm their presence in investigated samples; and ii) to compare the results obtained by FISH technique for Nectria. In order to fulfill these considerations, the following techniques were used: i) SEM analysis and MTT assay, useful for the evaluation of microflora proliferation and their biological activity [11]; and ii) molecular techniques, that allowed the recognition of the type of microorganisms present in the samples.

Evidence of biological contamination was observed in mortar microfragments analyzed by SEM. The image obtained by SEM allows the observation of fungal hyphae belonging to filamentous fungi penetrating the microstructure of the mortars inoculated with Nectria (Figure 3a). The images obtained from mortars, inoculated with SC396, reveal the presence of yeast cells (Figure 3c). Furthermore, EDX analyses (Figures 3b, 3d) of the structures of both samples confirm the presence of elements characteristics of organic material such as carbon and nitrogen, which points to the presence of microbial contamination. Thus, SEM-EDX results confirm the presence of fungi in the mortar samples.

Left to right:
Figure 1. Microscopic images of test mortars inoculated with Nectria fixed with paraformaldehyde (a–c) and ethanol (d-f). The same area is shown in the bright-field (a, d), a combination of dark– and bright–field (b, e) and dark–field (c, f) observation.
Figure 2. Visualization by FISH of Nectria in real mortars fixed with paraformaldehyde (a–c) and ethanol (d-f). The same area is shown in the bright-field (a, d), a combination of dark- and bright-field (b, e) and dark–field (c, f) observation.
Figure 3. SEM micrograph and EDX elemental composition map of mortars  inoculated with fungi.



4. Molecular Evidence of Fungi

The presence of fungi cells in mortar samples was also investigated by molecular techniques. The metagenomic DNA was extracted and amplified by PCR. The amplified products were run on an agarose gel. DNA from the mortar samples showed positive amplifications of 18S rDNA fungal primers NS1/ Fung-GC (Figure 4). The size of the PCR products of approximately 400 bp suggested the presence of fungi on mortar samples.

Metabolic activity was also investigated on mortar samples inoculated with fungal suspensions using the MTT assay, as the appearance of purple formazan crystals, proportional to the living cells in the sample, can be a useful and fast marker. A more intense colour is associated with a higher cell viability in a sample. Thus, the more intense coloured solution obtained for SC396 sample, (associated to a higher relative MTT response), in comparison with that correspondent to the Nectria sample, points to a higher metabolic activity in the mortar inoculated with SC396 rather than in the Nectria sample as it is shown in Figure 5.

Left to right:
Figure 4. Gel electrophoresis of PCR products amplified from DNA extracted from mortars inoculated with SC396 and Nectria (1 and 2, respectively). Lane M corresponds to the 100 bp DNA molecular-weight size marker and lane C to the control.
Figure 5. Relative MTT response for the mortars inoculated with SC396 and Nectria. The relative responses were calculated using the absorbance at 570 nm for 107 living cells of SC396 as a reference.


The results obtained by complementary techniques for the analysis of real mortars inoculated with Nectria, confirmed the presence of fungi contamination, which are in agreement with the results obtained by FISH technique. This positive correlation of results obtained by various analytical techniques shows that FISH technique could be applied to detect at least filamentous fungi. However, none FISH signals can be visualized for mortars inoculated with yeast, despite the fact that complementary approaches used (SEM-EDX, PCR-based and MTT,) showed the presence of yeast cells and their high metabolic activity.

The absence of FISH signals could be due to the problems that have been previously reported for FISH technique [3]: a low cellular ribosome content, the impermeability of cell walls or the accessibility of probe target sites. Further investigations should be carried out to improve the analytical method for SC396 detection in mortars.

Future work should involve testing various fixing agents, fixation times and the design of specific probes for specific biodeteriorating agents to improve their detection as well as identification in mortars.


5. Conclusions

This preliminary investigation demonstrates that a simple, rapid and cheap FISH analytical technique could be applied for the detection of at least filamentous fungi on mortars, with further need to test the analytical protocol for each probe. The results obtained by FISH were confirmed by SEM-EDX and PCR-based techniques, that revealed fungi contamination, together with the use of MTT assay, which revealed the presence of active metabolic cells in mortar samples.


6. Acknowledgments

The authors would like to thank the “IMAGOS – Innovative Methodologies in Archaeology, Archaeometry and Geophysics – Optimizing Strategies X APOLLO - Archaeological and Physical On-site Laboratory- Lifting Outputs“ Project (ALENT-07-0224-FEDER-001760) for supporting the research reported in this work.


7. References

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[2] A. Moter, U. B. Göbel, Fluorescence in situ hybridization (FISH) for direct visualization of microorganisms, Journal of Microbiological Methods 41(2), 2000, pp. 85-112, doi: 10.1016/S0167-7012(00)00152-4

[3] B. Bottari, D. Ercolini, M. Gatti, E. Neviani, Application of FISH technology for microbiological analysis: current state and prospects, Applied Microbiology and Biotechnology 73(3), 2006, pp. 485-494, doi: 10.1007/s00253-006-0615-z

[4] Y. Aoi, In situ identification of microorganisms in biofilm communities, Journal of Bioscience and Bioengineering 94(6), 2002, pp. 552-556, doi: 10.1016/S1389-1723(02)80194-3

[5] F. Cappitelli, C. Sorlini, Microorganisms Attack Synthetic Polymers in Items Representing Our Cultural Heritage, Applied and Environmental Microbiology 74(3), 2008, pp. 564-569, doi: 10.1128/AEM.01768-07

[6] C. Urzì, V. La Cono, E. Stackebrandt, Design and application of two oligonucleotide probes for the identification of Geodermatophilaceae strains using fluorescence in situ hybridization (FISH), Environmental Microbiology 6(7), 2004, pp. 678- 685, doi: 10.1111/j.1462-2920.2004.00619.x

[7] V. La Cono, C. Urzı̀, Fluorescent in situ hybridization applied on samples taken with adhesive tape strips, Journal of Microbiological Methods 55(1), 2003, pp. 65-71, doi: 10.1016/S0167-7012(03)00115-5

[8] C. Urzı̀, F. De Leo, Sampling with adhesive tape strips: an easy and rapid method to monitor microbial colonization on monument surfaces, Journal of Microbiological Methods 44(1), 2001, pp. 1-11, doi: 10.1016/S0167-7012(00)00227-X

[9] R. I. Amann, B. J. Binder, R. J. Olson, S. W. Chisholm, R. Devereux, D. A. Stahl, Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations, Applied and Environmental Microbiology 56(6), 1990, pp. 1919-1925

[10] A. Xufre, H. Albergaria, J. Inácio, I. Spencer-Martins, F. Gírio, Application of fluorescence in situ hybridisation (FISH) to the analysis of yeast population dynamics in winery and laboratory grape must fermentations, International Journal of Food Microbiology, 108(3), 2006, pp. 376-384, doi: 10.1016/j.ijfoodmicro.2006.01.025

[11] T. Rosado, M. R. Martins, M. Pires, J. Mirão, A. Candeias, A. T. Caldeira, Enzymatic Monitorization of Mural Paintings Biodegradation and Biodeterioration, International Journal of Conservation Science 4(SI), 2013, pp. 603-612

[12] L. M. Duong, R. Jeewon, S. Lumyong, K. D. Hyde, DGGE coupled with ribosomal DNA gene phylogenies reveal uncharacterized fungal phylotypes, Fungal Diversity 23(1), 2006, pp. 121-138

[13] T. Rosado, J. Mirão, A. Candeias, A.T. Caldeira, Microbial communities analysis assessed by pyrosequencing — a new approach applied to conservation state studies of mural paintings, Analytical and bioanalytical chemistry 406(3), 2014, pp. 887-895, doi: 10.1007/s00216-013-7516-7



Marina González
Physical Chemist, Postdoc in Biochemistry
HERCULES Laboratory, University of Évora (Portugal)
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Ricardo Vieira
Biochemist, Master student in Biochemistry
Chemistry Department, University of Évora (Portugal)
HERCULES Laboratory, University of Évora (Portugal)
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Patrícia Nunes
Biochemist, Bachelor in Biochemistry
HERCULES Laboratory, University of Évora (Portugal)
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Tânia Rosado
Biochemist, PhD candidate in Biochemistry
Chemistry Department, University of Évora (Portugal)
HERCULES Laboratory, University of Évora (Portugal)
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Sérgio Martins
Chemist, PhD candidate in Organic Chemistry
HERCULES Laboratory, University of Évora (Portugal)
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António Candeias
PhD in Chemistry, specialization in Surface Chemistry and Heritage Sciences
Chemistry Department, University of Évora (Portugal)
Évora Chemistry Centre, University of Évora (Portugal)
HERCULES Laboratory, University of Évora (Portugal)
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António Pereira
PhD in Chemistry, specialization in Organic Chemistry
Chemistry Department, University of Évora (Portugal)
Évora Chemistry Centre, University of Évora (Portugal)
HERCULES Laboratory, University of Évora (Portugal)
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Ana Teresa Caldeira
PhD in Chemistry, specialization in Biochemistry/ Biotechnology
Chemistry Department, University of Évora (Portugal)
Évora Chemistry Centre, University of Évora (Portugal)
HERCULES Laboratory, University of Évora (Portugal)
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