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Microscopy: Science, Technology, Applications and Education
A. Méndez-Vilas and J. Díaz (Eds.)
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Morphological changes induced in bacteria as evaluated by electron
microscopy
J. Díaz-Visurraga1, G. Cárdenas1,2, and A. García3
1
CIPA Chile, Research Center of Advanced Polymers, Edmundo Larenas 129, Concepción, VIII Region, Chile.
Advanced Materials Laboratory, Department of Polymers, Faculty of Chemical Sciences, University of Concepcion,
Concepción, Chile.
3
Department of Microbiology, Faculty of Biological Sciences, University of Concepcion, Concepción, Chile.
2
In terms of the physical bacteria-biocide interaction and molecular mechanisms associated, several interesting questions
remain to be answered. How do explain the apparent discrepancy between increased culture turbidity and decreased
bacterial viability? How do the biocides kill bacteria without lysis? Cell death is always accompanied by cell wall
degradation? The information acquired from microscopic images permits the partial understanding of bacterial physiology
and mechanisms of biocide action. Electron microscopic approaches in the study of bacteria and their physiological
consequences when it is affected by biocides permit the observation of characteristic morphological defects of cell wall
and provide valuable insights into processes involved on bacterial death. This article focuses on techniques for preparing
samples of damaged bacteria for conventional transmission (TEM) and scanning electron microscopy (SEM) and the
resulting observations that suggest morphological alterations in bacteria such as blockage of nutrients flow, disaggregation
of colonies, increase of cell wall roughness, cell disruption with loss of intracellular material, cytolytic damage, process of
morphological transformation, filamentation and bacteriolysis.
Keywords: Scanning Electron Microscope, Transmission Electron Microscope, artifacts, filamentation, cytolytic damage,
bacteriolysis.
1. Introduction
Nowadays, there are several known bacterial mutants resistant to all available antibiotics [1, 2]. The increased
prevalence of pathogens causing opportunistic infections in humans and animals underscores the imperative need to
develop new and effective biocides. Since 1946, several papers were published on the use of electron microscopy as a
means to provide relevant information about discrete cellular phenomena and damage which are inaccessible via
traditional means [3-5]. However, the information provided by electron micrographs must be supported with the
examination of metabolic activities (biochemical assays) of the organism. The in vitro time-kill assays are the first
approach in order to elucidate the extent of antibacterial activity of a biocide.
The knowledge of bacterial cell morphology is required for the understanding of the probable mechanisms of action of
biocides. In principal, the use of electron microscopy may provide information on the number, size, shape of cells and,
information of the microcolony at the same time. The typical size and shape of bacterial cells is dependent on numerous
variables that are unique to the particular environments associated with the lifestyle of each species [6].
From the microscopic observations it could be speculate various phenomena associated with cell damage:
1. Selectivity and binding site. The modes of action of biocides depend on the type of bacterial organisms and
are mainly related to the structure of the cell wall and outer membrane arrangement. This is likely due to the
significant differences in the outer layers of Gram-negative and Gram-positive bacteria. Gram-negative
bacteria possess an outer membrane and a unique periplasmic space not found in Gram-positive bacteria.
2. Ability to bind the cell wall. The attack of biocide to an organism might reflect their ability to bind and
penetrate the cell wall to gain access to target molecules in the cell membrane. For example, compared with
macromolecular biocides, the nanoparticles are expected to yield higher local concentrations of charged
particles at the bacterial membrane, thus enhancing the antimicrobial activity.
3. Morphological effect. The morphological effect is evidenced by the change of form, promoting
interactions and disruption of the cell wall. For instance, Penicillin-binding-proteins (PBPs) located in the
cytoplasmic membrane, which bind penicillins and other β-lactam antibiotics, determine the bacterial cell
shape [7], so changes in cell morphology could be involved in PBPs-biocide interaction.
4. Cofactors. The leakage of divalent and monovalent cations is an early sign of increased permeability of the
plasma membrane of bacterial cell. Potassium is a major intracellular cation that plays a crucial role in the
generation of membrane potential, activation of cytoplasmic enzymes and maintenance of turgor pressure in
cells [8].
5. Bacterial resistance. Bacteria retain genes that encode proteins that are dedicated to the purposeful
alteration of the overall bacterial cell length under certain conditions [9]. For instance, wwhatever the
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mechanism of action of biocides against E. coli, the final balance between lysis and production of nonviable,
structurally intact bacterial filaments clearly depends on the relative degree of inhibition of theirs PBPs [10].
6.Modulation of lysis or lethality. The modulation of lysis but not lethality appears when the inhibition of
PBPs occurs simultaneously with the inhibition of other PBPs [10]. For example, using β-lactam
antimicrobial agents of high affinity for a single PBP (in the appropriate concentration range), it has been
demonstrated that the inhibition of PBP lb [11] is associated with rapid killing and lysis [12], whereas
inhibition of PBP 2 produces round cells and inhibition of PBP 3 produces filaments [13] in E. coli cells.
The purpose of this paper is to point to some of the applications and contributions of conventional electron microscopy
to elucidate the mechanism of action of biocides and to highlight the advantages. In the following, the experimental
procedure of electron microscopy techniques and thin sections is outlined. Present findings will be illustrated by results
from our current research projects.
2. Experimental procedure
Organisms and materials
Methicillin resistant Staphylococcus aureus UA 2120 (MRSA, clinical isolate from wound secretion, Hospital Regional
de Antofagasta, MIC > 512 µg/mL Oxacillin, resistant to P-AMP-SAM-OXA-EM-RIF-GE-CIP-C-AK, sensitive to TEVA-SXT, β-lactamase: +, gene Blaz PCR: +, gene mecA PCR: +) was kindly gifted by Prof. Juan Silva (Departamento
de Microbiología, Facultad de Ciencias de la Salud, Universidad de Antofagasta, Antofagasta, Chile). This strain is
available upon request to Dr. Silva. Salmonella enterica serovar Typhimurium (clinical isolate from human feces) and
Pseudomonas aeruginosa ATCC 27853 (P. aeruginosa), Staphylococcus aureus ATCC 25923 and ATCC 6538P
(Staph. aureus ATCC 25923 and 6538P), Escherichia coli ATCC 25922 (E. coli) were obtained from American Type
Culture Collection (Rockville, MD, USA).
The bacteria were maintained in Trypticase soy broth (HiMedia Laboratories, Mumbai, India) as 15% glycerol stock
at -70 °C. For all experiments, the bacterial inoculum was prepared from previously frozen inocula by thawing on
Mueller Hinton agar (MHA, HiMedia Lab) plates. Bacteria from previously frozen inocula were diluted with 6 mL
MHB and incubated at 37°C for 24 h.
Lactobacillus rhamnosus 25A (L. rhamnosus, clinical isolate from human gastric mucosa) were identified at the
Diagnostic Microbiology Laboratory of Department of Microbiology, Faculty of Biological Sciences, University of
Concepcion.
Lactobacillus casei Shirota strain (L. casei Shirota strain) was obtained from the fermented milk product Yakult. L.
casei was grown overnight at 37 °C in DeMan-Rogosa-Sharpe agar (MRS, Oxoid Ltda, Cambridge, UK) and
resuspended in nanopure water.
Stock culture of both Lactobacillus were maintained in MRS broth as 15% glycerol stock at -70ºC. Lactobacillus
were prepared from previously frozen inocula by thawing on MRS agar plates, later, the inocula were diluted with 6 mL
MRS broth and incubated at 37°C for 24 h.
Helicobacter pylori ATCC 43504 (H. pylori) was grown in Columbia blood agar base (Oxoid Ltda, Cambridge, UK)
plates containing 5% horse blood and H. pylori-selective supplement of Dent (Oxoid Ltda) in microaerophilic
conditions (CampyGen, Oxoid Ltda) at 37°C for 5 days [14].
Inhibition zone test and liquid cultures
Suspensions of bacteria at 1.0 x 105 CFU/mL were made and streaked in at least three directions over the surface of the
Mueller-Hinton agar to obtain an uniform growth. After the plates were dried for 5 min, ~0.06 g of different biocides
(polysaccharide-metal complexes and cationic polysaccharide derivatives as films) were placed on the top of agar. The
plates were incubated at 37 °C overnight. For liquid cultures, an aliquot of 10 µL of the culture (1.0 x 105 CFU/mL) was
transferred to 30 mL sterile MHB in a 100 mL Erlenmeyer flask. Biocides equivalent to 0.06-0.32 g were added to the
test flask. The flask was placed in an incubator at 37°C/overnight, and time interval sampling was performed to observe
the bacteria at microscope. The wash procedure was as follows: rinse with sterile nanopure water, centrifuge at
10000g/min for 3 min, remove supernatant and repeat this procedure 2 times.
Artifacts. In order to illustrate the recognition of artifacts, inocula of MRSA UA 2120 strain were grown for 16 h in
MHA and resuspended in saline buffer.
Process of morphological transformations. In order to evidence the change from spiral to coccoid form of H. pylori,
an agar well diffusion assay was employed [15]. H. pylori cultures were plated on fresh Columbia blood agar plates
without antibiotics (107 CFU per plate), and wells (7 mm) were drilled into the agar by using sterile Pasteur pipettes. 50
µL aliquots of fresh L. rhamnosus cultures (108 CFU/mL) were suspended in the agar wells. After incubation in
microaerophilic conditions at 37°C for 5 days [14], the bacteria around the edge of inhibition growth zone (inhibition
zone test) were collected, transferred to Eppendorf tubes and washed three times with sterile nanopure water.
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Transmission electron microscopy and image preprocessing
Treated bacterial and control cells were processed by the same procedure for all techniques.
Thin section. Following the overnight incubation, the bacteria around the edge of inhibition growth zone produced by
films (inhibition zone test) were collected, transferred to Eppendorf tubes and washed three times with sterile nanopure
water. Finally, the bacteria were resuspended in 1 mL sterile nanopure water. Fractions of bacterial suspension were
fixed at 4°C for 90 min with 2% glutaraldehyde in 0.1 mol/L sodium phosphate buffer, pH 7.4. Cells were postfixed in
1% OsO4 for 90 min at 4°C and stained with 0.25% uranil acetate at 4°C for 1 h. Finally, the cells were embedded into
Spurr resin and left to polymerize in an oven at 60°C for 24 h. Resins were sectioned by cutting an 80-nm film at 25°C
using an ultramicrotome Sorvall MT 5000 (Dupont, Boston, MA) equipped with a diamond knife. Thin section was
mounted on copper grids covered with carbon film and observed using a TEM JEOL-JEM 1200 EX II (Jeol Ltd, Tokyo,
Japan) at an accelerating voltage of 80 kV.
Deposition on grid. Aliquots of 20 µL of the liquid cultures were deposited on TEM copper grids covered with
carbon film. Bacterial cells were then fixed with 20 µL of 5% formaldehyde [16] without staining. The samples were
promptly observed under the TEM.
Image processing. All images were acquired as 512 x 512 pixels, digitized and stored in an image analysis program
(DigitalMicrograph TM 3.7.0., Gatan Inc. [17]). Specimens that showed artifacts of cutting and staining for TEM were
excluded. Also, the public domain Java image processing software ImageJ 1.40g (Wayne Rasband, NIH, USA [18]) and
MBF ImageJ collection of plugins and macros were used to process the selected micrographs (binarization,
segmentation and extraction of local and dynamic threshold) [19]. It is important to note that the processing was
performed ONLY to make visualization of micrographs easier.
Scanning electron microscopy. For scanning electron microscopy, a drop of bacterial suspension was mounted onto a
gold-coated glass slide and air-dried before the image collection in a JEOL-JEM 6380 LV SEM (Jeol Ltd), operating at
an accelerating voltage of 20 kV. Histograms of cell length and width were obtained using the software Origin 6.0 [20].
EDX spectra were acquired with an INCA X-Sight 7582 EDX device (OXFORD Instruments, UK).
3. Applications and findings
3.1
Morphology
Untreated cells must be studied as control (untreated bacteria) to ensure the observed differences at electron microscope
between control and the bacterial cells exposed to the biocide. Fig. 1a shows SEM image of P. aeruginosa and their
corresponding histograms of cell length and width (Fig. 1b and 1c). Scanning electron microscopy of P. aeruginosa
showed that they exist as colonies of rod-shape bacteria in control culture with an average (n = 100) cell length of 1.04
µm and cell width of 0.41 µm. This Gram-negative rod (Fig. 1d) exhibit extra-cellular filamentous materials detectable
as smoothly curved filaments (Fig. 1a).
Figure 1e shows the thin section TEM micrograph of P. aeruginosa control with a typical cell wall, outer and
cytoplasmic membrane, periplasmic space, cytoplasmic content with few electron-dense areas. The cell envelope
showed distinct layers, the most prominent being the inner cytoplasmic membrane and the outer cell envelope
membrane. The electron-dense areas at the center of the cell as filamentous densities might be interpreted as the
bacterial chromosomal DNA. Note that there is no structural evidence of chromosomal DNA exchange between close
contact bacteria, although the exchange of small portions cannot be excluded.
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Fig. 1 SEM micrograph of control P. aeruginosa a) Colony, bar 1µm; Distribution of cell size b) length and c) width; d) Rod shape
cell, bar 0.5µm; e) TEM micrograph (thin section), bar 0.2µm; f) Close up view of Fig. 1f: (1) outer membrane, (2) periplasmic
space, (3) inner membrane, (4) cytoplasm.
3.2
Example I: Artifacts
Morphological changes (peripheral cell surface, ghost formation and cell disintegration phenomena) are better observed
with TEM micrographs from collected cells deposited on grid. On the other hand, ultrastructural changes (cell wall
thickening and increase in cell wall roughness) are better studied with thin cross section micrographs. However, the thin
section technique includes chemical fixation, dehydration, heavy metal staining and embedding on resin, introducing
many artifacts; moreover, these artifacts are aggravated by long treatment times [21]. Obviously, the advantage of
viewing thin sectioned samples is the observation of ultrastructural morphology and ultrastructural changes in treated
cells. Micrographs containing artifacts related to knife marks and compression along the cutting direction (Fig. 2a),
stain contamination (Fig. 2b), rupture of carbon support film and double covered sections because of the rupture of
support film (Fig. 2c), and air bubbles (Fig. 2d) were recognized because of their extensive report in the literature [2224]. It is evident that the incorrect washing and remotion of supernatant introduce artifacts and increase the difficulty of
image interpretation (Fig. 2e and 2f).
Fig. 2 Artifacts related to a) knife marks (dotted white arrows) and compression along the cutting direction (dotted white circle),
bar 1µm; b) Stain contamination (dotted white circle), bar 1µm; c) Double-covered sections by rupture of support film, bar 1µm; d)
Air bubbles, bar 0.5µm; e) Incorrect washing with saline buffer (NaCl small crystallites), bar 1µm; f) Poor washing of L. casei
Shirota strain, 0.5µm.
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3.3
Example II: Blockage of nutrients flow, disaggregation of colonies and increase in cell wall roughness
As shown in Fig 3a, a polymeric membrane encapsules the bacterial cell surface of Staph. aureus ATCC 6538P. The
ionic interactions between the cationic polymers and negatively charged lipopolysaccharides (as lipoteichoic acid, a
component of the thick peptidoglycan layer of Gram-positive bacteria) in the outer membrane can be the responsible for
the growth inhibition and lysis, through blockage of important nutrients flow (Ca+2, Mg+2, etc.) entering the cell [25,
26]. The grape-like cluster morphology of Staph. aureus ATCC 25923 was altered as observed in Figs. 3b and 3c (black
arrows). The affinity of metal cationic particles for cell surface may be explained by the binding to positive components
of bacterial cell surface. This binding could produce the disaggregation of the exopolysaccharide matrix, separation of
cell and their rearrangement into small clusters. A small endocytotic vesicle (periplasmic space altered) and contracted
cell Staph. aureus ATCC 6538P with wrinkled cell wall are also observed at Fig. 3d. Several cells with a roughness
surface were clearly evident from Fig. 3e. Fig. 3f presents a close up view of the Fig 3e. The coccoidal shape of Staph.
aureus ATCC 6538P cells was not preserved and deformations/indentations of the cell surface can be observed. This
increase in roughness and amorphous mass could be associated with the perforation of the cell wall with release of
intracellular material and subsequent cell wall deformation and cell wall thickening [25].
Fig. 3 TEM micrographs of a) Staph. aureus ATCC 6538P treated with a cationic polysaccharide, bar 1µm; b) and c) Staph. aureus
ATCC 25923 treated with polysaccharide-metal complexes, bars 0.5µm; d-f) Thin section of Staph. aureus ATCC 6538P treated with
polysaccharide-metal complexes, bars 0.2µm.
3.4 Example III: Cell disruption with loss of intracellular material (ghost cells)
Does the extensive bacterial membrane damage is a key factor in the inactivation of bacteria by biocides? It has been
suggested that the resultant antibacterial effect is the sum of the individual inhibitory effects and in some cases
synergistic. Disruptions with release of intracellular material associated to Staph. aureus ATCC 25923 cells losing their
cytoplasm (empty and flaccid cells) were also observed in Fig 4a. Cell ghost is an empty intact cell envelope structure
devoid of cytoplasmic content including genetic material. The distortion of the physical structure of the cell could cause
the expansion and destabilization of the membrane and increase membrane fluidity, which in turn increases the passive
permeability and manifest itself as a leakage of various vital intracellular constituents, such as ions, ATP, nucleic acids,
sugars, enzymes and amino acids. In Figs. 4b and 4c, cell wall perforation with release of significant intracellular
components, projections of the cell wall and release of membrane vesicles were observed.
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Fig. 4 TEM micrographs of bacteria treated with polysaccharide-metal complexes a) Staph. aureus ATCC 25923, bar 0.2µm; b) P.
aeruginosa (thin section), bar 0.2µm; c) Staph. aureus ATCC 25923, bar 0.2µm.
3.5 Example IV: Cytolytic damage
H. pylori electron micrographs (Figs. 5a and 5b) showed an accumulation of membranous structures in the cytoplasm of
the biocides-treated cells, while no external morphological alterations were seen in the cells. The cytoplasm shows
granularity with regions where the DNA fibrils are evident. Similar granular deposits were observed in the cytoplasmic
region of E. coli cell (Figs 5c and 5d). This intracytoplasmatic coagulated material is probably resulting of the
microprecipitation of abnormal proteins or denatured membranes [27, 28]. Also, coagulated material was seen inside
the treated cells especially close to the cell’s envelope. Retraction of cytoplasm of S. Typhimurium (Fig. 5e) in high
density regions (Fig. 5f, dotted circle) in processed image (binarization) from the outer membrane is also a typical
consequence of the treatment of bacteria by metal cationic particles. The relative decrease in cell surface with respect to
cell volume plays the key role in the consequent reduction in attachment/uptake sites on the cell surface for the heavy
metals [29]. This relative reduction of the cell surface-volume ratio is an effective way for the cells to lowering the toxic
effects of environmental stress factors (SOS response) just by decreasing the attachable/exposed surface in relation to
the whole cell volume [30]. H
Fig. 5 TEM micrographs of bacteria treated with a cationic polysaccharides a) and b) H. pylori, bar 0.5µm; c) E. coli, bar 5µm; d)
Close up view of Fig 5c, bar 0.5µm; e) S. Typhimurium, bar 0.5µm; f) Processed image (binarization) obtained from Fig. 5d.
3.6 Example V: Process of morphological transformations
As observed by transmission electron microscopy, spiral form of H. pylori (Fig. 6a) was superseded by coccoid ones
(Fig. 6b). The inhibition against H. pylori was evident with the agar diffusion well assay only when there was a direct
involvement of viable L. rhamnosus cells. H. pylori changes from a spiral form to coccoid (a viable but non-culturable
form of H. pylori with reduced metabolic activity [31]) by the aggravation of its surrounding environment (such as
aerobiosis, extended incubation periods, antibiotic treatment, etc.). This process could means the species preservation
mechanism under environmental stress and not only the dying or the dormant condition of H. pylori [32]. On the other
hand, many studies have shown that the coccoid form is one of the stages of the putative H. pylori biological life cycle,
because it retains cellular structures compatible with viability and polyphosphate energy reserves [33, 34].
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Fig. 6 TEM micrographs of a) Spiral form of H. pylori, bar 0.5µm; b) Coccoid form of H. pylori after treatment with L. rhamnosus
culture, bar 0.2µm.
3.7 Example VI: Filamentation
Enlarged and filamented MRSA UA 2120 cells treated with polysaccharide-metal complexes were observed in Figs. 7a7c. It was reported that binding of some biocides to penicillin-binding proteins (PBP) 2 generally results in enlarged and
filamented cells, whereas binding to PBP 3 mainly produces cessation of septation and cluster formation and cell lysis
[25, 35]. Interactions between different types of PBPs and biocides which lead to modulation of lysis can be elucidated
by electron microscopy. Filamentation occurs when cell growth continues in the absence of cell division (mutation
and/or alteration of the stoichiometry of the cell-division components), and results in the formation of elongated
defective organisms that have multiple chromosomal copies and that present metabolic changes and damage in their
DNA [6, 25]. Accumulating evidence attributes important biological roles to filamentation in stressful environments,
including, but not limited to, sites of interaction between pathogenic bacteria and their hosts [6, 36-39].
Fig. 7 Bacteria treated with polysaccharide-metal complexes a) SEM micrograph of MRSA UA 2120, bar 1µm; b) and c) TEM
micrographs of Staph. aureus ATCC 25923, bars 2 and 1µm, respectively.
3.8 Example VII: Release of intracellular cations (cofactors)
Cations release at the cell surface can be evidenced by EDX spectra (Fig. 8a-8a.3). K+ release during exposure to
polysaccharide-metal complexes showed that the biocide components interact with cell membranes, changing their
permeability to cations K+, disorganizing the membrane and producing the cell lyses. Biocides may cause the release of
Ca2+, leading to destabilization of the membrane and inducing loss of cellular components. Ca2+ plays an important role
in cellular metabolism by binding with a variety of proteins and maintaining the lipopolysaccharide assembly on the cell
surface of Gram-negative bacteria [40-44].
Fig. 8 a) SEM micrograph of control MRSA UA 2120 treated with polysaccharide-metal complexes, bar 6 µm. (a.1-a.3) X-ray line
scan of MRSA UA 2120 after exposure to biocide.
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3.9 Example VIII: Bacteriolysis
In Figs 9a and 9b, the coccoidal shape of MRSA UA 2120 treated with polysaccharide-metal complexes was lost, and
the strain was observed as a homogeneous cell mass. Cell death is the result of the extensive loss of cell contents, the
exit of critical molecules and ions, or the initiation of autolytic processes [45]. On the other hand, the type of lysis can
be associated with the binding of biocide at differents PBPs [25]. For example, in E. coli, the killing without lysis may
arise from a modulating effect of PBP 3 inhibition on the lysis caused by inhibition of PBP 1a [46, 47]. The cooperation
between diferents PBPs may be important to exploit when considering improvements in biocides design.
Fig. 9 Bacteria treated with polysaccharide-metal complexes a) and b) TEM micrographs of MRSA UA 2120 after exposure to
biocide at 6 and 12h, respectively, bars 2 µm; c) SEM micrograph of MRSA UA 2120, bars 1 µm.
The physical bacteria-biocide interaction is a key step in all mechanism of antimicrobial activity. In this sense,
information acquired from microscopic images permits the partial understanding of bacterial physiology and
mechanisms of biocide action. Electron microscopic approaches in the study of bacteria and their physiological
consequences when it is affected by biocides permit the observation of characteristic defects of cell wall. Conventional
electron microscopy is a very useful and versatile technique to image the various structural elements in the bacteria as
well as the characterization of the morphological changes at nanometer-to-micrometer resolution.
Acknowledgements Financial support from the FONDECYT 1080704, INNOVA Bío Bío 8PC-S1-260 and DIUC 208.036.033-1.0
projects are gratefully acknowleged. J. Díaz-Visurraga is indebted to J.P. González for his experimental support. We thank S.
Céspedes and C. Retamal (Laboratory of Microbiology, University of Concepcion) for their technical assistance and also Dr Juan
Silva (University of Antofagasta) for contributing the MRSA UA 2120 specimen. We are grateful to Dr Ricardo Oliva, Alexis Stay,
Hugo Pacheco and Julio Pugin for their invaluable assistance with the electron microscopes and samples preparation. Microscopic
images were performed in the Microscopy Laboratory at the University of Concepcion.
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