Method for classifying microorganisms

Abstract

The present invention relates to a method for determining the biofilm-forming capacity of microorganisms. The present invention also relates to a method for classifying microorganisms according to the biofilm-forming capacity thereof. In particular, the present invention is useful in the fields of analysis, biological and enzymological research, pharmaceuticals, diagnostics and/or medicine. The present invention is also useful in the clinical, environmental and food-processing fields.

Claims

1. A process for determining the biofilm-producing capacity of a microorganism, comprising the following steps: a) introducing at least two particles into culture containers comprising a liquid culture medium suited to the growth of said microorganism, said particles resting on a surface submerged in the culture medium, b) introducing and independently inoculating the culture medium of the containers obtained in step a) with said microorganism at a concentration range of from 1×10.sup.−1 to 1×10.sup.−6 McFarland (McF) unit, each medium independently comprising a different concentration of microorganisms, c) maintaining the inoculated culture media in conditions which enable growth of said microorganism, d) applying a field capable of moving said at least two particles resting on a surface submerged in the culture medium, e) determining the biofilm-producing capacity of the microorganism by observing and measuring the aggregation of said particles as follows: the absence of aggregation of said particles appearing at a concentration of from 1×10.sup.−6 to 1×10.sup.−4 McF corresponding to a strongly biofilm-producing microorganism, the absence of aggregation of said particles appearing from a concentration of greater than 1×10.sup.−4 and less than 1×10.sup.−2 McF corresponding to a moderately biofilm-producing microorganism, the absence of aggregation of said particles appearing from a of greater than or equal to 1×10.sup.−2 McF corresponding to a weakly biofilm-producing microorganism, an aggregation of said particles, regardless of the concentration, corresponding to a non-biofilm-producing microorganism.

2. The process as claimed in claim 1, wherein the process comprises a step e′) for determining the Biofilm Index (BFI) value for each of the media by: analyzing red, green and blue color components of pixels of an image formed by observing the particles, making it possible to determine a particle density per pixel, calculating the correlation between the particles, calculating the aggregation density of the particles expressed as BFI, and determining the biofilm formation potential index per container (BPc) according to the following formula (I):
BPc=[1−(BFIe/BFIn)]  (I) wherein BFIe corresponds to the biofilm formation index value in the inoculated medium and BFIn corresponds to the biofilm formation index value in a container which does not comprise microorganisms (negative control).

3. The process as claimed in claim 2, comprising a step e″) subsequent to or simultaneous with the step e′) for determining the threshold value (S) according to the following formula (II):
S=1−[(mBFIn−3×stmBFIn)/2]/mBFIn  (II) wherein mBFIn is equal to the mean of the BFIs of the negative controls, stmBFIn is equal to the calculated standard deviation of the mean of the BFIs of the negative controls.

4. The process as claimed in claim 3, wherein the biofilm-producing capacity of the microorganism is determined by comparing the BPc value as a function of the concentration of microorganisms, as follows: a BPc value greater than or equal to the threshold value S from a concentration of microorganisms of from 1×10.sup.−6 to 1×10.sup.−4 McF corresponding to a strongly biofilm-producing microorganism, a BPc value greater than or equal to the threshold value S from a concentration of microorganisms of greater than 1×10.sup.−4 and less than 1×10.sup.−2 McF corresponding to a moderately biofilm-producing microorganism, a BPc value greater than or equal to the threshold value S from a concentration of microorganisms of 1×10.sup.−2 McF corresponding to a weakly biofilm-producing microorganism, a BPc value less than the threshold value S, regardless of the concentration, corresponding to a non-biofilm-producing microorganism.

5. A process for classifying microorganisms, comprising the following steps: a) introducing at least two particles into culture containers comprising a liquid culture medium suited to the growth of said microorganism, said particles resting on a surface submerged in the culture medium, b) introducing and independently inoculating the culture medium of the containers obtained in step a) with said microorganism at a concentration range of from 1×10.sup.−1 to 1×10.sup.−6 McF unit, each medium independently comprising a different concentration of microorganisms, c) maintaining the inoculated culture media in conditions which enable growth of said microorganism, d) applying a field capable of moving said at least two particles resting on a surface submerged in the culture medium, with the aim of obtaining an aggregation, in spot form, of said at least two particles, e) classifying the microorganism by observing the aggregation of said particles as follows: category I: absence of aggregation of said particles appearing at a concentration of from 1×10.sup.−6 to 1×10.sup.−4 McF category II: absence of aggregation of said particles appearing from a concentration of greater than 1×10.sup.−4 and less than 1×10.sup.−2 McF corresponding to a moderately biofilm-producing microorganism, category III: absence of aggregation of said particles appearing from a concentration of greater than or equal to 1×10.sup.−2 McF corresponding to a weakly biofilm-producing microorganism, category IV: aggregation of said particles, regardless of the concentration, corresponding to a non-biofilm-producing microorganism.

6. The process as claimed in claim 1, comprising six culture containers in which the concentration of microorganisms at step b) is respectively 1×10.sup.−6 McF, 1×10.sup.−5 McF, 1×10.sup.−4 McF, 1×10.sup.−3 McF, 1×10.sup.−2 McF and 1×10.sup.−1 McF.

7. The process as claimed in claim 1, wherein the inoculated culture media are maintained in conditions which enable growth of said microorganism for 1 to 8 hours.

8. The process as claimed in claim 1, wherein the inoculated culture media are maintained in conditions which enable growth of said microorganism for 5 to 12 generation times of said microorganism.

9. The process as claimed in claim 1, wherein said at least two particles are electrically charged, magnetic or magnetizable particles, or particles covered with at least one magnetic or magnetizable layer.

10. The process as claimed in claim 1, wherein said at least two particles are subjected to an electromagnetic field.

11. The process as claimed in claim 1, wherein said at least two particles are subjected to a gradual increase in the electromagnetic field.

12. The process as claimed in claim 1, wherein said magnetic field is generated by moving field generating means.

13. The process as claimed in claim 1, wherein said at least two particles are illuminated by means of a light source, to detect movement thereof.

14. The process as claimed in claim 1, wherein said at least two particles generate a signal.

15. The process as claimed in claim 14, wherein said at least two particles are excitable, fluorescent or phosphorescent or radioactive or chemiluminescent.

16. The process as claimed in claim 1, wherein one of said containers does not comprise said microorganism, said container representing a control for the medium.

Description

BRIEF DESCRIPTION OF THE FIGURES

(1) FIG. 1 is a photograph of a 96-well plate. It shows biofilm formation on 96-well polystyrene plates by Gram-negative bacteria: A) P. aeruginosa (Pa), B) K. pneumoniae (Kp) and C) R. mannitolilytica, isolated from the IFO hospital. The images were obtained after magnetization of the plates on the test block and digitization with the plate reader. The reference laboratory strains are indicated in bold. The absence of biofilm formation is revealed by the presence of the central black dot in the wells, corresponding to the coming together of the particles. The presence of biofilm is revealed by the absence of a black dot at the center of the wells. The wells corresponding to negative controls (Ctr (−)) containing solely BHI medium and the magnetic microparticles correspond to the bottom row of wells.

(2) FIG. 2 is a photograph of a 96-well plate. It shows biofilm formation on 96-well polystyrene plates for Gram-positive bacteria: A) S. aureus (SA), B) S. epidermidis (SE) and C) different Gram-positive bacteria isolated at the IFO hospital. The images were obtained after magnetization of the plates on the test block and digitization with a plate reader. The reference laboratory strains are indicated in bold. The absence of biofilm formation is revealed by the presence of the central black dot in the wells, corresponding to the coming together of the particles. The presence of biofilm is revealed by the absence of a black dot at the center of the wells. The wells corresponding to negative controls (Ctr (−)) containing solely BHI medium and the magnetic microparticles are outlined in red.

(3) FIG. 3 is a bar chart representing the percentage of clinical isolates of each species, classified as a function of their biofilm-producing capacity measured by the process, namely strongly producing (dark gray zone), moderately producing (white zone), weakly producing (light gray zone) and non-producing/non-adhering (black zone).

(4) FIG. 4 is a bar chart representing the results of the quantitative analysis of biofilm formation by crystal violet (CV) staining of clinical isolates of P. aeruginosa, K. pneumoniae, R. mannitolilytica, S. aureus, S epidermidis and other Gram-positive bacteria. The isolates were classified as a function of their threshold values during optical density (OD) measurements, at a wavelength of 570 nm (OD570), of the medium. When the OD570 value is <0.18, the isolates are non-producing/non-adhering, when the OD570 is between 0.18 and 0.37 they are weakly producing; when the OD570 is between 0.37 and 0.74 they are moderately producing and when the OD570 is greater than 0.74 they are strongly producing. The error bars indicate the standard error. Dashed lines (- - -) indicate the threshold (cut-off) values at OD570<0.37 and at OD570<0.74.

(5) Other advantages may also become apparent to those skilled in the art on reading the examples below, illustrated by the appended figures and given by way of illustration.

EXAMPLES

Example 1: Evaluation of the Microorganism Biofilm-Producing Capacity

(6) In the example below, an example of the process according to the disclosure is especially denoted characterization process or cBFRT process. In this example, the materials and methods used were the following:

(7) Strains and Growth Conditions

(8) Reference strain: Staphylococcus aureus American Type Culture Collection (ATCC) 25923, Staphylococcus aureus ATCC 6538, Staphylococcus epidermidis ATCC 14990, Staphylococcus epidermidis ATCC 12228, Klebsiella pneumoniae ATCC 700603, Klebsiella pneumoniae ATCC 13883, Pseudomonas aeruginosa ATCC 47085, Pseudomonas aeruginosa ATCC 9027, Pseudomonas aeruginosa PA14, Ralstonia mannitolilytica LMG 6866, Ralstonia mannitolilytica BK931 and the clinical isolates of bacteria were cultured aerobically on a nonselective agar (blood agar, chocolate agar, MacConkey agar) (Oxoid, Hampshire, United Kingdom) at 37° C.

(9) Selection of the Isolates

(10) A total of 52 clinical isolates collected from patients suffering from nosocomial infections, admitted to the IFO hospital in Rome, were evaluated. The bacteria were collected from different materials including chronic infections (ulcers), intravenous and urinary catheters, blood, urine, sputum and naso-bronchial lavage samples. An ulcer was classified as chronic if it had existed for at least 3 months (Dissemond, 2006). The study also comprised strains having an increased resistance to commonly used antibiotics, as determined according to the standard for antibiotic susceptibility tests by the VITEK2 system (bioMérieux, Florence, Italy). In particular, the strains of P. aeruginosa resistant to three or more classes of antibiotics were considered to be multidrug-resistant (MDR) microorganisms. The K. pneumoniae strains resistant to the majority of the beta-lactam antibiotics, including penicillins, cephalosporins, and aztreonam monobactam, growing on selective chromogenic medium chromID ESBL (bioMérieux, Florence, Italy) were classified as having extended-spectrum beta-lactamase (ESBL). The phenotypic detection of ESBL producers was further defined using the disk test (Oxoid, United Kingdom), as described in De Gheldre, 2003; Moremi, 2014. The K. pneumoniae strains resistant to carbapenems and identified by a selective chromogenic medium chromID CARBA (bioMérieux, Florence, Italy) are referenced as K. pneumoniae carbapenemase (KPC)-producing. The methicillin-resistant Staphylococcus aureus (MRSA) were penicillin binding protein (PBP)-producing after confirmation by the PBP2′ latex agglutination test kit (Oxoid Ltd, Basingstoke, United Kingdom). The vancomycin-resistant enterococci (VRE) were identified by susceptibility tests and confirmed by selective chromogenic media chromID VRE (bioMérieux, Florence, Italy). The levels of susceptibility to antibiotics were compared with those defined by the minimum inhibitory concentration (MIC) according to the interpretation criteria recommended by the European Committee on Antimicrobial Susceptibility Testing (EUCAST). The number of species and the characteristics of the strains tested are given in table 2. The isolates were stored at −70° C. in Cryobank tubes (Copan Italia SpA) and cultured overnight at 37° C. on a specific agar dish before the test.

(11) Preparation of the Inoculum

(12) The biofilm formation was evaluated by the BioFilm Ring Test (BFRT) (BioFilm Control, Saint-Beauzire, France), using the commercially available kit (BioFilm Control, Saint Beauzire, France). The toner solution (TON004) containing magnetic beads was mixed in a brain heart infusion medium (BHI, Difco, Detroit, Mich., USA) according to the manufacturer's instructions. In order to determine the biofilm-producing capacity/classification of the microorganisms, the inoculum was prepared as follows. A fresh culture of bacteria on agar, cultured overnight, was used for each strain to be tested. The cultured bacteria were then transferred by a sterile inoculation loop into a sterile tube containing 2 ml of 0.45% saline solution, at the equivalent to 1.0±0.3 McFarland turbidity standard (McF) and mixed thoroughly. Subsequently, 200 μl of the inoculum were transferred into the wells of a 96-well polystyrene plate. From this initial inoculum, serial dilutions by a factor of 10 were performed, from 1×10.sup.−1 to 1×10.sup.−6 McF, by transferring 20 μl of the microbial solution in 200 μl of the BHI/TON mixture.

(13) One or more laboratory strains were included in each plate as standard reference and quality control. One well containing the BHI/TON mix without microbial cells was used as negative control in each experiment.

(14) After five hours of incubation at 37° C. without stirring (static culture), the wells were covered with a few drops of contrast liquid (inert opaque oil used for the reading step, included in the BFRT kit), placed for 1 min on the block carrying 96 mini magnets (test block) and scanned with a specially designed plate reader (BIOFILM pack, BioFilm Control, Saint Beauzire, France). The adhesion strength of each strain was expressed as biofilm formation index (BFI), which was calculated by specialized software initially described in Chavant, 2007. The BFI values were used to measure the biofilm formation potential (BP), using the formula:
BPc=[1−(BFIe/BFIn)]  (I)

(15) wherein BFIe corresponds to the biofilm formation index value in the inoculated medium and BFIn corresponds to the biofilm formation index value in a container which does not comprise microorganisms.

(16) The threshold value (S) was calculated according to the following formula (II)
S=1−[(mBFIn−3×stmBFIn)/2]/mBFIn  (II)

(17) wherein mBFIn is equal to the mean of the BFIs of the negative controls, stmBFIn is equal to the calculated standard deviation of the mean of the BFIs of the negative controls.

(18) In this example, the BFIn values were from 18 to 20. The calculated value of S was equal to 0.53.

(19) BPc values above the calculated S value were considered as biofilm-producing. Thus, the final dilution for which the value is above the calculated S value made it possible to identify the biofilm-forming capacity of the microorganism. Consequently, the microorganisms were classified in the following categories: non-adhering cells, weakly biofilm-producing, moderately biofilm-producing and strongly biofilm-producing. Each microbial culture was analyzed in duplicate, and the experiments were repeated at least 3 times for each strain in order to evaluate the reproducibility, accuracy and precision of the measurement. The values were considered to be valid if the standard deviation between the duplicates was less than 8%. The replicas showed complete agreement in the classification/categorization of the microorganisms.

(20) Evaluation of Biofilm Formation with the Crystal Violet Assay

(21) 96-well polystyrene plates were inoculated with 200 μl of an initial bacterial suspension (10.sup.5 CFU/ml) in BHI medium and incubated at 37° C. for 24 and 48 hours without stirring. Each condition was carried out in triplicate. The medium was removed from the wells, which were washed 3 times with 200 μl of sterile distilled water. The plates were air-dried for 45 minutes and the adhering cells were stained with 200 μl of 0.1% crystal violet solution. After 20 minutes, the stain was eliminated and the wells were washed four times with 300 μl of sterile distilled water to eliminate the excess stain. The stain incorporated by the biofilm-forming cells was dissolved with 200 μl of an 80/20% ethanol/acetone mixture and the absorbance of each well was measured by spectrophotometry at 570 nm (OD.sub.570) using the automated PhD Ix™ system (Bio-Rad Laboratories, Hercules, Calif., USA).

(22) For the comparative analysis, OD.sub.570 values were use to semi-quantitatively classify the biofilm production for the bacterial strains, according to the process described in Stepanovic et al (Stepanovic, 2000). Briefly, the threshold optical density (ODc) was defined as three standard deviations above the mean optical density (OD) of the negative control. Thus, the strains were classified as follows: OD<ODc=non-adhering; ODc<OD<2×ODc=weakly biofilm-producing; 2×ODc<OD<4×ODc=moderately biofilm-producing; and OD>4×ODc=strongly biofilm-producing.

(23) All the assays were carried out in triplicate, with reference strains and clinical isolates tested three times over to evaluate any potential variations in the conditions for assaying the biofilm.

(24) Statistical Methods

(25) The kappa coefficient test was used to determine the agreement between the results obtained with the characterization process and crystal violet (CV). The agreement was calculated according to Viera (Viera, 2005): kappa=0.81-1, very good; kappa=from 0.61 to 0.80, good; kappa 0.41 to 0.60, moderate; kappa=0.21 to 0.40, fair; kappa≤0.20, poor. The results of biofilm formation obtained with the two methods were compared also using McNemar's test. Values of p<0.05 were considered to be significant.

(26) Results and Discussion

(27) Standardization of the Bacterial Inoculums to McFarland Standards

(28) Given that different microorganisms, including mucoid or non-mucoid strains, or other characteristics such as the size, can affect the measurement of initial cellular concentration and consequently the reliability of the assay, the precision of the initial inoculum was measured with a densitometer and verified by counting the colony forming units (CFUs) (Welch, 2012).

(29) The data reported in table 1 showed that one McF unit varied from 0.6×10.sup.9 CFU/ml for R. mannitolilytica to 1.4×10.sup.9 CFU/ml for P. aeruginosa. This result demonstrates that measuring the initial inoculum with the densitometer does not generate significant differences. The mean value of the CFU/ml for the different bacterial strains from 1 McF corresponds to 1.0×10.sup.9±3.6×10.sup.8, which is in the same range of values as indicated previously for bacteria (Eng et al, 1984). Consequently, the CFU values show that the McF standard is highly reproducible, independently of the bacterial species or microbial phenotypes, ensuring the precision of the assay at the moment of the inoculum.

(30) TABLE-US-00001 TABLE 1 CFU/ml correspondence to 1 McFarland for the bacteria used to inoculate the cBFRT assay. The values represent the mean CFU/ml ± SD of two repetitions. Microbial strain CFU/ml P. aeruginosa 1.4 × 10.sup.9± 7.2 × 10.sup.8 K. pneumoniae 0.9 × 10.sup.9 ± 7.9 × 10.sup.8 R. mannitolilytica 0.6 × 10.sup.9 ± 2.6 × 10.sup.8 S. aureus 1.3 × 10.sup.9 ± 8.4 × 10.sup.8 S. epidermidis 1.1 × 10.sup.9 ± 7.1 × 10.sup.8 Other Gram+ 1.1 × 10.sup.9 ± 8.7 × 10.sup.8 Mean 1.0 × 10.sup.9 ± 3.2 × 10.sup.8

(31) Evaluation of Biofilm Production in Gram-Negative and Gram-Positive Bacteria

(32) The biofilm-forming ability was initially evaluated on isolates of P. aeruginosa, which is a Gram-negative opportunistic pathogen with a noteworthy biofilm-forming capacity, which ensures its persistence in the environment and in chronic infectious diseases (Høiby et al, 2010).

(33) Among the 12 strains tested, three were laboratory strains with a known biofilm-forming capacity. These comprised, respectively, Pa47085 (moderate capacity) (Schaber, 2004), PA14 (weak capacity) (Rahme, 1995), and Pa9027 (strong capacity), (Stapleton, 1993), while the 9 remaining strains of P. aeruginosa were clinical isolates collected from hospitalized patients (table 2).

(34) TABLE-US-00002 TABLE 2 Characteristics of the clinical isolates used in this study. Multidrug-resistant (MDR), having extended-spectrum beta- lactamase (ESBL), K. pneumoniae carbapenemase (KPC)-producing, methicillin-resistant S. aureus (MRSA), vancomycin-resistant enterococci (VRE). Catheter-associated urinary tract infections (CA-UTI), central venous catheter (CVC), catheter-associated blood infection (CA-BI). Bacterial Clinical species isolates Phenotype Site of isolation Gram-negative bacteria P. aeruginosa 9 MDR (2) CA-BI (2) MDR (1) Chronic ulcer (2) — Wound (1) — Urine (1) — Respiratory (3) K. pneumoniae 8 KPC (1) CVC (1) — Blood (2) — Wound (1) ESBL (1) CA-UTI (2) — Respiratory (2) R. mannitolilytica 8 — Blood (8) Gram-positive bacteria S. aureus 10 MRSA (1) CVC (1) MRSA (3) Chronic ulcer (6) — Skin (2) — Respiratory (1) S. epidermidis 8 — Blood (6) — Wound (2) Other Gram+ 9 — Blood (2) VRE (1) Wound (6) — Urine (1) Gram-negative bacteria P. aeruginosa 9 MDR (2) CA-BI (2) MDR (1) Chronic ulcer (2) — Wound (1) — Urine (1) — Respiratory (3) K. pneumoniae 8 KPC (1) CVC (1) — Blood (2) — Wound (1) ESBL (1) CA-UTI (2) — Respiratory (2) R. mannitolilytica 8 — Blood (8) Gram-positive bacteria S. aureus 10 MRSA (1) CVC (1) MRSA (3) Chronic ulcer (6) — Skin (2) — Respiratory (1) S. epidermidis 8 — Blood (6) — Wound (2) Other Gram+ 9 — Blood (2) VRE (1) Wound (6) — Urine (1)

(35) After five hours of incubation, the reference strain PA14 immobilized the magnetic beads only at the highest concentration of cells (McF=1×10.sup.−2), which suggests a weak capacity for adhesion and qualifies it as weakly biofilm-producing (FIG. 1). Similarly, PA14 was classified as weakly biofilm-producing (table 3A). Pa47085 adheres more readily to the surface of the wells and blocks the magnetic beads at a concentration of McF=1×10.sup.−3, thereby identifying this strain as moderately biofilm-producing. Similarly, Pa9027 was confirmed as being strongly biofilm-producing at a cellular concentration of McF=1×10.sup.−6. These results are in agreement with those described previously for these laboratory strains (Schaber, 2004; Rahme, 1995; Stapleton, 1993).

(36) Among the clinical isolates, six strains proved to be strongly biofilm-producing. In particular, this includes the three MDR strains, including two originating from patients suffering from catheter-associated blood infections (Pa6020-IFO, Pa5252-IFO), chronic ulcers (Pa5797-IFO), and the strains isolated from bronchioalveolar lavage from a patient suffering from cystic fibrosis (Pa0629-IFO), from pleural fluid (Pa5291-IFO) and originating from a patient with an infected wound (Pa0118-IFO). Two strains were classified as moderately biofilm-producing, deriving from a patient suffering from a chronic ulcer (Pa3019-IFO) and from another bronchoalveolar lavage from a patient suffering from cystic fibrosis (Pa0628-IFO). Conversely, the only weakly biofilm-producing strain was a mucoid strain isolated from a urinary infection (Pa0115-IFO).

(37) The assay was then use to evaluate the biofilm-forming capacity of Klebsiella pneumoniae, one of the most significant nosocomial pathogens (Podschun, 1998). The results, summarized in FIG. 1B and in table 3B, show that the reference strains Kp700603 and Kp13883 were, respectively, non-adhering/biofilm producing and moderately biofilm-producing, confirming previous reports (Naparstek, 2014). Regarding the clinical strains Kp0068-IFO (urinary infection), Kp5553-IFO (an ESBL strain from a urinary sound), Kp5668-IFO (a KPC strain from a central venous catheter) and Kp5776-IFO (from pleural fluid), they were found to be moderately biofilm-producing. Conversely, the strains isolated from blood cultures (Kp5656-IFO, Kp5281-IFO, Kp5783-IFO, Kp3040-IFO) but not originating from catheter-associated bacterial blood infections, exhibited a mucoid phenotype and were deemed to be non-adhering and non-biofilm-producing, This data is in agreement with previous reports indicating a higher frequency of biofilm-forming strains in K. pneumoniae isolated from non-fluid physiological environments, which may contribute to explaining the difficulty in eradicating these infections once they are established in solid tissues (Sanchez, 2013). Another important observation emerging from the limited number of clinical isolates analyzed was the relatively high incidence of non-adhering strains. More precisely, we observed that only 50% of the K. pneumoniae included in this study were capable of producing biofilm. This value is in agreement with previous in vitro studies which demonstrate that only 40% of K. pneumoniae isolated from different materials were able to produce biofilm (Yang, 2008).

(38) TABLE-US-00003 TABLE 3 Analysis of the plates obtained with the characterization process for A) P. aeruginosa, B) K. pneumoniae and C) R. mannitolilytica. A) Pa6020- Pa0115- Pa3019- Pa5797- Pa0628- Pa0629- Pa5252- Pa0118- Pa5291- McF Pa47085 Pa14 Pa9027 IFO IFO IFO IFO IFO IFO IFO IFO IFO 1 × 10.sup.−1 0.97 0.98 0.99 0.98 0.90 0.97 0.99 1.00 1.00 0.97 0.98 1.00 Non-adhering/ weak 1 × 10.sup.−2 0.97 0.72 0.99 0.96 0.33 0.99 1.00 1.00 1.00 0.95 0.97 1.00 Weak 1 × 10.sup.−3 0.96 0.01 0.95 0.92 0.12 0.98 0.99 0.98 0.98 0.98 0.93 1.00 Moderate 1 × 10.sup.−4 0.01 −0.02 0.97 0.74 0.06 0.64 0.87 0.58 0.96 0.96 0.89 1.00 Moderate 1 × 10.sup.−5 0.03 0.03 0.96 0.57 0.07 −0.02 0.53 0.07 0.87 0.92 0.78 1.00 Strong 1 × 10.sup.−6 0.05 0.05 0.52 0.39 0.08 0.00 0.49 −0.01 0.71 0.87 0.56 0.93 Strong 1 2 3 4 5 6 7 8 9 10 11 12 B) Kp5553- Kp5776- Kp5668- Kp5281- Kp3040- Kp0068- Kp5656- Kp5783- McF Kp13883 Kp700603 IFO IFO IFO IFO IFO IFO IFO IFO 1 × 10.sup.−1 0.07 0.98 0.92 0.86 1.00 0.01 0.04 1.00 −0.01 0.15 Non-adhering/ weak 1 × 10.sup.−2 0.08 0.94 0.91 0.81 0.94 0.01 0.04 1.00 0.00 0.05 Weak 1 × 10.sup.−3 0.03 0.82 0.84 0.76 0.95 −0.02 0.04 0.99 −0.01 0.00 Moderate 1 × 10.sup.−4 0.07 0.22 0.37 0.26 0.92 0.01 0.06 0.75 0.04 0.02 Moderate 1 × 10.sup.−5 −0.01 −0.03 0.05 0.00 0.25 0.02 0.00 0.01 0.04 −0.01 Strong 1 × 10.sup.−6 0.00 0.02 0.02 0.02 0.11 0.04 0.01 −0.05 0.03 0.01 Strong 1 2 3 4 5 6 7 8 9 10 C) LMG Rm1- Rm2- Rm3- Rm4- Rm5- Rm6- Rm7- Rm8- McF 6866 BK931 IFO IFO IFO IFO IFO IFO IFO IFO 1 × 10.sup.−1 0.88 0.66 0.88 0.99 0.59 0.57 0.91 0.51 0.57 0.94 Non-adhering/ weak 1 × 10.sup.−2 0.26 0.14 0.17 0.61 0.13 0.11 0.16 0.12 0.15 0.54 Weak 1 × 10.sup.−3 0.09 0.03 0.10 0.13 0.05 0.07 0.07 0.10 0.09 0.15 Moderate 1 × 10.sup.−4 0.00 0.04 0.07 0.01 0.11 0.12 0.02 0.10 0.10 0.04 Moderate 1 × 10.sup.−5 0.02 0.06 0.06 0.10 0.09 0.12 0.02 0.09 0.10 0.10 Strong 1 × 10.sup.−6 0.03 0.10 0.05 0.03 0.09 0.13 0.01 0.09 0.11 0.09 Strong 1 2 3 4 5 6 7 8 9 10

(39) In the table above, the bacteria were classified as a function of the BP value, also referred to as BPc, measured. The cut-off/threshold was established at 0.53. A value of between 1 and 0.53 corresponds to biofilm formation, a value of less than 0.53 to a possible absence of biofilm, or a non-significant biofilm formation.

(40) R. mannitolilytica is rarely isolated from clinical samples and represents the most widespread species of the genus Ralstonia found in patients suffering from cystic fibrosis (Coenye, 2005). R. mannitolilytica has been associated with epidemics around the world (Ryan, 2014) and the hypothesis of the biofilm-forming capacity of this bacterium has been raised numerous times as a strategy for survival in difficult environments and in clinical settings, although it has rarely been tested. The present strains analyzed were isogenic clinical isolates from an epidemic occurring in the oncology department in the IFO hospital in 2014. Two reference strains, R. mannitolilytica LMG 6866 (De Baere, 2001) and R. mannitolilytica BK931 (Marroni, 2003), were evaluated in comparison, although their biofilm-producing capacity is unknown. The results from FIG. 10 and table 3C revealed that the reference strains LMG 6866 and BK931 were weakly biofilm-producing. Similarly, the 8 strains of R. mannitolilytica, isolated from blood culture, were weakly biofilm-producing/non-adhering.

(41) Subsequently, the biofilm-forming ability was studied for different Gram-positive bacteria. More precisely, strains of Staphylococcus aureus, which have been associated with different types of human infections, comprising skin infections, endocarditis, bone infections, septic shock and biofilm-associated chronic infections (Otto, 2008) were evaluated. The reference strains Sa6538 and Sa25923, known to form biofilms (Latimer, 2012; Croes, 2009), were confirmed as strongly biofilm-producing and moderately biofilm-producing (FIG. 2A and table 4a). Among the clinical isolates, five strains were deemed to be moderately biofilm-producing (Sa3074-IFO, Sa3050-IFO, Sa0186-IFO, Sa5674-IFO and Sa5826-IFO) and three strains isolated from patients suffering from chronic ulcers were classified as strongly biofilm-producing (Sa3146-IFO, Sa3079-IFO and Sa3065-IFO). Among the four MRSA strains, two were moderately biofilm-producing (Sa0186-IFO, Sa5826-IFO) and two were strongly biofilm-producing (Sa3146-IFO and Sa3065-IFO). The only two weakly biofilm-producing strains originated from children suffering from mild atopic dermatitis (Sa3032-IFO and Sa0073-IFO). None of the strains analyzed was classified as non-adhering or non-producing. This result is in agreement with previous reports describing the effectiveness of S. aureus in producing biofilm (Otto, 2008; Periasamy, 2012).

(42) Staphylococcus epidermidis, a major component of the normal microbial population of human skin, is a significant nosocomial pathogen in patients with predisposing factors such as a probe or an implanted device (Otto, 2009). While S. epidermidis has a weak pathogenic potential, it has been isolated in 40% of bacterial blood infections (Suetens, 2007). This high occurrence probably originates from the fact that it is a ubiquitous colonizer of human skin, and consequently is a possible source of contamination for devices when they are inserted or removed (Otto, 2009). Nonetheless, the biofilm-forming capacity of S. epidermidis is considered to be the most relevant determinant of its virulence (O'Gara, 2001; Götz, 2002; Cerca, 2005). In our assays, the reference strains Se12228 (Zhang, 2003) and Se14990 (Stepanovic, 2003) were classified, respectively, as weakly biofilm-producing and moderately biofilm-producing (FIG. 2B and table 4B). These results are in agreement with the data available in the literature. The analysis of the adhesion capacity by the assay revealed that 5 of the 6 clinical isolates derived from blood cultures (Se5287-IFO, Se5669-IFO, Se5934-IFO, Se5752-IFO and Se5845-IFO), were weakly biofilm-producing. One strain isolated from a catheter-associated blood infection (Se5993-IFO) proved to be moderately biofilm-producing. The two other strains of S. epidermis, isolated from wounds, were both classified as moderately biofilm-producing.

(43) TABLE-US-00004 TABLE 4 analysis of the plates obtained with the process A) S. aureus B) S. epidermidis and C) various Gram-positive bacteria. A) Sa3074- Sa3032- Sa3050- Sa0073- Sa0186- Sa5674- Sa5826- Sa3146- Sa3079- Sa3065- McF Sa6538 Sa25923 IFO IFO IFO IFO IFO IFO IFO IFO IFO IFO 1 × 10.sup.−1 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 Non-adhering/ weak 1 × 10.sup.−2 0.99 0.99 0.98 0.99 0.94 1.00 1.00 0.99 0.99 0.98 0.99 0.99 Weak 1 × 10.sup.−3 0.95 0.99 0.92 0.09 0.84 0.44 0.98 0.99 0.98 0.98 0.99 0.97 Moderate 1 × 10.sup.−4 0.05 0.94 0.11 0.10 0.08 0.04 0.56 0.45 0.33 0.98 0.97 0.98 Moderate 1 × 10.sup.−5 0.09 0.54 0.12 0.03 0.10 0.04 0.05 0.02 0.02 0.93 0.74 0.99 Strong 1 × 10.sup.−6 0.03 0.01 0.03 0.04 0.05 0.03 0.00 0.03 −0.01 0.70 0.64 0.80 Strong 1 2 3 4 5 6 7 8 9 10 11 12 B) Se5287- Se5669- Se5899- Se1501- Se5934- Se5752- Se5845- Se5993- McF Se12228 Se14990 IFO IFO IFO IFO IFO IFO IFO IFO 1 × 10.sup.−1 0.99 1.00 0.94 0.97 0.99 0.98 0.97 0.99 0.99 1.00 Non-adhering/ weak 1 × 10.sup.−2 0.59 0.99 0.62 0.98 0.99 0.91 0.84 0.99 0.99 0.98 Weak 1 × 10.sup.−3 0.29 0.81 0.27 0.28 0.64 0.53 0.45 0.46 0.34 0.98 Moderate 1 × 10.sup.−4 0.20 0.27 0.20 0.00 0.05 0.20 0.16 0.12 0.05 0.87 Moderate 1 × 10.sup.−5 0.13 0.28 0.13 0.01 0.03 0.02 0.03 0.01 0.02 0.44 Strong 1 × 10.sup.−6 0.01 0.02 0.03 0.06 0.02 0.02 0.05 0.03 0.05 0.01 Strong 1 2 3 4 5 6 7 8 9 10 C) Ss5800- Sh5529- Sh5592- Sag0140- Sag0093- Efm5304- Efm5515- Efs0044- Efs5269- McF Sa6538 IFO IFO IFO IFO IFO IFO IFO IFO IFO 1 × 10.sup.−1 1.00 0.87 0.96 1.00 0.93 1.00 0.62 0.94 0.98 0.07 Non-adhering/ weak 1 × 10.sup.−2 0.99 0.21 0.83 0.90 0.97 1.00 0.48 0.83 0.97 0.06 Weak 1 × 10.sup.−3 0.94 0.13 0.35 0.64 0.91 0.95 0.12 0.62 0.95 0.07 Moderate 1 × 10.sup.−4 0.06 −0.06 0.07 0.12 0.87 0.81 −0.02 0.15 0.39 0.05 Moderate 1 × 10.sup.−5 0.04 0.14 0.06 0.01 0.22 0.63 −0.03 0.05 −0.04 0.05 Strong 1 × 10.sup.−6 0.01 0.04 0.04 0.03 0.24 0.52 0.00 0.06 0.03 0.06 Strong 1 2 3 4 5 6 7 8 9 10

(44) Other Gram-positive bacteria, recognized as significant nosocomial pathogens, were analyzed. The S. aureus strain Sa6538 was used as reference strain and the bacteria analyzed included: Staphylococcus haemolyticus, Streptococcus sanguinis, Streptococcus agalactiae, Enterococcus faecium and Enterococcus faecalis. More precisely, the strains of S. agalactiae (Sag0140-IFO and Sag0093-IFO) isolated from infected ulcers were found to be moderately/strongly biofilm-producing (FIG. 2C and table 4C). E. faecium (Efm5304-IFO) was classified as weakly biofilm-producing while the strain of ERV (Efm5515-IFO) was identified as moderately producing (FIGS. 3C and 4C).

(45) Analysis of the Species-Specific Distribution of the Biofilm Production Phenotype

(46) The analysis of biofilm production as a function of the different bacterial species indicates that, among the Gram-negative bacteria, P. aeruginosa had the most consistent biofilm-producing phenotype. In fact, 6 strains (67%) were strongly biofilm-producing, 2 (22%) were moderately biofilm-producing and 1 (11%) was weakly biofilm-producing, respectively (FIG. 3). Conversely, 4 (50%) of the K. pneumoniae isolates were moderately biofilm-producing and 4 strains (50%) proved to be non-adhering/non-producing bacteria. Interestingly, the two clinical isolates and the laboratory strains of R. mannitolilytica had the weakest capacity for adhesion. Among the Gram-positive bacteria, 3 (20%) of the S. aureus strains were strongly biofilm-producing, 5 (50%) were moderately biofilm-producing and 2 were deemed to be weakly biofilm-producing (20%). In the group of clinical isolates of S. epidermis, 6 strains had a weakly biofilm-producing phenotype (75%), while the 2 other strains (25%) were moderately biofilm-producing. Overall, the analysis of the 52 clinical isolates revealed that more than 44% ( 23/52) were strongly/moderately biofilm-producing and 85% ( 44/52) were capable of producing biofilm. Despite the fact that the distribution of these 52 isolates does not reflect clinical reality, this value is in correlation with the 80% estimated by Romling and Balsalobre (Romling, 2012; NIH Parent Grant Announcement, 2002). The most relevant results were obtained with P. aeruginosa (89%) and S. aureus (80%) as the most effective biofilm producers (FIG. 3).

(47) It is interesting to note that the multidrug-resistant strains all proved to belong to the strongly/moderately biofilm-producing groups. Indeed, as indicated previously, multidrug-resistant organisms are more frequently associated with strong biofilm production (Kwon, 2008; Rao, 2008; Sanchez, 2013). This point appears to represent a key element which may promote antimicrobial resistance by selecting highly resistant strains exposed to subinhibitory antimicrobial concentrations and by offering favorable conditions for gene transfer (Wang, 2010).

(48) Categorization with Crystal Violet (CV) Staining

(49) The biofilm production phenotype of the clinical isolates was also evaluated by CV assay. The mean light absorbance for the different biofilm-producing bacteria obtained by analysis with CV is represented in FIG. 6. The results revealed that the reproducibility of the CV assay was generally good with minor differences observed between the means of the results of the repetitions. In the clinical isolates and the reference strains, the biofilm formation was heterogeneous according to the bacterial species and the isolates from the same species.

(50) The results of the CV assay were subsequently compared with the data collected by the process according to the disclosure. Complete agreement was defined as the percentage of isolates which were in the same category with both methods; agreement was considered to be partial when the process according to the disclosure obtained the same classification as the CV OD570±standard deviation. The isolates in disagreement were considered to be inconsistent.

(51) Complete and partial agreement between the characterization process and the CV assay for P. aeruginosa were 83% and 92%, corresponding to two and one disagreements, respectively. For K. pneumoniae, the characterization process showed partial precision of 70% compared to CV staining, with three disagreements. In detail, two strains classified as being in disagreement by the characterization process (Kp3040-IFO; Kp5783-IFO) were classified as weakly biofilm-producing with CV and one moderate strain (Kp0068-IFO) with the characterization process was strongly biofilm-producing according to CV staining. A possible explanation for this reduced agreement for K. pneumoniae could be associated with the production of thick mucus in these strains, which could have partially influenced the agreement between the tests. Indeed, it has been demonstrated that the CV tests may sometimes give false results due to non-specific staining properties (Pan, 2010; Merritt, 2005; Skogman, 2012). The strains of R. mannitolilytica showed 90% agreement between the tests, corresponding to a classification error. Complete agreement between the tests was observed for both S. aureus and S. epidermidis, while for other Gram-positive bacteria the agreement was 87%, corresponding to a classification error (Ef5515-IFO). Overall, complete agreement for Gram-negative strains was 68% while for Gram-positive bacteria it was more than 77%, showing an overall agreement of 72.5% of the samples analyzed. Thus, partial agreement was more than 95%, corresponding to 3 classification errors in 63 strains analyzed. It is important to note that in all the results in disagreement, the classification differs by a single category.

(52) Thus, total agreement was analyzed for the classification process and CV staining using McNemar's test. The results indicate that, for the identifications in disagreement, a statistically significant difference exists between the tests (p=0.007). More precisely, the classification process underestimates the biofilm production compared to CV. This result is entirely unsurprising, since it results from the non-specific staining property of CV. Indeed, crystal violet is known to bind to negatively-charged surface molecules, which are present both on bacteria and the extracellular matrix of the biofilm (Extremina, 2011). This may lead to overestimating the real adhesion capacity of different strains (Pan, 2010; Merritt, 2005; Skogman, 2012).

(53) TABLE-US-00005 TABLE 5 overall results for adhesion of the different bacterial species, obtained by the characterization process and CV staining Agreement (%) Characterization Total Bacterial process CV staining and species Na W M H Na W M H Total partial P. aeruginosa 0 2 3 7 0 2 3 7 83 100 K. pneumoniae 5 0 5 0 1 4 3 2 40 80 R. mannitolilytica 1 9 0 0 0 10 0 0 80 100 S. aureus 0 2 6 4 0 1 7 4 92 100 S. epidermidis 0 7 3 0 0 7 2 1 70 100 Other Gram+ 1 3 4 1 0 6 1 2 60 90 Na: non-biofilm-producing, W: weakly biofilm-producing, M: moderately biofilm-producing, H: strongly biofilm-producing

(54) Agreement between the characterization process according to the disclosure and CV was measured statistically by the kappa coefficient. The results showed good agreement between the tests for the weak (kappa=0.71±0.09; specificity=94.3%; precision=85.7%), moderate (kappa=0.63±0.11; specificity=84.8%; precision=84.1%) and strong (kappa=0.73±0.10; specificity=97.9%; precision=90.5%) biofilm producers, while for non-adhering/non-producing cells, the agreement was moderate (kappa=0.42±0.2; specificity=91.8%; precision=92.1%). Overall, the strength of agreement between these methods was good (kappa=0.66±0.07), in particular the characterization process demonstrating high levels of specificity and precision (specificity=92.2%; precision=88.1%).

(55) This example therefore clearly demonstrates that the characterization process, also referred to as cBFRT, is a rapid and reliable process for a quantitative evaluation of bacterial biofilm production. This process was tested with different bacterial species, with different biofilm formation phenotypes, including laboratory strains and clinical isolates. The ability of the process to evaluate biofilm production was compared with the CV test, which is a widely used method for biofilm quantification (Christensen, 1985; Stepanovic, 2000, Peeters, 2008). According to the kappa coefficient, the overall agreement between the tests was good (kappa=0.66±0.07) with the process showing high specificity (92.3%) and precision (88.1%) for classifying biofilm-producing bacteria. For all the reference strain results, the process was especially in agreement/consistent with the data reported in the literature. Complete category agreement between the process and CV staining was 72.5% while partial category agreement exceeded 96%, with only two incorrectly classified samples in the 63 strains analyzed. Complete category agreement was 68% for the Gram-negative bacteria and 77% for the Gram-positive bacteria, while partial category agreement was respectively 93.3% and 96.6%. The greatest number of results in disagreement is located in the weakly biofilm-producing/non-adhering group, and, in particular, all these strains exhibited a mucoid phenotype which affected the sensitivity of the CV test. It is worth noting that CV is a non-specific colorimetric assay, which stains living and dead bacteria and also the biofilm matrix. Thus, this test, which merely provides indirect quantification of the biofilm, may lead to overestimation of the results (Pan, 2010; Merritt, 2005; Skogman, 2012), in particular during assay on mucoid strains. Indeed, it has been demonstrated that the overproduction of mucus does not play a significant role in bacterial adhesion and biofilm matrix formation; however, it has a protective role against the host's immune response (Stapper, 2004; Leid, 2005). Conversely, the classification process provides a direct estimation of the true capacity of different bacterial strains to aggregate or adhere, thereby ensuring greater specificity. In fact, the different specificity of the two methods was indirectly confirmed by McNemar's test, which revealed that, in the case in which the methods are in disagreement, the classification process has a significant tendency (p=0.007) to underestimate the results. The other important differences between the classification process and CV are associated with the preparation and the implementation time and also the reproducibility of the data. With the classification process, the preparation of a full plate and inoculation requires for example 30 minutes plus for example five hours of incubation and for example a few minutes for analysis of the plate. Conversely, the CV assay enabled biofilm detection only after 24/48 hours of incubation and requires repeated washing steps and laborious staining procedures (Stepanovic, 2000; Djordjevic, 2002). The requirement for several non-standardized handling operations in the CV test has been associated with large intra- and inter-experimental variations leading to large standard deviations (Peeters, 2008). The lack of reproducibility is also one of the major drawbacks of other conventional techniques, in particular in high-throughput screening (Peeters, 2008).

(56) Nonetheless, the development of standardized methods to identify the phenotype of bacteria is of critical importance in clinical practice, since it may add a key decision-making element to support effective therapeutic management of “difficult infections” such as those associated with surgical devices (namely catheter-associated infections), which often lead to treatment failure and the removal of the apparatus, despite putting in place apparently suitable therapeutic strategies (Costerton, 2003). In fact, once the biofilm is established on the device, the individual cells have an increased tolerance to antimicrobial agents and antibiotic treatment alone is often insufficient. In vitro and in vivo experiments have demonstrated that in a biological biofilm matrix, the cells show a much higher minimum inhibitory concentration (MIC) (approximately 10-1000 times higher) than the same bacterial cells examined under planktonic growth conditions (Høiby, 2011; Hengzhuang, 2012). The effective in vivo antibiotic MIC for eradicating the biofilm may therefore be impossible to achieve by administering antibiotics at doses that appear to be effective for planktonic growth, due to the toxicity and side effects of the medications, including the limitations imposed by renal and/or hepatic function. Thus, the timely recognition of a strong biofilm producer, before the growth of a mature biofilm matrix, may contribute to the appropriate targeting of the therapeutic intervention (type, doses, duration) and decision-making (for example removing the catheter).

(57) As demonstrated, the classification process has been developed to work in combination with traditional microbiological clinical procedures, without altering the daily work routine. Immediately after isolation of the microorganism, for example on an agar plate, it is possible to analyze the biofilm-producing capacity.

(58) As demonstrated in this example, the process of the disclosure makes it possible to characterize biofilm-producing microorganisms, for example bacteria, yeasts etc. It is therefore obvious that the process may be used for characterizing the formation of biofilms by other organisms such as yeasts, with a similar benefit and applications to those claimed for bacteria.

(59) In addition, the classification process may provide information on the capacity of an unknown bacterial isolate to form a biofilm even before the microorganism is identified with conventional phenotyping techniques. Thus, the process according to the disclosure has numerous applications, for example in the field of dentistry, where biofilms are associated with major dental diseases such as tooth decay and periodontal diseases. In addition to these clinical applications, the classification process may represent an invaluable tool for example for the food processing industry and also the sanitation industry, including water systems.