microorganisms Article A High-Throughput Microtiter Plate Screening Assay to Quantify and Differentiate Species in Dual-Species Biofilms Víctor Campo-Pérez 1,2 , Júlia Alcàcer-Almansa 1,3 , Esther Julián 2, * and Eduard Torrents 1,3, * 1 Bacterial Infections and Antimicrobial Therapies Group, Institute for Bioengineering of Catalonia (IBEC), The Barcelona Institute of Science and Technology (BIST), Baldiri Reixac 15-21, 08028 Barcelona, Spain; vcampo@ibecbarcelona.eu (V.C.-P.); jalcacer@ibecbarcelona.eu (J.A.-A.) 2 Departament de Genètica i de Microbiologia, Facultat de Biociències, Universitat Autònoma de Barcelona, 08193 Barcelona, Spain 3 Microbiology Section, Department of Genetics, Microbiology and Statistics, Faculty of Biology, University of Barcelona, 643 Diagonal Ave., 08028 Barcelona, Spain * Correspondence: esther.julian@uab.cat (E.J.); etorrents@ibecbarcelona.eu or eduard.torrents@ub.edu (E.T.) Abstract: Pathogenic bacteria form biofilms during infection, and polymicrobial biofilms are the most frequent manifestation. Biofilm attachment, maturation, and/or antibiotic sensitivity are mainly evaluated with microtiter plate assays, in which bacteria are stained to enable the quantification of the biomass by optical absorbance or fluorescence emission. However, using these methods to distinguish different species in dual-species or polymicrobial biofilms is currently impossible. Colony- forming unit counts from homogenized dual-species biofilms on selective agar medium allow species differentiation but are time-consuming for a high-throughput screening. Thus, reliable, feasible, and fast methods are urgently needed to study the behavior of polymicrobial and dual-species communities. This study shows that Pseudomonas aeruginosa and Burkholderia cenocepacia strains expressing specific fluorescent or bioluminescent proteins permit the more efficient study of dual- species biofilms compared to other methods that rely on measuring the total biomass. Combining fluorescence and bioluminescence measurements allows an independent analysis of the different microbial species within the biofilm, indicating the degree of presence of each one over time during a dual-species biofilm growth. The quantitative strategies developed in this work are reproducible and Citation: Campo-Pérez, V.; recommended for dual-species biofilm studies with high-throughput microtiter plate approaches Alcàcer-Almansa, J.; Julián, E.; Torrents, E. A High-Throughput using strains that can constitutively express fluorescent or bioluminescent proteins. Microtiter Plate Screening Assay to Quantify and Differentiate Species in Keywords: crystal violet; biomass quantification; Pseudomonas aeruginosa; Burkholderia cenocepacia; Dual-Species Biofilms. Microorganisms dual-species biofilms 2023, 11, 2244. https://doi.org/ 10.3390/microorganisms11092244 Academic Editor: Alexis Bazire 1. Introduction Received: 7 August 2023 Polymicrobial infections denote diseases involving multiple species of microorgan- Revised: 31 August 2023 isms, a situation commonly associated with chronic infections [1]. The interactions between Accepted: 4 September 2023 pathogens can vary from competition for the same space or nutrients to cooperative in- Published: 6 September 2023 terspecies mechanisms that enhance joint survival and the evasion of host responses and increase antibiotic susceptibility [2]. These infections have become more frequent and relevant in recent years and are especially worrisome because biofilm-forming microbes are Copyright: © 2023 by the authors. associated with higher mortality rates [3]. Biofilms are composed of bacteria embedded in Licensee MDPI, Basel, Switzerland. an extracellular polymeric matrix consisting of polysaccharides, protein, and extracellular This article is an open access article DNA [4]. This growth behavior confers a broad range of advantages on pathogens, such distributed under the terms and as quorum sensing interactions and interspecies DNA exchanges that lead to metabolic conditions of the Creative Commons cooperation and, in some cases, a high tolerance to antibiotics. This provides a competitive Attribution (CC BY) license (https:// advantage and significantly enhances the resistance capacity inside the host [5]. Pathogenic creativecommons.org/licenses/by/ biofilm-forming bacteria present an increasing multidrug resistance to the currently avail- 4.0/). able antibiotics, so biofilm development is a relevant treatment target in the fight against Microorganisms 2023, 11, 2244. https://doi.org/10.3390/microorganisms11092244 https://www.mdpi.com/journal/microorganisms Microorganisms 2023, 11, 2244 2 of 12 chronic infections, and new anti-biofilm strategies are needed [6,7]. In this sense, over the last few years, strategies based on the delivery of anti-biofilm compounds, nanoparticles, and enzymes have been developed. Some studies have focused on inhibiting bacterial adhesion using compounds and enzymes that affect structures and biomolecules involved in cell adhesion, a key feature in the development of biofilms [8]. Others develop micropar- ticles based on polymeric carriers, dispersibility enhancer molecules, and anti-biofilm compounds [9]. The use of metal oxide nanoparticles has reported significant anti-biofilm efficiencies, but cytotoxicity issues in eukaryotic cells makes this nanoparticle applicability controversial [7]. The different in vitro methods developed for biofilm growth studies are categorized into static, continuous flow or open systems, and microfluidic biofilm culture [10]. The microtiter plate assay is the most widespread system used to study the growth of static biofilms and can be used for high-throughput screenings. This consists of growing biofilm- forming microorganisms in polystyrene microtiter plates using a suitable culture medium. Cells become attached to the walls of the wells, usually at the culture medium–air inter- face, and start to develop a biofilm growth form. After washing the wells to remove the planktonic cells, the adhered cells are allowed [11] to develop a mature biofilm. Biofilm development is then analyzed by the indirect quantification of the biomass, with crystal violet cell staining as the most commonly used method [12]. This method presents several advantages, such as low economic cost, technical simplicity, easy setup, and applicability to the high-throughput screening of biofilm-forming strains [13], and represents an ideal proposal for an anti-biofilm drug testing platform [14]. However, crystal violet staining also causes concern due to its toxicity to the environment and carcinogenic effects [15,16]. In recent years, alternatives to crystal violet, such as classic safranin, another cationic dye that binds to negatively charged bacterial cell walls, and the genetic material fluorescent dye SYTO 9 or acridine orange [17], have been used. These two new dyes stain live and dead cells and extracellular matrix components, thus generally overestimating biomass values [18]. Nevertheless, both are used for optical absorbance or fluorescence measure- ments of monomicrobial [19] and dual-species biofilms [20]. An outstanding alternative solution to this problem is resazurin-based viability staining approaches [15,17,21], which allow the quantification of biofilm-forming live cells exclusively by the metabolic reduction of nonfluorescent resazurin to fluorescent resorufin. In addition, this method enables a correlation of the fluorescence emitted with the number of cells in the biofilm, although it must first be optimized and calculated for each species [21]. However, general biomass staining methods and resazurin assays, as described previ- ously, are very limiting in studies of dual-species biofilms, since they stain nonspecifically, impeding the independent quantification of each individual microbial species present in the mixed biofilm. This is a recurring problem in these types of studies, which end up without the evaluation of which bacterial species percentage represents the mature dual-species biofilm. To reduce the impact of this limitation, homogenizing the dual-species biofilms and plating the bacterial suspensions in selective agar media allow the determination of the number of culturable cells of each microbial species, although this strategy is time- consuming and sometimes does not allow the online monitoring of antibiotic susceptibility testing of a high number of samples or not in a high-throughput screening. To the best of our knowledge, a reproducible, fast, easy-to-perform, and reliable method for the measurement of dual-species bacterial biofilms is greatly desired. New strategies that allow the study of dual-species biofilms in all their complexity need to be developed. The present study describes an applicable methodology based on the use of bacteria constitutively expressing fluorescent or bioluminescent proteins, and as a proof of concept, we specifically used the strains of Pseudomonas aeruginosa and Burkholderia cenocepacia species. These microbes have remarkable clinical relevance producing severe nosocomial infections associated with the formation of polymicrobial biofilms, especially in cystic fibrosis and other immunocompromised patients [22]. The formation of dual-species biofilms with these strains in microtiter plates using this developed methodology allows Microorganisms 2023, 11, 2244 3 of 12 their independent quantification, making it possible to determine the percentage of each bacterial species within the biofilm in a reproducible, fast, and reliable way and without the necessity of using advanced microscopy techniques. 2. Materials and Methods 2.1. Bacterial Strains and Culture Conditions Reference laboratory strains of P. aeruginosa PAO1 (ATCC 15692) and B. cenocepacia J2315 (ATCC BAA-245) were grown in Luria–Bertani (LB; tryptone 10 g/L, yeast extract 5 g/L, and sodium chloride 10 g/L) medium (Scharlab, Barcelona, Spain) and in tryptic soy broth (TSB; casein peptone 17 g/L, soy peptone 3 g/L, sodium chloride 5 g/L, dipotassium phosphate 2.5 g/L, and dextrose 2.5 g/L) medium (Scharlab, Barcelona, Spain) at 37 ◦ C and 30 ◦ C, respectively. P. aeruginosa PAO1::eGFP (MK171) [23] constitutively expressing a green fluorescent protein marker from the chromosome (eGFPmut3) and P. aeruginosa PAO1 chromosomally modified by the integration of a mini-Tn7-lux constitutively expressing the luciferase genes [24] were also used, supplementing the media with 50 µg/mL gentamicin as the selective pressure. B. cenocepacia was transformed with pETS248-Tc-E2Crimson plasmid (see the plasmid construction and bacterial transformation and its low copy number vector) and growth supplementing media with 80 µg/mL of tetracycline. The strains were preserved in 20% glycerol stocks at −80 ◦ C and were freshly revived for each experiment. 2.2. Plasmid Construction and B. cenocepacia Transformation We constructed a plasmid expressing an E2-Crimson red fluorescent protein. Recom- binant DNA techniques were performed using standard procedures on Escherichia coli DH5α cells [25]. To construct this plasmid, the promoter probe vector pETS134-GFP con- taining the promoter region of the P. aeruginosa nrdA gene was used as the backbone [26]. The gentamycin resistance gene fragment was removed from pETS134-GFP by double BglII and NsiI digestion (Thermo Fisher, Waltham, MA, USA). In parallel, a tetracycline resistance gene was amplified from the pEX18Tc vector [27] by a polymerase chain reac- tion (PCR) using the primers TcR_Fw (50 -GCGATGCATGACGTCAGGTGGCACTTTTC-30 ; NsiI) and TcR_Rv (50 -GCGAGATCTATTCACAGTTCTCCGCAAG-30 ; BglII). The resulting DNA amplification was gel-purified using a GeneJET Gel Extraction Kit (Thermo Fisher, Waltham, MA, USA), digested with NsiI and BglII, and ligated into the pETS134-GFP band fragment without gentamycin, generating the pETS247-Tc-GFP plasmid. Next, the GFP present in this plasmid was replaced by the E2-Crimson red fluorescent protein by digesting pETS247-Tc-GFP with ApaI and NdeI. Then, the DNA encoding the E2-Crimson fluorescent protein was amplified from the pUCP20T-E2Crimson vector [28] by PCR using the primers E2C_Fw (50 -GGGCATATGGGCGAGCTCGATAG-30 ; NdeI) and E2C_Rv (50 - ATTGGGCCCTTACAATTCGTCGTGCTTGTAC-30 ; ApaI). The amplified DNA fragment corresponding to E2-Crimson was gel-purified using a GeneJET Gel Extraction Kit (Thermo Fisher, Waltham, MA, USA), digested with ApaI and NdeI (Thermo Fisher, Waltham, MA, USA), and ligated into the digested pETS247-Tc-GFP to form pETS248-Tc-E2Crimson. The amplified DNA fragment corresponding to E2-Crimson was gel-purified using a GeneJET Gel Extraction Kit (Thermo Fisher, Waltham, MA, USA), digested with ApaI and NdeI (Thermo Fisher, Waltham, MA, USA), and ligated into the digested pETS247-Tc-GFP to form pETS248-Tc-E2Crimson. For B. cenocepacia transformation, a previously easily described procedure was used [29]. The electroporation parameters were 2.5 kV and 25 µF using a GenePulser Xcell Electropo- ration System (Bio-Rad, Hercules, CA, USA). The DNA-to-cell ratio was 0.45 µg plasmid DNA in 150 µL of competent bacterial cells. Microorganisms 2023, 11, 2244 4 of 12 2.3. Plasmid Maintenance Depending on Antibiotic Selection Pressure in the B. cenocepacia Transformed Strain The B. cenocepacia transformed strain integrity was confirmed by images detecting E2-Crimson in ON culture cells under a Nikon inverted fluorescence microscope, ECLIPSE Ti-S/L100 (Nikon, Tokyo, Japan). To test the plasmid maintenance in the transformed B. cenocepacia strain J2315 carrying pETS248-Tc-E2Crimson, we developed static 48 h biofilms in microtiter plates with or without the selective antibiotic (80 mg/mL of tetra- cycline). After 48 h, the biofilm was resuspended with 100 µL of 1× phosphate-buffered saline (PBS) (Fisher Scientific, Hampton, VA, USA), and the absorbance (Abs550nm ) and relative fluorescence units (RFU) of E2-Crimson per well (excitation at 580 ± 20 nm and emission at 635 ± 35 nm) were measured in a Spark multimode microplate reader (Tecan, Männedorf, Switzerland). 2.4. Development of a Microtiter Plate Screen for Dual-Species Biofilm Biomass Quantification In this work, as a proof of concept, we established a dual-species biofilm of a luminescence-expressing strain of P. aeruginosa and a fluorescence-expressing strain of B. cenocepacia. Overnight cultures of P. aeruginosa PAO1 chromosomally carrying the mini- Tn7-lux [24] and B. cenocepacia J2315 transformed with pETS247-Tc-E2Crimson were grown with the corresponding antibiotics in LB and TSB medium (Scharlab, Barcelona, Spain), respectively. Note that, in the case of undertaking studies using other bacterial species, different vectors with replication compatibility should be used. Then, both ON cultures were adjusted to OD550nm = 0.1, corresponding to 107 colony-forming units (CFU/mL), in TSB + 0.2% glucose without antibiotic pressure (Panreac Quimica, Barcelona, Spain) using a UV Mini 1240 spectrophotometer (Shimadzu, Kyoto, Japan) (see Figure 1A). The use of glucose is necessary to promote the development of biofilm in these species [30,31]. Then, 200 µL/well was added to Costar® 96-well flat black polystyrene plates (Corning, NY, USA), and the plates were incubated statically at 37 ◦ C in a compact mini-incubator (Labnet International, Edison, NJ, USA) with humidity saturation conditions to allow cell adhesion to the plastic walls and biofilm formation in the air–liquid interface. After 48 h, planktonic bacteria were removed by washing three times with 300 µL/well of 1× PBS (pH 7.4) to retain exclusively the attached biofilm-forming bacteria. At this point, for the development of the dual-species biofilm, strains prepared from freshly ON culture were inoculated together into wells containing a previously formed monomicrobial biofilm following the same steps previously explained (a schematic representation is shown in Figure 1). After an additional 24 h of incubation, the wells were washed 3× with PBS to remove planktonic bacteria, and the cell biomass of the biofilms was quantified using crystal violet staining. Fluorescence, bioluminescence, or CFU counting was carried out. Wells filled with the medium alone were used as a negative control, and the average values of fluorescence and bioluminescence reported in these wells were subtracted from all the data. For the crystal violet staining procedure (see Figure 1B), 200 µL/well methanol (Fisher Scientific, Hampton, VA, USA) was added for 15 min to fix the biofilms. Then, the methanol was removed, and the wells were allowed to evaporate until they were completely dry. Then, 200 µL/well of crystal violet (1%) (Sigma-Aldrich, St. Louis, MO, USA) was added for 5 min, and the excess dye was removed through water washes. Finally, the crystal violet was dissolved with 200 µL/well of acetic acid (33%) (Fisher Scientific, Hampton, VA, USA) under agitation (approximately 150 rpm), and the absorbance was measured at 570 nm for the semiquantitative analysis of the biofilm biomass. The fluorescence and biolumines- cence of the formed biofilms were quantified simply by homogenizing the dual-species biofilms in PBS through strong pipetting and pipette tip scraping on the walls of the wells (see Figure 1B). Microorganisms 2023, 11, x FOR PEER REVIEW 5 of 12 Microorganisms 2023, 11, 2244 5 of 12 Figure 1. Figure Schematicsummary 1. Schematic summaryofofthe themicrotiter microtiter plate plate assay assay forfor thethe formation formation andand quantification quantification of of static static biofilms. biofilms. (A) (A) TheThe procedures procedures followed followed to form to form monomicrobial monomicrobial and and dual-species dual-species biofilms. biofilms. (B) (B) Quantification Quantification methods methods usedused foranalysis for the the analysis of biofilms: of biofilms: CFUCFU counting, counting, crystal crystal violetviolet staining, staining, and fluorescence (RFU,(RFU, and fluorescence relative fluorescence relative units)units) fluorescence or bioluminescence (RLU, (RLU, or bioluminescence relativerelative light units) lightmeas- units) urements developed measurements as a novel, developed reproducible, as a novel, and reliable reproducible, alternative and reliable method. alternative method. The the For values of crystal crystal violetviolet stainingabsorbance procedure(Abs (see ), lux 1B), Figure 570nm luminescence 200 µL/well (relative light methanol units, RLU), (Fisher and E2-Crimson Scientific, Hampton, VA, fluorescence USA) wasexpression added for 15 (excitation min to fixat the ± 20 nm Then, 580biofilms. and emis- the methanol ± 35removed, sion at 635was nm) wereand quantified the wellsindependently were allowed in each well using to evaporate they® were a Spark until multimode com- microplate pletely dry. reader (TECAN). Then, 200 µL/wellToofestablish the percentages crystal violet of growth of (1%) (Sigma-Aldrich, St.each Louis,species MO, inUSA)the dual-species biofilms, the level of fluorescence/bioluminescence of was added for 5 min, and the excess dye was removed through water washes. Finally, the monospecies biofilms grown violet crystal underwas the dissolved same conditions with 200 inµL/well the same of microtiter acetic acidplate (33%)and during (Fisher the same Scientific, time Hamp- was VA, ton, quantified, and the USA) under mean of(approximately agitation these values was 150established rpm), and as the100% biofilm growth. absorbance was meas- ured at 570 nm for the semiquantitative analysis of the biofilm biomass. The fluorescence 2.5. Colony-Forming Unit Counts from Biofilms and bioluminescence of the formed biofilms were quantified simply by homogenizing the To quantify dual-species the in biofilms biofilms by CFU PBS through counts, strong the bacterial pipetting biofilm and pipette formed on tip scraping on the walls the walls of each well was removed of the wells (see Figure 1B). by scraping with a pipette tip and resuspended by pipetting in 100 of 1×ofPBS. TheµLvalues Then, crystal eachabsorbance violet biofilm cell(Abssuspension was transferred to a 1.5 mL 570nm), lux luminescence (relative light Eppendorf units, RLU),tube, disaggregated and E2-Crimson by pipetting, fluorescence and vortexed expression for 30 (excitation s to±separate at 580 20 nm and theemis- cells in the biofilm. Bacterial suspensions were then serially diluted in 1 sion at 635 ± 35 nm) were quantified independently in each well using a Spark multimode × PBS and ® plated on selective tryptic soy agar (TSA; casein peptone 15 g/L, soy peptone 5 microplate reader (TECAN). To establish the percentages of growth of each species in theg/L, sodium chloride 5 g/L, and agar dual-species 15 g/L) biofilms, themedium level of (Scharlab, Barcelona, Spain). TSA fluorescence/bioluminescence plates and TSA of monospecies plates biofilms supplemented with 2 µg/mL ciprofloxacin (Sigma-Aldrich, St. Louis, grown under the same conditions in the same microtiter plate and during the same time MO, USA) were used P. aeruginosa B. cenocepacia, was quantified, and the mean of these values was established as 100% biofilm growth. of to quantify and respectively. After 24 h and 48 h of incubation P. aeruginosa and B. cenocepacia, respectively, the colonies were counted, and the CFU/mL were determined. Ciprofloxacin 2.5. Colony-Forming Unit Counts from used since it inhibits the growth of P. aeruginosa but wasBiofilms allows the growth of B. cenocepacia. On TSA plates, P. aeruginosa grows earlier due to its To replication higher quantify thespeed, biofilms by CFU counts, preventing the bacterial the growth biofilm formed of B. cenocepacia, while, ononthe TSAwalls of plates each well was removed by scraping with a pipette supplemented with ciprofloxacin, only B. cenocepacia grows. tip and resuspended by pipetting in 100 µL of 1× PBS. Then, each biofilm cell suspension was transferred to a 1.5 mL Eppen- dorf tube, disaggregated by pipetting, and vortexed for 30 s to separate the cells in the Microorganisms 2023, 11, 2244 6 of 12 2.6. Fluorescence Microscopy to Visualize Biofilm Formation Monomicrobial or dual-species biofilms were developed in microtiter plates, as de- scribed above. Briefly, 200 µL/well of P. aeruginosa or B. cenocepacia ON suspensions with an OD550 nm = 0.1 were inoculated on the microtiter plates and incubated in TSB + 0.2% glucose at 37 ◦ C, maintaining the humidity saturation conditions over 48 h. Af- ter this time, the wells underwent three rounds of washing with PBS to remove plank- tonic bacteria. Dual-species biofilms were performed by the inoculation and growth of P. aeruginosa or B. cenocepacia suspensions into wells that already contained a developed monomicrobial biofilm. For microscopic observation, biofilms developed at the air–liquid interface were carefully detached from the walls of the wells using a micropipette tip, trying to minimize the possible decomposition or alteration of the original biofilm structure, and placed on a microscope slide (Thermo Fisher, Waltham, MA, USA). The appearance of the biofilm was analyzed with a Nikon inverted fluorescence microscope, ECLIPSE Ti-S/L100 (Nikon, Tokyo, Japan), coupled with a DS-Qi2 Nikon camera (Nikon, Tokyo, Japan). Fluorescent representative images were obtained with a 100× objective and subsequently processed using Fiji ImageJ software [32]. 3. Results and Discussion To establish a high-throughput system to precisely quantify the relative abundance of each microbial species in a dual-species biofilm, bacteria were labeled with designed low copy number vectors to express proteins that emit fluorescence or bioluminescence or chromosomically inserted in a single copy. In this study, we worked with P. aeruginosa PAO1 with a mini-Tn7-lux chromosomically inserted, a previously described strain consti- tutively expressing bioluminescence [24], and a B. cenocepacia J2315 strain transformed with pETS248-Tc-E2Crimson, a plasmid that expresses a red fluorescent protein (see Section 2.2). However, any plasmid with these properties can be used to label any biofilm-forming bacteria of interest. These two species were selected due to their frequent simultaneous presence in polymi- crobial or dual-species biofilms related to severe respiratory airway nosocomial infections. A critical clinical trait of P. aeruginosa is its capacity to interact and coexist with other mi- croorganisms in multispecies communities [33,34]. For example, bronchiectasis produced by chronic obstructive lung disease (COPD), cystic fibrosis (CF), and lung injury, among others, is usually related to P. aeruginosa and B. cenocepacia coinfections. Furthermore, these species demonstrate a propensity to associate intimately with polymicrobial biofilm communities [35], so its study in dual-species biofilms has key biological relevance. The strains of P. aeruginosa and B. cenocepacia used in this study contain resistance genes against gentamicin and tetracycline as selection markers, respectively. However, the expression of luciferase (lux) and E2-Crimson was not affected independently of the presence/absence of an antibiotic. P. aeruginosa mini-Tn7-lux was chromosomically inserted in a single copy expressing itself constitutively, and in B. cenocepacia, the plasmid was maintained in the cytoplasm, and the expression of E2-Crimson was maintained without an antibiotic when the strain developed in a static biofilm (Figure 2). Our strains were free of different antibiotic pressures during the development of the experiment. Microorganisms 2023, 11, x FOR PEER REVIEW 7 of 12 was maintained in the cytoplasm, and the expression of E2-Crimson was maintained with- Microorganisms 2023, 11, 2244 7 of 12 out an antibiotic when the strain developed in a static biofilm (Figure 2). Our strains were free of different antibiotic pressures during the development of the experiment. Figure2.2. Evaluation Figure Evaluation of of E2-Crimson E2-Crimson fluorescence fluorescence expression expression in the B. in the B. cenocepacia cenocepacia transformed transformed strain, strain, dependingon depending onthe thepresence/absence presence/absence of of the the plasmid plasmid selection selection antibiotic antibiotic tetracycline. tetracycline. (A) (A) Fluorescence Fluorescence microscopy image microscopy image at at100 100× ON culture × of an ON culture from from aasuccessfully successfullyelectrotransformed electrotransformed B. B.cenocepacia cenocepacia J2315expressing J2315 expressingthetheE2-Crimson E2-Crimsonfluorescent fluorescentprotein. protein.(B) (B) Relative Relative fluorescent fluorescent units units (RFU)/OD550nm (RFU)/OD 550nm comparison of comparison of three three independent independent replicates replicates(n (n==12) 12)after after4848 h of biofilm-forming h of biofilm-forming B. cenocepacia E2- B. cenocepacia Crimson growth with and without the presence of 80 µg/mL of tetracycline. ns, nonsignificant dif- E2-Crimson growth with and without the presence of 80 µg/mL of tetracycline. ns, nonsignificant ferences (unpaired t-test). differences (unpaired t-test). Once it Once it was was demonstrated demonstratedthat thatthetheantibiotic antibiotic selection selectionpressure pressure waswas unnecessary, unnecessary, the biofilm the development biofilm development experiments experiments werewere carried out without carried antibiotics out without in the culture antibiotics me- in the cul- dium. We first validated our method by growing monomicrobial ture medium. We first validated our method by growing monomicrobial biofilms of biofilms of P. aeruginosa P.PAO1 mini-Tn7-lux aeruginosa and B. cenocepacia PAO1 mini-Tn7-lux pETS248-Tc-E2Crimson and B. cenocepacia and quantifying pETS248-Tc-E2Crimson their bio- and quantifying mass by directly measuring the crystal violet and their fluorescence their biomass by directly measuring the crystal violet and their fluorescence and biolu- and bioluminescence (Figure 1) (see minescence Section (Figure 2.4).Section 1) (see Once the biofilm 2.4). Oncewas the formed, biofilm was the reproducibility of both strat- formed, the reproducibility egies of bothwas further evaluated, strategies was further showing evaluated,that showing the coefficient of variation that the coefficient(CoV; the ratio (CoV; of variation of the standard deviation to the mean) of all biofilm biomass values was the ratio of the standard deviation to the mean) of all biofilm biomass values was similar similar between bio- films quantified by both methods (crystal violet and fluorescence between biofilms quantified by both methods (crystal violet and fluorescence and biolumi- and bioluminescence) (Figure 3A). nescence) In the3A). (Figure P. aeruginosa PAO1 microtiter In the P. aeruginosa plate biofilm, PAO1 microtiter platea biofilm, 15% CoV was found a 15% CoV was be- tweenbetween found crystal violet replicates, crystal while a slight violet replicates, whiledecrease (11%) was a slight decrease observed (11%) using biolumi- was observed using nescence measurements. bioluminescence measurements.The sameThe tendency same tendencywas observed was observedin B.in cenocepacia B. cenocepacia biofilms but biofilms with but the the with highest variability, highest with variability, 47%47% with CoV CoVin the crystal in the violet crystal versus violet versus31% 31%in the fluores- in the fluo- cence measurements rescence measurements(Figure (Figure3A). 3A).TheThevariability variabilityof ofthe thebiofilm biofilm biomass quantification quantification in in the B. cenocepacia biofilms between the different wells was higher than the B. cenocepacia biofilms between the different wells was higher than for the P. aeruginosa for the P. aeruginosa biofilms,which biofilms, whichwere were more more homogeneous homogeneous betweenbetween replicates, replicates, independently independently of the of the method method of of quantification quantification used. These used. These results results confirmed confirmed that using that using bacterial bacterial label label biofilm tags allows tags al- lows biofilm quantification quantification results comparableresultsto comparable those obtained to those obtained with the withcrystal traditional the traditional crys- violet, avoid- ing tal its limitations violet, avoiding anditsreducing the and limitations variability reducing between replicates.between the variability Once thereplicates. validity ofOnce the measurements the validity of the wasmeasurements demonstrated,was dual-species demonstrated,biofilms of P. aeruginosa dual-species biofilmsandofB.P.cenocepacia aeruginosa were and established B. cenocepacia to quantify each biofilm-forming were established to quantify species eachindependently biofilm-forming (Figure 3B). Note species inde- that using crystal pendently (Figureviolet will quantify 3B). Note that using thecrystal total biomass violet willwithout distinguishing quantify the total biomass the specific with- contribution of eachthe out distinguishing bacterial specificspecies. contribution of each bacterial species. Monomicrobial mature biofilms of P. aeruginosa PAO1 mini-Tn7-lux and B. cenocepa- cia pETS248-Tc-E2Crimson were first established (48 h) and further co-inoculated with a culture of the opposite species for an additional 24 h to characterize their effect on the original mature biofilm (Figure 1). The results showed that the addition of P. aeruginosa significantly reduced the initial B. cenocepacia biofilm by approximately 50% (Figure 3B). cepacia cells when P. aeruginosa was present [35]. However, no direct growth-inhibitory activity has been proven between the two species. It was also seen that, in 24 h, P. aeru- ginosa was accountable for 80% of the biofilm cell abundance in a mature biofilm (72 h), despite growing together with B. cenocepacia. This result demonstrates the expected dom- Microorganisms 2023, 11, 2244 inant profile of P. aeruginosa over B. cenocepacia, as reported in previous studies [33,36], 8 of 12 even early in cocultures, validating our method. Figure 3. Assays validating the novel biofilm biomass quantification method for measuring fluo- Figure 3. Assays rescence validating the novel and bioluminescence. biofilm biomasstest (A) Reproducibility quantification comparing method forviolet the crystal measuring assayfluores- with the cence and bioluminescence. (A) Reproducibility test comparing the crystal violet assay with the bi- bioluminescence and fluorescence assay methodology on monomicrobial biofilms; the results corre- oluminescence and fluorescence assay methodology on monomicrobial biofilms; the results corre- spond to three independent replicates (n = 72). (B) Bioluminescence quantification of P. aeruginosa lux spond to three independent replicates (n = 72). (B) Bioluminescence quantification of P. aeruginosa and lux B. cenocepacia and E2-Crimson B. cenocepacia E2-Crimson forming formingdual-species biofilms. dual-species The biofilms. 100% The 100%biofilm growth biofilm growthvalue is the value is average expression reported by a monomicrobial biofilm grown for 72 h under the the average expression reported by a monomicrobial biofilm grown for 72 h under the same condi- same conditions as the tions as dual-species biofilms. the dual-species TheThe biofilms. values correspond values to three correspond independent to three independent experiments experiments(n =(n72); the = 72); the data data areare presented presented as the as the meanmean± standard ± standard deviation. deviation. The The coefficient coefficient of of variation variation (CoV) (CoV) shows shows the the variability variability in relation in relation to the to meantheinmean in each condition. each condition. (C) Colony-forming (C) Colony-forming unit (CFU) unit (CFU) quantification of P. aeruginosa lux and B. cenocepacia E2-Crimson forming mono- and dual-species biofilms; the results correspond to four CFU recounts of each condition. *** p < 0.001; ns, nonsignificant differences (unpaired t-test). Monomicrobial mature biofilms of P. aeruginosa PAO1 mini-Tn7-lux and B. cenocepacia pETS248-Tc-E2Crimson were first established (48 h) and further co-inoculated with a culture of the opposite species for an additional 24 h to characterize their effect on the original mature biofilm (Figure 1). The results showed that the addition of P. aeruginosa significantly reduced the initial B. cenocepacia biofilm by approximately 50% (Figure 3B). Detailed observation by fluorescence microscopy showed that B. cenocepacia remained in the base of the biofilm well, and the growth of P. aeruginosa began to infringe on it (Figure 4). Microorganisms 2023, 11, 2244 9 of 12 Microscopic images confirmed the coexistence of both bacterial species in biofilm growth. Previous dual-species biofilm studies have already reported a decrease in B. cenocepacia Microorganisms 2023, 11, x FOR PEER REVIEW 9 of 12 cells when P. aeruginosa was present [35]. However, no direct growth-inhibitory activity has been proven between the two species. It was also seen that, in 24 h, P. aeruginosa was accountable for 80% of the biofilm cell abundance in a mature biofilm (72 h), despite quantification of P. aeruginosa growing together lux and B. cenocepacia with B. cenocepacia. E2-Crimson This result formingthe demonstrates mono- and dual-species expected dominant biofilms; the results correspond to four CFU recounts of each condition. *** p < 0.001; ns, nonsignif- profile of P. aeruginosa over B. cenocepacia, as reported in previous studies [33,36], even early icant differences (unpaired t-test). in cocultures, validating our method. Figure 4. Representative fluorescence microscopy images of P. aeruginosa and B. cenocepacia mo- Figure 4. Representative fluorescence microscopy images of P. aeruginosa and B. cenocepacia mo- nomicrobial and dual-species biofilms. P. aeruginosa eGFP is reported in green, and B. cenocepacia nomicrobial and dual-species biofilms. P. aeruginosa eGFP is reported in green, and B. cenocepacia E2-Crimson is reported in red. Monomicrobial biofilm wells were grown for 48 h and then sub- E2-Crimson is reported in red. Monomicrobial biofilm wells were grown for 48 h and then subse- sequently quently inoculated inoculated withwith the opposite the opposite bacteria bacteria to form to form dual-species dual-species biofilms biofilms (growth (growth for an for an addi- additional tional 24 h).24 h). On the other hand, the growth of the P. aeruginosa biofilm remained unchanged when On the other hand, the growth of the P. aeruginosa biofilm remained unchanged when B. cenocepacia was later inoculated over a mature biofilm (48 h), reaching similar values to B. cenocepacia was later inoculated over a mature biofilm (48 h), reaching similar values to those of the monomicrobial P. aeruginosa biofilm (100% growth). However, B. cenocepacia those of the monomicrobial P. aeruginosa biofilm (100% growth). However, B. cenocepacia grew and settled into the formed P. aeruginosa biofilm, although its growth was only 35% of grew and settled into the formed P. aeruginosa biofilm, although its growth was only 35% the total growth of a monomicrobial B. cenocepacia mature biofilm (Figure 3B). These results of the total growth of a monomicrobial B. cenocepacia mature biofilm (Figure 3B). These obtained through quantification by fluorescence and bioluminescence measurements were results obtained through quantification by fluorescence and bioluminescence measure- verified by observing the appearance of the biofilms by fluorescence microscopy. While ments were verified by observing the appearance of the biofilms by fluorescence micros- B. cenocepacia forms a dense biofilm when it grows alone, its development is seriously copy. While B. in compromised cenocepacia forms the coculture a dense with biofilm since P. aeruginosa, whenthe it grows alone, latter is its development the predominant is species seriously compromised (Figure 4). Importantly,inwetheobserved coculturethewith P. aeruginosa, establishment ofsince the latter isbiofilm a B. cenocepacia the predom- over a inant species (Figure 4). Importantly, we observed the establishment of a B. cenocepacia biofilm over a mature P. aeruginosa biofilm, which always showed some dominant growth over other types of bacterial species [33]. The results obtained by the fluorescence and bioluminescence measurements were further validated by analyzing the CFU counts for every tested condition. As Figure 3C Microorganisms 2023, 11, 2244 10 of 12 mature P. aeruginosa biofilm, which always showed some dominant growth over other types of bacterial species [33]. The results obtained by the fluorescence and bioluminescence measurements were further validated by analyzing the CFU counts for every tested condition. As Figure 3C shows, the CFUs of P. aeruginosa were stable regardless of the inoculation time point and the presence of B. cenocepacia in the culture. In contrast, the CFUs/mL of B. cenocepacia were the highest in the absence of P. aeruginosa, whereas there was a significant decrease in CFUs after the addition of P. aeruginosa to a mature B. cenocepacia biofilm. Specifically, the number of B. cenocepacia colonies obtained in the dual-species biofilms was three orders of magnitude lower than that in the control monomicrobial biofilms. When incubated for 24 h over a P. aeruginosa mature biofilm, the viability of B. cenocepacia decreased compared to its monoculture, although it was still able to form a biofilm. These results are in line with the fluorescence and bioluminescence measurements described in Figure 3A,B and therefore validate our biofilm quantification method. Using genetically transformed bacterial strains to express fluorescence and biolumi- nescence in microtiter plates has excellent advantages in studying dual-species biofilms. In this study, a broad host range of replicative plasmids with the origins of the replications compatible for the tested bacteria was used. However, using recombinant strains containing antibiotic resistance genes could limit the assays focused on antibiotics. Therefore, the choice of the selection marker in the vector to avoid interference in the study is crucial. Different constructions and plasmids will be needed if other microorganisms are used in further studies. The most important advantage of this method is the possibility of performing individu- alized quantifications of each bacterial species forming a dual-species biofilm, determining the percentage that each one occupies. It facilitates the independent monitoring of each species in dual-species biofilm evolution and the measurement of solely live microbial cells by preventing the overestimation of the biomass by measuring dead cells in the biofilm matrix. In this sense, the constitutive expression of the fluorescence/bioluminescence proteins shown in our method could be stopped in dormant cells in biofilms and may produce an underestimation of a biofilm biomass being a methodological noncontrollable limitation. Note that, in most of the current dual-species biofilm biomass quantification methods for static setups, it is impossible to easily evaluate the contribution of each bac- terial species to biofilm development [37,38]. Additionally, this problem is accentuated during the evaluation of different antimicrobial therapies and the study of the specific effect of each bacterial species on a dual-species biofilm [39]. Furthermore, the biofilm quantifications reported here by fluorescent and biolumines- cent strains are reproducible, with the coefficients of variation between replicates lower than those reported using the traditional crystal violet method, reflecting improved reproducibil- ity. Also, the fluorescent and bioluminescent procedure is faster and more direct than that of crystal violet, which involves several steps with waiting periods. In addition, fluorescence is a methodology broadly used to evaluate and quantify biofilm formation [2,23]. 4. Conclusions The study and characterization of dual or polymicrobial static biofilms in vitro remain challenging, and the most widely employed method for this purpose is the development of biofilms in microtiter plates. However, the lack of specificity of the current methods hinders the individual identification and quantification of each species within a biofilm. In this context, this study demonstrated that utilizing genetically modified strains emitting fluorescence or bioluminescence enables the rapid, reproducible, and reliable determination of individual species proportions within a biofilm. These biofilm quantification results, as well as the reliability of the genetically transformed strains, have been verified and validated using time-demanding techniques such as CFU counting and observation through fluorescence microscopy. Microorganisms 2023, 11, 2244 11 of 12 Despite its significant advantages, the application of the presented method requires attention to specific details. It is essential to genetically transform the desired strains and to carefully select the appropriate selectable marker (often an antibiotic resistance gene) within the vectors to avoid interference with subsequent studies. In essence, we presented a straightforward, faster, and more reproducible strategy compared to the current methods for quantifying dual-species biofilms. Author Contributions: V.C.-P. and J.A.-A. performed the experiments. E.J. and E.T. directed the research and revised the experimental data. All authors have read and agreed to the published version of the manuscript. Funding: This study was supported by grants from the Ministerio de Ciencia, Innovación y Universi- dades and Agencia Estatal de Investigación (AEI), Spain, co-funded by Fondo Europeo de Desarrollo Regional, FEDER, European Union (RTI2018-098777-B-I00, RTI2018-098573-B-I00, PID2021-125801OB- I00, PID2021-122331OB-I00, PDC2022-133577-I00, and PLEC2022-009356); the CERCA programme and AGAUR-Generalitat de Catalunya (2021SGR-00092 and 2021SGR-00145); the European Regional Development Fund (FEDER); Catalan Cystic Fibrosis Association; and Obra Social “La Caixa”. V.C.-P. is a recipient of a Ph.D. contract (FI) from the Generalitat de Catalunya. J.A.-A. is thankful to MCIN for its financial support through a PRE2021-098703 grant funded by MCIN/AEI/ 10.13039/501100011033 and by ESF investing in your future. Data Availability Statement: Research data will be shared by the corresponding authors upon request. Conflicts of Interest: The authors declare no conflict of interest. References 1. Wolcott, R.; Costerton, J.W.; Raoult, D.; Cutler, S.J. The Polymicrobial Nature of Biofilm Infection. Clin. Microbiol. Infect. 2013, 19, 107–112. [CrossRef] 2. Gabrilska, R.A.; Rumbaugh, K.P. Biofilm Models of Polymicrobial Infection. Future Microbiol. 2015, 10, 1997–2015. [CrossRef] [PubMed] 3. Brogden, K.A.; Guthmiller, J.M.; Taylor, C.E. Human Polymicrobial Infections. Lancet 2005, 365, 253–255. [CrossRef] [PubMed] 4. Limoli, D.H.; Jones, C.J.; Wozniak, D.J. Bacterial Extracellular Polysaccharides in Biofilm Formation and Function. Microbiol. Spectr. 2015, 3, 10. [CrossRef] [PubMed] 5. Høiby, N.; Ciofu, O.; Johansen, H.K.; Song, Z.J.; Moser, C.; Jensen, P.Ø.; Molin, S.; Givskov, M.; Tolker-Nielsen, T.; Bjarnsholt, T. The Clinical Impact of Bacterial Biofilms. Int. J. Oral Sci. 2011, 3, 55–65. [CrossRef] 6. Pecoraro, C.; Carbone, D.; Deng, D.; Cascioferro, S.M.; Diana, P.; Giovannetti, E. Biofilm Formation as Valuable Target to Fight against Severe Chronic Infections. Curr. Med. Chem. 2022, 29, 4307–4310. [CrossRef] 7. Mishra, S.; Gupta, A.; Upadhye, V.; Singh, S.C.; Sinha, R.P.; Häder, D.P. Therapeutic Strategies against Biofilm Infections. Life 2023, 13, 172. [CrossRef] 8. Pecoraro, C.; Carbone, D.; Parrino, B.; Cascioferro, S.; Diana, P. Recent Developments in the Inhibition of Bacterial Adhesion as Promising Anti-Virulence Strategy. Int. J. Mol. Sci. 2023, 24, 4872. [CrossRef] 9. Xiroudaki, S.; Sabbatini, S.; Pecoraro, C.; Cascioferro, S.; Diana, P.; Wauthoz, N.; Antognelli, C.; Monari, C.; Giovagnoli, S.; Schoubben, A. Development of a New Indole Derivative Dry Powder for Inhalation for the Treatment of Biofilm-Associated Lung Infections. Int. J. Pharm. 2023, 631, 122492. [CrossRef] 10. McLean, R.J.C.; Bates, C.L.; Barnes, M.B.; McGowin, C.L.; Aron, G.M. Methods of Studying Biofilms. In Microbial Biofilms; Wiley: Hoboken, NJ, USA, 2014; pp. 379–413. [CrossRef] 11. Peeters, E.; Nelis, H.J.; Coenye, T. Comparison of Multiple Methods for Quantification of Microbial Biofilms Grown in Microtiter Plates. J. Microbiol. Methods 2008, 72, 157–165. [CrossRef] 12. Christensen, G.D.; Simpson, W.A.; Younger, J.J.; Baddour, L.M.; Barrett, F.F.; Melton, D.M.; Beachey, E.H.; Christensen, G.; Simpson, W.; Beachey, E.; et al. Adherence of Coagulase-Negative Staphylococci to Plastic Tissue Culture Plates: A Quantitative Model for the Adherence of Staphylococci to Medical Devices. J. Clin. Microbiol. 1985, 22, 996. [CrossRef] [PubMed] 13. Coffey, B.M.; Anderson, G.G. Biofilm Formation in the 96-Well Microtiter Plate. Methods Mol. Biol. 2014, 1149, 631–641. [CrossRef] [PubMed] 14. O’Toole, G.A. Microtiter Dish Biofilm Formation Assay. J. Vis. Exp. 2011, 47, e2437. [CrossRef] 15. Ommen, P.; Zobek, N.; Meyer, R.L. Quantification of Biofilm Biomass by Staining: Non-Toxic Safranin Can Replace the Popular Crystal Violet. J. Microbiol. Methods 2017, 141, 87–89. [CrossRef] [PubMed] 16. Mani, S.; Bharagava, R.N. Exposure to Crystal Violet, Its Toxic, Genotoxic and Carcinogenic Effects on Environment and Its Degradation and Detoxification for Environmental Safety. Rev. Environ. Contam. Toxicol. 2016, 237, 71–104. [CrossRef] Microorganisms 2023, 11, 2244 12 of 12 17. Amador, C.I.; Stannius, R.O.; Røder, H.L.; Burmølle, M. High-Throughput Screening Alternative to Crystal Violet Biofilm Assay Combining Fluorescence Quantification and Imaging. J. Microbiol. Methods 2021, 190, 106343. [CrossRef] 18. Merritt, J.H.; Kadouri, D.E.; O’Toole, G.A. Growing and Analyzing Static Biofilms. Curr. Protoc. Microbiol. 2011, 22, 1B. [CrossRef] 19. Xu, Z.; Liang, Y.; Lin, S.; Chen, D.; Li, B.; Li, L.; Deng, Y. Crystal Violet and XTT Assays on Staphylococcus aureus Biofilm Quantification. Curr. Microbiol. 2016, 73, 474–482. [CrossRef] 20. Peters, B.M.; Ward, R.M.; Rane, H.S.; Lee, S.A.; Noverr, M.C. Efficacy of Ethanol against Candida albicans and Staphylococcus aureus Polymicrobial Biofilms. Antimicrob. Agents Chemother. 2013, 57, 74–82. [CrossRef] 21. Van den Driessche, F.; Rigole, P.; Brackman, G.; Coenye, T. Optimization of Resazurin-Based Viability Staining for Quantification of Microbial Biofilms. J. Microbiol. Methods 2014, 98, 31–34. [CrossRef] 22. Eberl, L.; Tümmler, B. Pseudomonas aeruginosa and Burkholderia cepacia in Cystic Fibrosis: Genome Evolution, Interactions and Adaptation. Int. J. Med. Microbiol. 2004, 294, 123–131. [CrossRef] [PubMed] 23. Blanco-Cabra, N.; López-Martínez, M.J.; Arévalo-Jaimes, B.V.; Martin-Gómez, M.T.; Samitier, J.; Torrents, E. A New BiofilmChip Device for Testing Biofilm Formation and Antibiotic Susceptibility. NPJ Biofilms Microbiomes 2021, 7, 62. [CrossRef] [PubMed] 24. Moya-Andérico, L.; Admella, J.; Fernandes, R.; Torrents, E. Monitoring Gene Expression during a Galleria mellonella Bacterial Infection. Microorganisms 2020, 8, 1798. [CrossRef] [PubMed] 25. Maniatis, T.; Fritsch, E.F.; Sambrook, J. Cloning: A Laboratory Manual; Cold Spring Harbor Laboratory: Cold Spring Harbor, NY, USA, 1982; p. 545. 26. Sjöberg, B.M.; Torrents, E. Shift in Ribonucleotide Reductase Gene Expression in Pseudomonas aeruginosa during Infection. Infect. Immun. 2011, 79, 2663. [CrossRef] 27. Hoang, T.T.; Karkhoff-Schweizer, R.R.; Kutchma, A.J.; Schweizer, H.P. A Broad-Host-Range Flp-FRT Recombination System for Site-Specific Excision of Chromosomally-Located DNA Sequences: Application for Isolation of Unmarked Pseudomonas aeruginosa Mutants. Gene 1998, 212, 77–86. [CrossRef] 28. Barbier, M.; Damron, F.H. Rainbow Vectors for Broad-Range Bacterial Fluorescence Labeling. PLoS ONE 2016, 11, e0146827. [CrossRef] 29. Dubarry, N.; Du, W.; Lane, D.; Pasta, F. Improved Electrotransformation and Decreased Antibiotic Resistance of the Cystic Fibrosis Pathogen Burkholderia cenocepacia Strain J2315. Appl. Environ. Microbiol. 2010, 76, 1095–1102. [CrossRef] 30. Ramli, N.S.K.; Eng Guan, C.; Nathan, S.; Vadivelu, J. The Effect of Environmental Conditions on Biofilm Formation of Burkholderia pseudomallei Clinical Isolates. PLoS ONE 2012, 7, e44104. [CrossRef] 31. She, P.; Wang, Y.; Liu, Y.; Tan, F.; Chen, L.; Luo, Z.; Wu, Y. Effects of Exogenous Glucose on Pseudomonas aeruginosa Biofilm Formation and Antibiotic Resistance. Microbiologyopen 2019, 8, e933. [CrossRef] 32. Schindelin, J.; Arganda-Carreras, I.; Frise, E.; Kaynig, V.; Longair, M.; Pietzsch, T.; Preibisch, S.; Rueden, C.; Saalfeld, S.; Schmid, B.; et al. Fiji: An Open-Source Platform for Biological-Image Analysis. Nat. Methods 2012, 9, 676–682. [CrossRef] 33. Cendra, M.D.M.; Torrents, E. Pseudomonas aeruginosa Biofilms and Their Partners in Crime. Biotechnol. Adv. 2021, 49. [CrossRef] [PubMed] 34. Bragonzi, A.; Farulla, I.; Paroni, M.; Twomey, K.B.; Pirone, L.; Lorè, N.I.; Bianconi, I.; Dalmastri, C.; Ryan, R.P.; Bevivino, A. Modelling Co-Infection of the Cystic Fibrosis Lung by Pseudomonas aeruginosa and Burkholderia cenocepacia Reveals Influences on Biofilm Formation and Host Response. PLoS ONE 2012, 7, e52330. [CrossRef] 35. Berlutti, F.; Morea, C.; Battistoni, A.; Sarli, S.; Cipriani, P.; Superti, F.; Ammendolia, M.G.; Valenti, P. Iron Availability Influences Aggregation, Biofilm, Adhesion and Invasion of Pseudomonas aeruginosa and Burkholderia cenocepacia. Int. J. Immunopathol. Pharmacol. 2005, 18, 661–670. [CrossRef] [PubMed] 36. Schwab, U.; Abdullah, L.H.; Perlmutt, O.S.; Albert, D.; William Davis, C.; Arnold, R.R.; Yankaskas, J.R.; Gilligan, P.; Neubauer, H.; Randell, S.H.; et al. Localization of Burkholderia cepacia Complex Bacteria in Cystic Fibrosis Lungs and Interactions with Pseudomonas aeruginosa in Hypoxic Mucus. Infect. Immun. 2014, 82, 4729–4745. [CrossRef] [PubMed] 37. Ren, D.; Madsen, J.S.; de la Cruz-Perera, C.I.; Bergmark, L.; Sørensen, S.J.; Burmølle, M. High-Throughput Screening of Multispecies Biofilm Formation and Quantitative PCR-Based Assessment of Individual Species Proportions, Useful for Exploring Interspecific Bacterial Interactions. Microb. Ecol. 2013, 68, 146–154. [CrossRef] [PubMed] 38. Magana, M.; Sereti, C.; Ioannidis, A.; Mitchell, C.A.; Ball, A.R.; Magiorkinis, E.; Chatzipanagiotou, S.; Hamblin, M.R.; Had- jifrangiskou, M.; Tegos, G.P. Options and Limitations in Clinical Investigation of Bacterial Biofilms. Clin. Microbiol. Rev. 2018, 31, e00084-16. [CrossRef] 39. Koo, H.; Allan, R.N.; Howlin, R.P.; Stoodley, P.; Hall-Stoodley, L. Targeting Microbial Biofilms: Current and Prospective Therapeutic Strategies. Nat. Rev. Microbiol. 2017, 15, 740–755. [CrossRef] Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.