**Bacteriophage Therapy for Clinical Biofilm Infections: Parameters That Influence Treatment Protocols and Current Treatment Approaches**

### **James B. Doub**

Division of Infectious Diseases, University of Maryland School of Medicine, Baltimore, MD 21201, USA; jdoub@ihv.umaryland.edu

Received: 30 September 2020; Accepted: 10 November 2020; Published: 12 November 2020 -

**Abstract:** Biofilm infections are extremely difficult to treat, which is secondary to the inability of conventional antibiotics to eradicate biofilms. Consequently, current definitive treatment of biofilm infections requires complete removal of the infected hardware. This causes significant morbidity and mortality to patients and therefore novel therapeutics are needed to cure these infections without removal of the infected hardware. Bacteriophages have intrinsic properties that could be advantageous in the treatment of clinical biofilm infections, but limited knowledge is known about the proper use of bacteriophage therapy in vivo. Currently titers and duration of bacteriophage therapy are the main parameters that are evaluated when devising bacteriophage protocols. Herein, several other important parameters are discussed which if standardized could allow for more effective and reproducible treatment protocols to be formulated. In addition, these parameters are correlated with the current clinical approaches being evaluated in the treatment of clinical biofilm infections.

**Keywords:** biofilm; bacteriophage therapy; prosthesis related infections; hardware infections; left ventricular assist devices

### **1. Introduction**

When bacteria attach to surfaces they can form an extracellular matrix comprised of proteins, polysaccharides, extracellular DNA and water [1–5]. The extracellular matrix and the bacteria that reside within this matrix are what comprise biofilms. Contrary to planktonic bacteria that are free floating, biofilm bacteria are sessile. Bacteria in these sessile states have drastically different characteristics than planktonic bacteria causing conventional antibiotics to have limited ability to eradicate biofilms [1–5]. This stems from the reduced metabolic activity of biofilm bacteria and the architecture of biofilm itself [1]. The minimal inhibitory concentration of antibiotics to biofilm bacteria can be up to 1000 times that of planktonic bacteria [1]. Therefore to definitively cure these infections surgical removal of all the hardware (Figure 1) that harbor biofilms, in combination with prolonged systemic antibiotic therapy, is required. However, this causes significant morbidity and mortality to the patients who suffer from these infections. As a result, new antimicrobial methods are needed that can treat these biofilm infections without removal of the hardware. Bacteriophages might be such an adjuvant therapeutic.

ǻǼȱ

ǻǼȱ

ǻǼȱ

**Figure 1.** Examples of a few types of "hardware" that once infected require removal for definitive cure of these biofilm infections. (**A**) A lumbar posterior spinal rods and pedicle screw construct. (**B**) Total knee arthroplasty with long stem femoral and tibial components. (**C**) Total hip arthroplasty.

### *1.1. Bacteriophages*

Bacteriophages are viruses with a very narrow spectrum of activity to only certain strains of a certain bacterial species. Infection of human cells has not been observed and therefore bacteriophages are attractive therapeutics to use in bacterial infections [6,7]. These viruses can either be lytic or lysogenic. Lytic bacteriophages hold the most promise in treating infections given their ability to lyse bacteria. Lysogenic bacteriophages incorporate into bacterial DNA and do not induce bacterial lysis until reactivated at a later time, making them not advantageous in the treatment of infections. In nature, the majority of bacteria live in sessile states associated with biofilms and through evolution bacteriophages have coevolved to be able to infect and lyse bacteria inside biofilms [6,7].

### *1.2. Bacteriophage Activity in Biofilms*

In order to eradicate a clinical biofilm, an effective agent must be able to penetrate the biofilm and kill the bacteria that are present in various metabolic states while also degrading the biofilm extracellular matrix. Bacteriophages possess these abilities, but are not motile agents [6,7]. Therefore if bacteriophages can establish an infection within a biofilm, high rates of replication can occur given the high densities of biofilm bacteria in a structured space [8]. Bacteriophages even retain lytic activity against reduced metabolically active bacteria [9,10]. In the deepest regions of a biofilm, bacteria known as persister cells are semi-dormant [11]. All conventional therapeutics have limited activity to persister cells [11]. However, bacteriophages have the ability to infect persister cells and then lyse these bacteria once they become metabolically active again [12].

Bacteriophages also can enzymatically degrade the biofilm extracellular matrix thus allowing for dissemination within the biofilm. This occurs through use of endolysins and depolymerases [13,14]. Enodolysins are enzymes produced by bacteriophages to weaken the bacterial cell wall allowing for lysis to occur releasing their progeny [13]. Endolysins also have activity in degrading the extracellular matrix [13]. Depolymerases are enzymes attached to some bacteriophages that can also degrade the biofilm matrix in functionally different ways to endolysins [13]. Unique to bacteriophages is their ability to self-replicate and increase their own concentrations. This occurs when bacteriophage induced bacterial lysis causes release of progeny into the local environment to infect other bacterial cells. In the confined space of a biofilm this could be advantageous allowing for bacteriophages to infect biofilm bacteria and slowly degrade the biofilm [6]. However bacteriophages are not motile agents and finding biofilm bacteria may be an arduous undertaking if not directly applied to the biofilm.

Several preclinical animal studies support the use of bacteriophage therapy in clinical biofilm infections [15–23]. These studies show that local administrations of bacteriophages to the site of biofilm infections result in biofilm reduction [15–23]. In addition, these studies show that, without local administration of bacteriophage therapy, reduction in biofilms on hardware is not significantly reduced [19]. One of the most relevant preclinical studies was conducted by Morris et al. [19]. Rats were implanted with replica orthopedic prosthetics and then infected with *Staphylococcus aureus.* A total of 3 weeks later rats were given intraperitoneal bacteriophage therapy for 3 days. Results show synergistic activity of bacteriophage therapy with vancomycin in local infected tissues but no statistical reductions in biofilm burden on infected prosthetic material [19]. These findings support other preclinical testing that direct instilment of bacteriophage therapy to the site of biofilm infection is needed to achieve significant biofilm reduction. The intrinsic abilities of bacteriophages and results of animal studies support evaluation of bacteriophages in the treatment of clinical biofilm infections. However bacteriophages are not like conventional antibiotics and several parameters need to be understood before using this therapeutic in vivo.

### **2. Parameters that Impact Treatment Protocols**

Unlike conventional antibiotics, bacteriophage therapy is not a one size fits all antimicrobial therapeutic. Rather a bacteriophage that has robust activity to a clinical isolate of a specific bacterial species may have widely different activity or no activity to another clinical isolate of the same bacterial species. Many other aspects of bacteriophage therapy are poorly understood and not standardized, making creation of treatment protocols an arduous undertaking. At the present time, standardization of protocols can only be achieved with respect to bacteriophage titers and duration of therapy. However, this limits bacteriophage therapy to be used similarly to conventional antibiotics and does not incorporate many other parameters that need to be considered to devise advantageous, reproducible treatment protocols. Herein several additional important parameters are discussed.

### *2.1. Current "Susceptibility" Testing*

At the present time, bacteriophage therapy requires a clinical isolate to be tested against either a library of individual bacteriophages or to a set cocktail of bacteriophages to ensure susceptibility. Given the narrow spectrum of activity, even with the use of bacteriophage cocktails, susceptibility testing is warranted. There is no proverbial gold standard of susceptibility testing and no standardized "breakpoints" are available to determine if a bacteriophage has adequate activity to be used clinically. Therefore, it is vital to understand how in vitro susceptibility testing is conducted to be able to extrapolate these findings to in vivo use. Testing for phage susceptibility usually includes two methods.


Bacteriophages that form plaques and can inhibit bacterial growth are considered potential therapeutic options. Complicating this testing is that different MOIs can have drastically different growth inhibition durations. For instance an MOI of 100 might inhibit growth for 24 h while an MOI of 10 for the same bacteriophage might not inhibit growth at all. Figure 1 reinforces this for a *Staphylococcus epidermidis* clinical isolate in which PM448, PM472, PM421 have different growth inhibition durations for different MOIs of 100 and 10. This has ramifications when treating biofilm infections as reproducing the high MOI seen in vitro may not occur unless direct bacteriophage application is applied to biofilms. This can also have implications for resistance formation which will be discussed below.

Another important implication of this testing is the lack of standardization with respect to the duration of growth inhibition. A bacteriophage that inhibits growth for 48 h likely has different therapeutic potential compared to a different bacteriophage that only inhibits growth for 8 h. In correlation, different in vivo bacterial metabolic states may require different levels of growth inhibition. For instance, biofilm bacteria are less metabolically active then planktonic bacteria and therefore less in vitro growth inhibition might be needed compared to if bacteriophage therapy is being used to treat a planktonic infection. No standardized growth inhibition duration has been proposed, thereby exposing treatment protocols to potential reproducibility issues. To improve reproducibility, it may be important to standardize what is considered adequate growth inhibition depending on how a bacteriophage therapy is going to be used (intravenously vs. directly applied to biofilms). It should also be mentioned that "susceptibility" testing is usually only conducted against planktonic bacteria. Routine testing for a bacteriophage's ability to remove in vitro biofilms is usually not conducted. However, in the treatment of biofilm infections, determining the ability of a candidate bacteriophage (or cocktail) to reduce in vitro biofilms should be considered as an additional susceptibility testing step once adequate growth inhibition and formation of plaques has been proven.

**Figure 2.** Bacterial growth inhibition curves or "Phagogram" for a compassionate use *Staphylococcus epidermidis* case in a recalcitrant prosthetic joint infection. Different bacteriophages are indicated by PM241-PM472. MOI refers to multiplicity of infection. Growth control is the bacterial isolate with no bacteriophages. Each box has time on the *x*-axis from 0 to 48 h. This figure shows how growth inhibition testing is conducted to determine potential bacteriophage candidates (PM448, PM472, and PM421). This figure also shows how different MOIs can cause different growth inhibition durations as seen with bacteriophages: PM448, PM472, and PM421.

### *2.2. Pharmacology*

The main routes of phage administration that are being investigated in western medicine for the treatment of biofilm infections are local administration directly applied to the infected hardware and intravenous therapy. Eastern European studies have had limited success with topical or oral phage therapies in the treatment of biofilm infections and therefore little interest is present for these methods beyond topical application for burns and wounds [24–26]. Given the novelty of this therapeutic there is a paucity of data with respect to pharmacokinetics of local administration of bacteriophage therapy to biofilms. No data are present to suggest how long locally administrated bacteriophage reside at the infection site, how much is systemically absorbed or the safety of this approach. On the other hand, intravenous bacteriophage therapy has been more widely used and data are present to help guide treatment protocols. Therefore discussion about simple pharmacokinetics is limited to intravenous use.

Distribution: Bacteriophages are expected to be diluted in the whole body volume when given intravenously [27]. In numerous animal studies the titers of bacteriophages after intravenous infusions can be reduced 100–100,000-fold within 30 min of infusion [27]. Animal studies have shown distribution to various other organs including but not limited to heart, lungs, brain, skeletal muscle, bone marrow, and genitourinary tract [27]. However there are no data on intravenous bacteriophage therapy distribution to joints, spinal hardware, Left ventricular assist devices (LVADs) or other spaces that could have poor vascularization.

Metabolism: This is the chemical modification of bacteriophage therapy to reduce its activity. With bacteriophage therapy this occurs mainly through inactivation by the human immune system by neutralizing antibodies [28]. Neutralizing antibody responses have occurred with all forms of bacteriophage administration and this is a theoretical concern for long duration bacteriophage therapies [28,29].

Elimination: Elimination occurs mainly through hepatic clearance. In 1969, using a T4 bacteriophage, Inchley demonstrated that the liver phagocytosed and eliminated more than 99% of the bacteriophages within 30 min after systemic administration [30]. Other studies have supported this extensive hepatic elimination [30–33]. In one compassionate use case, 50 min after intravenous administration no bacteriophage could be detected in patient's serum [34].

Based on these data, intravenous bacteriophage therapies are likely to have significant reduction in titers, secondary to volume of distribution and hepatic elimination. Therefore, achieving MOIs similar to what occurs with in vitro susceptibility testing may be difficult. With the use of bacteriophage therapy applied directly to biofilm infections, MOIs may be similar to what was observed with in vitro susceptibility testing. However, limited pharmacological data have been found to help direct dosing, duration or safety of local bacteriophage administration.

### *2.3. Safety*

Unbeknownst to most, humans are exposed to low titers of bacteriophages on a continual basis [35]. Eastern European medicine has used bacteriophage therapy for close to 100 years with few significant adverse reactions being reported [36]. However, given the extensive hepatic clearance, western medicine is entertaining the use of high titers (greater than 109) of intravenous bacteriophage therapy and direct injection of high titers of bacteriophages directly to biofilm infections. Limited safety studies have been conducted using these techniques beyond a phase 1 clinical trial evaluating a three-bacteriophage cocktail with titers of 1 × 10<sup>9</sup> plaque-forming unit (PFU) twice a day for 14 days to *Staphylococcus aureus* bacteremia [37].

While intravenous bacteriophage therapy has been used in the past with limited adverse events, recent compassionate use cases have shown two adverse events [38,39]. One occurred in the treatment of chronic pseudomonas left ventricular assist device (LVAD) infection in which no success occurred with low titers of intravenous bacteriophage therapy and subsequent bacteriophage therapy with high titers of 1 × 10<sup>11</sup> PFU induced fever, shortness of breath and wheezing [38]. These symptoms resolved with supportive medical care but continued with repeat dosing with the same titers. When titers were diluted to 1 × 1010 PFU, the authors document that no adverse events occurred [38]. Endotoxin units were well below the United States Food and Drug Administration approved limit. In the other case, a significant transaminitis occurred after three doses of daily intravenous bacteriophage therapy with titers of 2.7 × 10<sup>9</sup> PFU in the treatment of a recalcitrant methicillin-resistant *Staphylococcus aureus* prosthetic joint infection. No causative etiology other than bacteriophage therapy could be found [39]. After cessation of bacteriophage therapy liver function returned to normal after 14 days. These two cases suggest that an upper limit may exist with respect to the titers that can be intravenously infused without exposing patients to potential adverse events. However only further safety trials with high titers given intravenously or directly to sites of biofilm infections will be able to assess safety and if there is a ceiling for the amount of titers that can be given.

### *2.4. Resistance Development*

With longer bacteriophage therapies, concern arises for the development of resistance. Resistance usually occurs from bacterial modifications of cell surface receptors or down regulation of receptors used in phage–bacteria attachment [40,41]. Other means of resistance can occur through adaptive systems such as the CRISPR–Cas9 system that cleaves phage DNA thus not allowing for progeny phage to be created [40]. Means to overcome or prevent resistance from occurring include use of cocktails of bacteriophages and bacteriophage substitutions. Bacteriophage cocktails are a group of different bacteriophages that theoretically use different attachment receptors. Therefore, if resistance develops to one receptor, the cocktail should continue to be effective. A recent study showed the frequency of spontaneous induction of resistance to a cocktail of three *Staphylococcus aureus* bacteriophages was no greater than 3 × 10−<sup>9</sup> [42]. Bacteriophage substitutions are simply changing therapy to a different bacteriophage that has lytic activity to the bacterium.

Bacteriophage resistance can occur rapidly causing formation of resistant variants that are immune to further bacteriophage infection [41]. This could impede effectiveness of bacteriophage therapy but resistance may also come at a cost to the bacterium especially when antimicrobial agents are present [41]. Moreover, bacteriophage-resistant bacteria often lack important surface features that are responsible for bacterial virulence [41]. Nonetheless resistance is an important factor that should be accounted for especially with prolonged bacteriophage treatments. In a case series of 10 intravenous bacteriophage only cases, resistance occurred in a significant portion of patients and required bacteriophage substitutions [38]. Resistance development is dependent on complex interplays of MOIs, growth inhibition durations and other bacteriophage–bacteria interactions [41]. Therefore resistance might develop rapidly or slower depending on these complex interactions. Determining in vitro resistance development to a clinical isolate is not routinely conducted, but could be easily assessed with susceptibility testing by evaluating the bacterial overgrowth for resistant variants. Resistance development is an important parameter that can have ramifications on efficacy and reproducibility of treatment protocols. Therefore it might be prudent to routinely test for and standardize what an acceptable level of in vitro resistance development is for different infectious processes to reduce further problems of reproducibility and improve efficacy of treatment protocols.

### *2.5. Synergistic or Antagonistic Activity with Antibiotics*

While resistance is an important parameter so is compatibility with systemic antibiotics which may have synergistic or antagonistic activity with bacteriophage therapy. Theoretically, antibiotics that inhibit protein synthesis (rifampin, tetracyclines, linezolid and others) can inhibit phage gene expression and therefore be antagonistic [43]. Antibiotics that inhibit cell wall synthesis inhibitors such as beta-lactams are potentially more synergistic [43]. These findings have been reinforced in numerous in vitro studies [43]. It has also been documented that concentrations of antibiotics also have important ramification of synergistic or antagonistic activity with higher antibiotic concentrations tending to be more antagonistic compared to lower concentrations which tend to be more synergistic [43]. It is interesting that in vivo studies have shown more synergistic activity of antibiotics with bacteriophage therapy then antagonism [15–23]. Spatial and temporal interactions of antibiotics and bacteriophages in vivo likely account for this synergistic activity [43]. It should be reinforced that biofilm bacteria reside where there is poor vascularization and therefore very low concentrations of systemic antibiotics may make phage–antibiotic compatibility less of an issue in vivo [43]. Testing forin vitro for phage–antibiotic compatibility is not commonly conducted. As with resistance development and susceptibility testing, it may be prudent to ensure bacteriophage–antibiotic compatibility with the systemic antibiotics that are planned to be used thus allowing for more reproducible treatment protocols.

### *2.6. Clinical Biofilms*

In vitro bacteriophage biofilm studies are traditionally conducted in static environments. These studies are usually devoid of human plasma proteins, lack in vivo stressors and normally remove planktonic infections before experiments are conducted. However, in vivo, planktonic infections overly clinical biofilm infections and are what causes the majority of the symptoms that patient's experience. Without eradicating these planktonic bacteria, bacteriophage therapy protocols will have to account for the planktonic infection and the biofilm infection. This adds complexity to the use of bacteriophage therapy and potentially further hinders reproducibility given the heterogeneity of these planktonic infections.

Other clinical factors that should be considered include: stability of infected hardware and importance of manual debridement of biofilms. Stability of hardware must be assessed to ensure retaining these materials is possible. This is usually conducted by imaging but manual inspection and manipulation is the most advantageous way to assess the ability to retain these materials. Manual debridement of biofilms has also been shown to be synergistic with respect to bacteriophage activity in biofilms [16,17]. This synergistic activity is likely a result of better bacteriophage penetration into biofilm and exposing biofilm bacteria to bacteriophages [16,17].

### *2.7. Conclusions*

The parameters discussed show that currently bacteriophage therapy is not a therapeutic that can be used similarly to conventional antibiotics. In addition relying mainly on bacteriophage's ability to self-replicate is unlikely to be beneficial beyond isolated case reports given the complexity of these other parameters. Rather thoughtful consideration of many parameters needs to be considered to devise effective, reproducible treatment protocols. Most of these parameters discussed are intertwined but standardization is lacking. Given the heterogeneity of these parameters glaring issues of reproducibility are present at this nascent stage of bacteriophage therapy. To reduce these reproducibility issues standardizing some of these parameters might be needed which include: minimal duration of growth inhibition, resistance testing, bacteriophage–antibiotic compatibility and ensuring in vitro bacteriophage biofilm activity. This may allow for more rigorous testing of reproducible protocols to therefore definitively determine if this therapeutic has efficacy in treating clinical biofilm infections.

### **3. Current Theoretical Bacteriophage Protocols for Chronic Biofilm Infections**

Many recent compassionate use cases have been conducted recently to treat clinical biofilm infections (prosthetic joint infections, LVAD infections, vascular graft infections and others). Two main approaches are being used in western medicine which include: intravenous bacteriophage therapy and the use of surgical interventions to directly inject bacteriophages to site of the biofilm. Table 1 discusses the advantages and disadvantages to each approach in relation to the parameters discussed above. Both approaches involve adjuvant bacteriophage therapy in combination with standard of care systemic antibiotics. Further review of recent case reports with respect to the different approaches is discussed here.

### *3.1. Case Studies of Intravenous Bacteriophage Therapy in Biofilm Infections*

Intravenous bacteriophage therapy has been attempted to treat prosthetic joint infections, ventricular assist devices, vascular graft infections and other hardware infections [38,44–46]. In one case series, the authors describe the use of bacteriophage therapy for two *Pseudomonas* LVAD infections with prolonged intravenous bacteriophage therapies with unsuccessful outcomes [38]. The same author also treated a *Staphylococcus aureus* LVAD infection with intravenous bacteriophage therapy and the was documented as a success, but the patient continued to have culture positive *Staphylococcus aureus* infection at the time of LVAD explant, suggesting the inability of intravenous bacteriophage therapy to eradicate the biofilm infection [38,44]. Another case report treated a *Klebsiella pneumonia* prosthetic

joint infection with 8 weeks of intravenous bacteriophage therapy with improvement of symptoms [45]. However the patient remains on chronic indefinite oral suppression antibiotics, limiting the ability to assess if eradication of the clinical biofilm was achieved.

Intravenous bacteriophage therapy is optically attractive in that no surgery is needed. However at the present time no case report has definitively shown the ability to eradicate clinical biofilms with this approach. The theoretical concern with intravenous bacteriophage therapy alone is the entrapment of bacteriophages in the planktonic infection limiting exposure of bacteriophages to the biofilm bacteria. In correlation, bacteriophages are extensively cleared by the liver and in vivo biofilms are usually poorly vascularized. Therefore achieving theoretical MOIs seen with in vitro "susceptibility" testing requires very high titers of infused bacteriophages. These high titers may be limited by potential adverse reactions [38,39]. Further complicating intravenous only therapies is the need for prolonged durations driven by limited bacteriophages reaching their bacterial biofilm targets thereby leading to the development of resistance and neutralizing antibodies. These variables when combined cause numerous confounding variables that may cause significant issues of reproducibility at this nascent stage of bacteriophage therapy.


**Table 1.** Advantages and disadvantages of intravenous and direct injection of bacteriophage therapy for clinical biofilm infections.

### *3.2. Case Studies of Direct Injection of Bacteriophages to Biofilms with Surgical Intervention*

The addition of bacteriophage therapy with debridement and irrigation surgeries is the other approach that has been used in several case studies [38,47–50]. This approach involves injection of high titers of bacteriophage phages directly at the site of the biofilm infection thereby circumventing hepatic clearance. The goal of this approach is to potentially cure these infections without need for removal of the hardware. However the risks of a surgical procedure are present and therefore this approach is optically less desirable.

There have been several compassionate use case reports that have shown successful eradication of biofilm infections which include: chronic orthopedic hardware infections, LVAD infection, vascular graft infection, cardiothoracic surgery infections [38,47–50]. In these case reports different durations of bacteriophage therapy were used. Some cases only required single doses given at the time of surgery while others used drains to continually instill bacteriophage therapy for 7 to 10 days. All cases described no recurrence of bacterial infections while patients were off antimicrobial therapies thereby showing eradication of bacterial biofilms. However with surgical debridement, successful eradication of bacterial biofilm is confounded by the uncertainty of whether it was bacteriophage therapy that was the reason for clearance or if it was the surgical intervention itself. This questioning occurs because success occurs with these surgical debridement procedures without adjuvant bacteriophage therapy albeit at low rates. In addition, limited data are present to help direct safety, appropriate dosing and durations of directly administered bacteriophage therapies but this approach may allow for more standardized reproducible protocols.

### **4. Conclusions**

Many aspects of bacteriophage therapy allow this therapeutic to be an attractive adjuvant therapeutic in the treatment of biofilm infections but much work is needed before definitive efficacy trials are to be conducted. The various parameters discussed here should allow researchers to be more cognizant of the current inherent limitations of bacteriophage therapy. However, given the heterogeneity of these parameters, projected issues of reproducibility are glaring. Therefore, standardizing some of these parameters is warranted to formulate reproducible protocols that will allow for rigorous testing of this therapeutic in the treatment of biofilm infections.

**Funding:** This research received no external funding.

**Acknowledgments:** Phagomed are acknowledged for providing and authorization of the use of Figure 2.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

© 2020 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Bacteriophage Cocktail-Mediated Inhibition of** *Pseudomonas aeruginosa* **Biofilm on Endotracheal Tube Surface**

**Viviane C. Oliveira 1,2 , Ana P. Macedo 2, Luís D. R. Melo <sup>3</sup> , Sílvio B. Santos <sup>3</sup> , Paula R. S. Hermann 1,4, Cláudia H. Silva-Lovato 2, Helena F. O. Paranhos 2, Denise Andrade <sup>1</sup> and Evandro Watanabe 1,5,\***


**Abstract:** Although different strategies to control biofilm formation on endotracheal tubes have been proposed, there are scarce scientific data on applying phages for both removing and preventing *Pseudomonas aeruginosa* biofilms on the device surface. Here, the anti-biofilm capacity of five bacteriophages was evaluated by a high content screening assay. We observed that biofilms were significantly reduced after phage treatment, especially in multidrug-resistant strains. Considering the anti-biofilm screens, two phages were selected as cocktail components, and the cocktail's ability to prevent colonization of the endotracheal tube surface was tested in a dynamic biofilm model. Phage-coated tubes were challenged with different *P. aeruginosa* strains. The biofilm growth was monitored from 24 to 168 h by colony forming unit counting, metabolic activity assessment, and biofilm morphology observation. The phage cocktail promoted differences of bacterial colonization; nonetheless, the action was strain dependent. Phage cocktail coating did not promote substantial changes in metabolic activity. Scanning electron microscopy revealed a higher concentration of biofilm cells in control, while tower-like structures could be observed on phage cocktail-coated tubes. These results demonstrate that with the development of new coating strategies, phage therapy has potential in controlling the endotracheal tube-associated biofilm.

**Keywords:** bacteriophage; biofilm; *Pseudomonas aeruginosa*; endotracheal tube

### **1. Introduction**

Ventilator-associated pneumonia (VAP) is a serious concern in critically ill patients occurring within the 48 h period following endotracheal intubation. The current COVID-19 pandemic is a predisposing factor for VAP, since it often requires mechanical ventilation, thus increasing the incidence and relevance of this infection [1]. VAP frequently involves high morbidity and excessive healthcare costs, and its incidence increases with the duration of ventilation [2–4]. The role of the endotracheal tube-associated biofilms in VAP etiology has been largely discussed. Commonly, biofilms on the device surface appear rapidly after intubation, promote a global covering of the internal side, and remain attached even after suctioning [2,5]. These biofilms represent a persistent source of pathogenic bacteria that can invade the lower airways, colonizing the lungs and causing VAP [6].

Strategies for biofilm growth inhibition on the endotracheal tube (ET) surface involve mainly suction systems [7], mucus shavers [8], and antimicrobial coatings [9–12]. Regarding

**Citation:** Oliveira, V.C.; Macedo, A.P.; Melo, L.D.R.; Santos, S.B.; Hermann, P.R.S.; Silva-Lovato, C.H.; Paranhos, H.F.O.; Andrade, D.; Watanabe, E. Bacteriophage Cocktail-Mediated Inhibition of *Pseudomonas aeruginosa* Biofilm on Endotracheal Tube Surface. *Antibiotics* **2021**, *10*, 78. https://doi.org/10.3390/ antibiotics10010078

Received: 1 December 2020 Accepted: 12 January 2021 Published: 15 January 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

biofilm removal, well-established strategies were not previously described in scientific literature, and direct administration of aerosolized antibiotics [13], cationic peptides [14], and ionized gas [15] have been suggested to control mature biofilm. Nonetheless, the available methods for both inhibiting and removing biofilm are not widely effective in controlling the microorganism layers on the ET surface, and innovative approaches to treat or prevent this contamination source should be investigated.

Even though the ET colonization is polymicrobial, the aerobic nosocomial bacterium *Pseudomonas aeruginosa* has been suggested to play a dominant role in the infection etiology [5,16,17]. In addition, *P. aeruginosa* has shown an enhanced ability to form huge biofilms and to develop antibiotic resistance, which in turn can be considered factors that ensure persistent infection [18,19]. Efforts have been made to reduce complications associated with *P. aeruginosa* colonization in artificial airways; nonetheless, none of them seem to be largely effective [3,20,21].

Considering the successful use of phage therapy in the treatment of *P. aeruginosa* acute respiratory infection in animal models [22], the use of phages is a promising and challenging alternative to deal with the ET-associated biofilms, mainly those formed by antibiotic-resistant strains. The advantages of using phage therapy involve low damage to the host microbiota, ability to self-replicate in the presence of host cells, host specificity, rapid selection and characterization, and low cost [23]. Aiming at treating acute infections caused by nosocomial pathogens, Aleshkin et al. reported that intragastric administration of a phage cocktail in patients with mechanical ventilation promoted an important reduction in bacterial burden [24]. Furthermore, the anti-biofilm activity of recently characterized new phages was demonstrated in vitro in an ET-associated *P. aeruginosa* biofilm model [25]. The authors demonstrated an extensive lytic activity with multidrug-resistant *P. aeruginosa* biofilm, suggesting that the new phages might be considered as good candidates for therapeutic studies [25]. Although encouraging results have revealed the anti-biofilm effect of the phage therapy, the action of immobilized phages on the ET surface, to control biofilm development, is unclear.

Along these lines, it is essential to clarify whether phages could be used for both controlling and preventing ET-associated *P. aeruginosa* biofilm. In the present study, antibiofilm activity of five recently characterized phages was evaluated by a high content screening assay. Subsequently, two phages were selected as cocktail components and applied as a preventive strategy to inhibit bacteria colonization in a dynamic biofilm model simulating endotracheal intubation. The null hypothesis of this study was that there is no difference in *P. aeruginosa* biofilm when challenged with bacteriophages.

### **2. Results**

### *2.1. Screening Phages for Anti-Biofilm Activity*

An initial screening was performed to select phages with stronger anti-biofilm activity. Biofilm-covered areas showed a significant reduction after phage treatment in 4/15 *P. aeruginosa* strains (Table S1), in which three were classified previously as multidrug-resistant [25]. Biofilm areas of four other *P. aeruginosa* strains were lower, but the difference was not significant. Even though phage infectivity had been previously determined, seven *P. aeruginosa* strains were not affected by the phage treatment. The analysis of the biofilm-covered areas indicated a statistically significant difference (*p* < 0.05) between phage-treated and control; however, statistically significant differences were not observed among the five different phages (Figure 1A). Therefore, based on the broader lytic spectrum of the phages with multidrug-resistant strains; the efficiency of plating and genomic differences, reported by Oliveira et al. [25]; and the anti-biofilm activity presented here (Figure 1B–G), the phages vB\_PaeM\_USP\_2 and vB\_PaeM\_USP\_18 were selected to compose a cocktail in the assays involving dynamic biofilm growth on the ET surface.

**Figure 1.** (**A**) Biofilm-covered areas, expressed in μm2, after phage treatment. Comparisons were conducted among groups by means of multiple comparisons considering strains and bacteriophages in a generalized linear model with Bonferroni correction. AB Different capital letters indicate statistically significant differences (*p* < 0.05). (**B**–**G**) Representative fluorescent images of *P. aeruginosa* illustrate the control group (**B**) and the action of phages vB\_PaeM\_USP\_1 (**C**), vB\_PaeM\_USP\_2 (**D**), vB\_PaeM\_USP\_3 (**E**), vB\_PaeM\_USP\_18 (**F**), and vB\_PaeM\_USP\_25 (**G**). Scale bar = 50 μm.

### *2.2. Replication of ET Adsorbed Phage During Biofilm Growth*

The phage cocktail that was adsorbed to the ET clearly showed the ability to replicate, since the number of phage particles increased over time. The initial log 3 phage population (0 h) was able to replicate in the presence of the biofilm cells, increasing to log 6 at 24 h and log 8 at 48 h. After 48 up to 168 h, phage concentration remained almost constant without variations among the strains (Figure 2A–C).

**Figure 2.** Phage cocktail presence on tube surfaces over 24 to 168 h of dynamic biofilm growth. (**A**) *P. aeruginosa* ATCC 27853; (**B**) *P. aeruginosa* ATCC 2110; (**C**) *P. aeruginosa* ATCC 2112.

### *2.3. Phage Cocktail Effect on P. aeruginosa Biofilms*

The in vivo contamination of an ET was mimicked using a continuous biofilm model system. Biofilm growth rates on non-coated tubes were similar among the three strains. However, on phage-coated tubes, a different growth pattern among the strains was observed (Table S2). This outcome indicated that the cocktail's action was strain dependent. Regarding metabolic activity, phage cocktail coating did not promote substantial changes in the biofilm response. Generally, the absorbance values were lower at early stages and higher in late stages of cultivation time (Figures 3B, 4B and 5B; Table S3).

**Figure 3.** Biofilm growth (**A**) and metabolic activity (**B**) of *P. aeruginosa* ATCC 27853 over 24 to 168 h of dynamic biofilm growth on non-coated and phage cocktail-coated tubes. Comparisons were conducted among groups, at each time point, by means of multiple comparisons considering strains and phage cocktail treatment in a generalized linear model with Bonferroni correction. \* indicates statistically significant difference at each time point (*p* < 0.05).

Comparing the CFU values, *P. aeruginosa* ATCC 27853 (Figure 3A) showed a significant reduction of the microbial load on phage cocktail-coated tubes at 24 (1.8 log; *p* < 0.001) and 120 h (0.9 log; *p* = 0.035) of treatment. Even though the CFU values at 48, 72, and 96 h of treatment indicated a slight biofilm reduction on phage cocktail-coated tubes (ranging from 0.1 to 0.6 log), the microbial load did not significantly differ from non-phage coated tubes.

In comparison to control, *P. aeruginosa* ATCC 27853 had higher metabolic activity on phage cocktail-coated tubes at 72 h and lower at 168 h of culture (Figure 3B).

Regarding *P. aeruginosa* ATCC 2110, significant reduction of the microbial load was observed only at 48 h (1 log; *p* = 0.004) of treatment (Figure 4A). The strain exhibited lower metabolic activity on phage cocktail-coated tubes at 72 h of culture in comparison to non-coated tubes (Figure 4B).

**Figure 4.** Biofilm growth (**A**) and metabolic activity (**B**) of *P. aeruginosa* ATCC 2110 over 24 to 168 h of dynamic biofilm growth on non-coated and phage cocktail-coated tubes. Comparisons were conducted among groups, at each time point, by means of multiple comparisons considering strains and phage cocktail treatment in a generalized linear model with Bonferroni correction. \* indicates statistically significant difference at each time point (*p* < 0.05).

The reduction of *P. aeruginosa* ATCC 2112 on phage cocktail-coated tubes ranged from 1.1 to 1.8 log (Figure 5A) during the entire treatment period (*p* ≤ 0.001). Nonetheless, no difference was observed in the evaluation of the metabolic activity (Figure 5B).

**Figure 5.** Biofilm growth (**A**) and metabolic activity (**B**) of *P. aeruginosa* ATCC 2112 over 24 to 168 h of dynamic biofilm growth on non-coated and phage cocktail-coated tubes. Comparisons were conducted among groups, at each time point, by means of multiple comparisons considering strains and phage cocktail treatment in a generalized linear model with Bonferroni correction. \* indicates statistically significant difference at each time point (*p* < 0.05).

SEM representative biofilm images of *P. aeruginosa* ATCC 2112 for all the cultivation times are shown in Figure 6. The microscopy images of phage cocktail-coated tubes were morphologically distinct from non-phage coated ones. A higher concentration of biofilm cells was noticed covering the tube surface in the control, while tower-like structures could be observed on phage cocktail-coated tubes. In the control tubes, the biofilm grew like a homogeneous layer, while on coated tubes the highest number of cells was observed in the clusters. In addition, on control tubes an extracellular polymeric matrix covered the entire biofilm layer. On phage cocktail-coated tubes, the extracellular polymeric matrix was detected as merely covering the tower-like structures, while in the surrounding areas less matrix and fewer isolated bacteria could be observed during the entire cultivation time.

### **3. Discussion**

In the current study, we assessed whether a phage treatment could efficiently reduce ET-associated biofilm. Firstly, the action of five different phages in a mature biofilm was evaluated. Subsequently, considering the lytic spectrum with multidrug-resistant strains, anti-biofilm screenings, and different efficiency of plating (EOP), two phages were selected as cocktail components and were applied as a strategy to prevent bacterial colonization and biofilm formation. Based on the results, the null hypothesis was rejected, since there were statistical differences for *P. aeruginosa* ET-associated biofilms.

According to biofilm-covered areas, our results demonstrated that phages applied solely exhibited effective anti-biofilm activity against a variety of *P. aeruginosa* strains. Nonetheless, biofilm-covered areas of seven *P. aeruginosa* strains remained unaffected when challenged with phages. Such an observation is in line with another in vitro study that showed varying degrees of biofilm disruption after phage treatment [22] and may be explained by the following reasons: (I) The efficiency of plating of the evaluated phages on five of these strains was considered low according to previously reported data [25]; (II) The antiviral mechanisms developed by the bacteria against phage adsorption, infection, and replication. The bacterial resistance systems have been extensively discussed in the scientific literature [26,27]; (III) Both treatment period and phage dosing might have been insufficient. According to Abedon, elimination of biofilms using phage therapy can require long treatment periods as well as repeated dosing [28]. Here, a single dose and treatment time was evaluated; and (IV) Staining methods and imaging tools have been considered useful for quantitative assessment and spatial structure visualization of biofilm; however, they can also result in misinterpretation of data due to laser penetration, absorption of the dye into the biomass, and auto-fluorescence [29,30]. LIVE/DEAD staining comprises two types of fluorescent stains, which differ in ability to penetrate viable and non-viable bacterial cells [31]. Here, the biofilm-covered areas were calculated according to total image fluorescence. It is known that cell concentration in biofilms can be distributed differently according to their thickness. Our image series may not have precisely recorded the biofilm density, which could in part explain unapparent anti-biofilm effects. Since agar plate counts detect all cultivable cells, the conflicting phage anti-biofilm results, reported by Oliveira et al., could be explained by the method used. The authors demonstrated superior phage anti-biofilm activity, against the same strains, by using colony forming unit counts [25]. Therefore, we consider that additional quantitative methods involving the determination of the number of viable cells by agar plate counts, flow-based cell counting, and assessment of biofilm dry mass or total protein content could lead to efficient determination of the biofilm density.

After describing the phage isolated action, a cocktail composed of vB\_PaeM\_USP\_2 and vB\_PaeM\_USP\_18 was investigated as an additional strategy for biofilm control on the ET surface. We hypothesized that surface coating using multiple phage strains prevents bacterial colonization considering that these two phages have different EOP, distinct lytic spectra with multidrug-resistant strains, and considerable genetic differences that could potentially lead them to bind to different receptors [32].

Efforts have been made to propose methods for phage coating in medical devices [33– 35]. For indwelling urological devices, the phage-coating is usually obtained by physical adsorption [33,36] and hydrogel conjugation [34,37,38]. The ETs used in this study were manufactured from reinforced polyvinyl chloride (PVC), and the scientific literature describes neither phage immobilization on PVC surfaces nor on other devices designed for mechanical ventilation. In this sense, we allowed physical adsorption of 1 × 10<sup>7</sup> PFU/cm<sup>2</sup> to create an antimicrobial surface. After 24 h, the phage immobilized on the tube surface was 1 × 10<sup>3</sup> PFU/cm2. The physical adsorption did not promote a large phage immobilization on the ET surface. We consider that the limited anti-biofilm effect, observed in the cocktail-coated tubes, was possibly due to the reduced phage attachment. It might be expected that by maximizing the density of the phages on the ET surface, an enhanced capacity to control *P. aeruginosa* growth would be reached. Even though the physical

adsorption comprises a simple and cost-effective method for phage immobilization, the low coverage seems to be unsuitable for producing a largely anti-biofilm effect. In this sense, different studies have been proposed aiming at the development of functionalized surfaces to immobilize phages efficiently. For instance, Wang, Sauvageau, and Elias exhibited that on the plasma-treated polyhydroxyalkanoate surface, the immobilization of phage T4 was greater than on the non-treated surface [39]. Therefore, investigation of different technologies for attaching phages to PVC surfaces, as well as phages displaying plastic-binding peptides, should be performed to ensure a high phage concentration on the ET surface.

Although an initial high phage titer was not immobilized on the tube surface, in the presence of bacterial strains an increasing concentration was observed after 48 h of treatment, which confirmed the phages' ability to replicate and compensate for the initial low dose [40]. Additionally, the 1 × 103 PFU/cm<sup>2</sup> was able to produce differences of bacterial colonization. According to mean differences at each specific time point, *P. aeruginosa* ATCC 27853 and *P. aeruginosa* ATCC 2112 showed evident reduction of biofilm growth on phage cocktail-coated tubes in the early stages of biofilm formation. This result can be related to the biofilm formation stage and the amount of extracellular exopolysaccharide matrix. The scientific literature has shown that inefficiency in phage penetration in mature biofilms is an important factor affecting the tolerance to phages [28]. The initial reduction in biofilm growth could be correlated both to the exponential increase of phage titer and thinner extracellular matrix layer. After 48 h, the constant titer of phages supports the idea that the thicker extracellular matrix could have hindered phage adsorption. On the other hand, a similar pattern was not observed for *P. aeruginosa* ATCC 2112, which exhibited a reduction of biofilm growth during the entire cultivation time. This distinct response can be associated with the metabolic activity of *P. aeruginosa* ATCC 2112. In comparison to *P. aeruginosa* ATCC 27853 and *P. aeruginosa* ATCC 2110, XTT assay revealed high absorbance values for *P. aeruginosa* ATCC 2112. As phages require metabolically active hosts to replicate [41], the cocktail could have more effectively infected this strain. In general, the lowest metabolic activity was observed for *P. aeruginosa* ATCC 2110, which exhibited reduction in biofilm growth only at 48 h. Taken together, these findings suggest that *P. aeruginosa* ATCC 2110 exhibits antiviral mechanisms that result in a phage-insensitive phenotype. Blocking of phage receptors, production of competitive inhibitors, prevention of bacteriophage DNA entry, slicing of bacteriophage nucleic acids, CRISPR/cas system activation, and abortive infection mechanisms are well-known bacterial resistance systems against phage infection [26,27].

The synthetic sputum medium used promotes the formation of *P. aeruginosa* aggregates with sizes similar to those observed in human cystic fibrosis lung tissue [42]. Here, however, coated and non-coated ETs had differences regarding distribution of aggregates. We suggest two different reasons to explain the formation of *P. aeruginosa* aggregates. First, on coated tubes, the formation of large bacterial aggregates, observed in SEM images, seemed to be a protection mechanism against phage invasion as it became more evident in the mature biofilms when phages reached the highest titer. Second, we speculate that this phenomenon may be caused not by the overgrowth of bacteria in some regions but by the lysis of cells by the phage cocktail causing holes on the biofilm. Structures similar to those observed here were also reported by Henriksen et al., who classified them as a defense strategy against phage infection [43]. According to the authors, the continuous phage exposure affected the biofilm growth by stimulating the formation of a highly organized and spatially heterogeneous structure.

Our results did not provide evidence regarding the different phage infection behavior of antibiotic sensitive and resistant strains. Phages have the demonstrated ability to infect both sensitive and multidrug-resistant *P. aeruginosa*. Loc-Carrillo and Abedon pointed out that resistance mechanisms against antibiotics do not affect phage infection [23]. Our results corroborate the author's statement and indicate that phage therapy could be applied as an auxiliary method to treat infections caused by resistant bacteria. Moreover, recombinant phage-encoded enzymes could be applied directly to the tube surface as an alternative to direct phage usage.

The relevance of the present study highlights the urgent need to investigate new therapeutic strategies to control *P. aeruginosa* biofilms on the ET surface. The intubation period through an ET for ≥ 8 days represents a risk factor for VAP occurrences [4]. Here, we demonstrated that phage therapy can reduce bacterial bioburden on the ET surface and therefore might contribute to reducing VAP episodes. Nonetheless, in view of the discrepant titers applied to both strategies in the study, biofilm treatment and biofilm prevention, we were unable to determine whether the phages would be more efficient in the treatment or prophylaxis of *P. aeruginosa* biofilms. Indeed, challenges and limitations of phage therapy are evident, and the scientific literature has reported that the therapeutic or prophylactic use of phages is dependent on the application area. For instance, in the food industry, prophylactic phage administration represents a promising sustainable solution to control pathogenic bacteria and reduce the massive use of antibiotics. In this field, phages are mainly used during food production, sanitization, and preservation [44]. For both animal and human infection treatment, the therapeutic use of phages and phage-encoded enzymes, alone or in combination with antibiotics, has aroused a growing interest in their potential use against multidrug-resistant bacteria, and different routes of administration and dosage effect have been suggested [45]. In order to reduce biofilm growth on implantable medical devices, we consider that immobilization of phages or phage-encoded products, as preventive agents, might decrease colonization more effectively than using them for biofilm removal. Thus, some issues remain and should be addressed in future studies. Experimental ventilator-associated pneumonia models and preclinical assessment would be useful to clarify if the biofilm removal/inhibition promoted by phages could prevent or reduce the severity of the VAP.

### **4. Materials and Methods**

### *4.1. Bacterial Strains, Growth Conditions, and Bacteriophages*

All bacterial strains used in this study are listed in Table 1. Bacteria were thawed and routinely grown in tryptic soy broth (TSB; BD Difco, Sparks, MN, USA) at 37 ◦C with agitation. After achieving the exponential growth phase, the culture was centrifuged (4200× *g*, 5 min) and washed twice in phosphate buffered saline (PBS), pH 7.4. The bacteria inoculum was prepared considering its optical density (OD625 nm) measured in a spectrophotometer (Thermo Scientific, Waltham, MA, USA).


**Table 1.** Bacterial strains used in the study.

† Human Exposome and Infectious Diseases Network collection. ‡ American Type Culture Collection. \* S: Susceptible to all antimicrobial agents tested; AMC: amoxicillin-clavulanic acid; AMK: amikacin; AMP: ampicillin; CFZ: cefazolin; CPD: cefpodoxime; CRO: ceftriaxone; CTX: cefotaxime; CXM: cefuroxime; FOX: cefoxitin; GEN: gentamicin; IMP: imipenem; NIT: nitrofurantoin; SAM: ampicillin-sulbactam; SXT: trimethoprim-sulfamethoxazole; TET: tetracycline; TGC: tigecycline. 173

Five bacteriophages were used in this study: vB\_PaeM\_USP\_1, vB\_PaeM\_USP\_2, vB\_PaeM\_USP\_3, vB\_PaeM\_USP\_18, and vB\_PaeM\_USP\_25. The isolation, characterization, and assessment of the lytic spectrum of the bacteriophages was described previously by Oliveira et al. [25].

### *4.2. Screening Phages for Anti-Biofilm Activity*

The anti-biofilm activity of the bacteriophages was utilized against 10 clinical isolates and five strains from the American Type Culture Collection (ATCC; Table 1). Two hundred microliters of TSB containing standardized bacteria suspension (10<sup>7</sup> colony forming units per milliliter—CFU/mL) was cultured (37 ◦C, 75 rpm) in black 96-well plates with a flat glass bottom (Corning, New York, NY, USA). After 24 h, half of the culture medium was removed, and the biofilm was supplied with freshly prepared culture medium, then plates were incubated for another 24 h. The culture medium was then discarded, and 200 μL of sterile TSB supplemented with 108 plaque forming units per milliliter (PFU/mL) of bacteriophages were added to each well (6 × 105 PFU/mm2). Culture medium without bacteriophages was used as a control. The plates were incubated for 24 h at 37 ◦C and 75 rpm.

To evaluate the anti-biofilm activity of the bacteriophages, the culture medium was discarded, and the wells were rinsed with 200 μL of PBS. The biofilm was stained for 15 min, protected from light, with LIVE/DEAD™ Biofilm Viability Kit (Molecular Probes, California, CA, USA) according to the manufacturer's protocol. Afterwards, the plates were scanned, and images were randomly collected (considering peripheral and central regions) with an Operetta CLS High-Content imaging system (PerkinElmer Waltham, MA, USA) at 40× magnification with 15 fields of view/well. The biofilm-covered areas (μm2) were then analyzed using Harmony High Content Imaging and Analysis Software (PerkinElmer, Version 4.8, MA, USA). The assay was conducted in triplicate.

### *4.3. Phage Cocktail Pretreatment of Endotracheal Tube Surfaces*

Under sterile conditions, the two ends of the ET (8.5 mm diameter, 300 mm length; Rüsh, Meridian, MS, USA) were removed to allow its connection to the tubing of the dynamic biofilm system. A phage cocktail containing 4 × 10<sup>7</sup> (PFU/mL) of vB\_PaeM\_USP\_2 and vB\_PaeM\_USP\_18 was prepared in elution buffer (SM) (1 M Tris HCl pH 7.5, Sigma-Aldrich, Saint Louis, MO, USA; 8 mM MgSO4, Sigma-Aldrich; 100 mM NaCl, Dinâmica, Indaiatuba, SP, Brazil; 0.002% (*w*/*v*) gelatin, Dinâmica). Seventeen milliliters of the suspension was added to the inner part of the tube. Then, the extremities were sealed, and the device was maintained under static conditions for 24 h at room temperature. This step was employed to allow phage adsorption to the ET surface. Afterwards, the suspension was discarded, and the tube was rinsed with SM buffer in order to remove unbound phages [33]. Two fragments of 1 cm2 were removed from the tube to assess the presence of bacteriophages, and the flow system was mounted under sterile conditions, as demonstrated in Figure 7.

### *4.4. Developing Biofilms on Endotracheal Tube Pretreated with the Phage Cocktail*

Three *P. aeruginosa* strains (ATCC 27853, ATCC 2110, and ATCC 2112) were selected for this assay. The strains ATCC 2110 (resistant to ampicillin, cefazolin, cefotaxime, cefoxitin, nitrofurantoin, trimethoprim-sulfamethoxazole, and tetracycline) and ATCC 2112 (resistant to amoxicillin-clavulanic acid, ampicillin, cefazolin, cefpodoxime, ceftriaxone, cefotaxime, cefuroxime, cefoxitin, nitrofurantoin, trimethoprim-sulfamethoxazole tetracycline, and tigecycline) were chosen due to their multidrug-resistant characteristics. The strain ATCC 27853 was selected due to the absence of antibiotic resistance. Each strain was evaluated solely in triplicate.

To simulate in vivo conditions, SCFM2 artificial sputum medium (4 g DNA salmon sperm, GoldBio, St Louis, MO, USA; 5 g swine stomach mucin, Sigma-Aldrich; 5 g casamino acids, Difco; 5.9 mg diethylenetriamine pentaacetic acid, Sigma-Aldrich; 5 g NaCl, Dynamic; 2.2 g KCl, Dynamic; 5 mL egg yolk emulsion; and 1000 mL distilled water, pH = 6.9) that

mimics a cystic fibrosis model was employed [42]. After connecting to the flow system, phage cocktail-coated and non-coated tubes were supplied with a continuous flow of SCFM2 culture medium inoculated with 1 × 10<sup>5</sup> CFU/mL of *P. aeruginosa* strains for 24 h. After 24 up to 168 h, the system was supplied with sterile SCFM2 culture medium without recirculation [33].

**Figure 7.** Schematic representation of the dynamic biofilm system showing that biofilm growth was monitored over 24 up to 168 h for colony forming units, metabolic activity, and biofilm morphology. Phage titer was also assessed during the entire cultivation time.

### *4.5. Analysis of the Phage Cocktail Effect on P. aeruginosa Biofilms*

The phage cocktail's ability to prevent ET colonization was determined by means of biofilm growth rates (CFU/cm2), metabolic activity of the biofilm (XTT), and scanning electron microscopy (SEM) at 24, 48, 72, 96, 120, 144, and 168 h under dynamic conditions. Therefore, at each time point, two fragments of 1 cm<sup>2</sup> of each tube (*n* = 3) were removed for CFU counts (*n* = 6) and XTT assessment (*n* = 6). For biofilm morphology evaluation (SEM), one representative fragment was processed. The CFU and XTT methodology were performed as described previously by Oliveira et al. [25].

For CFU quantification, each fragment was transferred to a tube containing 10 mL of phosphate buffered saline (PBS). The tubes were vortexed for 60 s, sonicated (200 W, 40 kHz; Altsonic, Clean 9CA, Ribeirão Preto, SP, Brazil) for 20 min and vortexed again for 2 min to ensure detachment of all aggregated biofilm. Ten-fold dilution aliquots were seeded in tryptic soy agar (BD Difco) and incubated at 37 ◦C for 24 h. The number of colonies was registered and expressed as log10CFU/cm2.

For the evaluation of metabolic activity, the strains were transferred to 24-well plates containing: 948 μL PBS supplemented with 100 mM glucose (Sigma-Aldrich), 240 μL XTT 1 mg/mL (Sigma-Aldrich), and 12 μL 0.4 mM menadione (Sigma-Aldrich). The plates were incubated, protected from light at 37 ◦C for 2 h, and the OD492 nm of the resulting solution was measured in triplicate. The mean of the readings was calculated subtracting the background absorbance.

For SEM analysis, the fragments were fixed with 2.5% glutaraldehyde (*v*/*v*) for 24 h and then dehydrated in a graded ethanol series (30%, 50%, 70%, 90%, and 100% (*v*/*v*)). After chemical drying using hexamethyldisilazane (Sigma-Aldrich), the specimens were mounted on an aluminum specimen holder and gold coated. The surface morphology of the biofilms was examined at a magnification of 3000× under high vacuum with a scanning electron microscope (EVO 10, CARL ZEISS, Jena, Germany).

### *4.6. Replication of ET Adsorbed Phage During Biofilm Growth*

The replication (infection ability) of phages from the cocktail that were adsorbed on the ET surface was confirmed, at all the time points (from 0 to 168 h), by double-layer-agar plating (tryptic soy agar soft (0.8% agar)—TSAS; BD Difco) [46]. In brief, the suspension employed for CFU quantification was centrifuged and diluted in SM buffer (100–10<sup>−</sup>6). Ten microliters were dropped onto a TSAS medium, with *P. aeruginosa* lawns, and incubated at 37 ◦C for 24 h. After the incubation period, the phage titer (PFU/mL) was determined by the number of phage plaques observable on the TSAS.

### *4.7. Statistical Analysis*

The adherence of the data to normal distribution (Shapiro–Wilk test) and homogeneous variance (Levene test) was tested. The data set did not exhibit normal distribution and were analyzed by multiple comparisons considering strains and bacteriophages, at specific time points, in a generalized linear model with Bonferroni correction. Comparisons among time points were not conducted in view of significantly phenotypic changes in biofilm growth. The statistical tests were performed through the IBM SPSS Statistics 25.0 software (IBM Corp Armonk, NY, USA). The significance level was set to 0.05.

### **5. Conclusions**

This study is the first step toward enhancing our understanding of biofilm growth in phage-coated ETs. The observed reduction depicts a favorable result but is not enough, suggesting that phages may be used not as an alternative but as a complementary strategy to control biofilms on ET, which can be improved with a better immobilization method. Since this low number of adherent phages caused significant changes in treatment, even better results are expected with an increased immobilization method. Furthermore, special attention should be paid to the potential development of phage resistance mechanisms, since over time phage treatment favors phage-insensitive phenotype development.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/2079-6 382/10/1/78/s1, Table S1: Biofilm-covered areas (μm2) of *Pseudomonas aeruginosa* strains after 24 h in the presence of five different bacteriophages. Table S2: Colony forming units (log10CFU/cm2) of different *Pseudomonas aeruginosa* strains after 24, 48, 72, 96, 120, 144, and 168 h of culture in continuous flow, on the surface of endotracheal tubes, in the presence and absence of bacteriophage cocktail. Table S3: Metabolic activity (absorbance at 492 nm) of different *Pseudomonas aeruginosa* strains after 24, 48, 72, 96, 120, 144, and 168 h of culture in continuous flow, on the surface of endotracheal tubes, in the presence and absence of bacteriophage cocktail.

**Author Contributions:** Conceptualization, V.C.O. and E.W.; data curation, C.H.S.-L., H.F.O.P. and D.A.; formal analysis, A.P.M. and P.R.S.H.; funding acquisition, E.W.; investigation, V.C.O.; methodology, V.C.O. and A.P.M.; project administration, E.W.; supervision, E.W.; validation, L.D.R.M. and S.B.S.; visualization, L.D.R.M. and S.B.S.; writing—original draft, V.C.O.; writing—review and editing, E.W., C.H.S.-L., H.F.O.P., L.D.R.M. and S.B.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by the São Paulo Research Foundation (FAPESP) under the grants 2018/09757-0, 2019/13271-9 and 2020/03405-5, as well as the National Council for Scientific and Technological Development (CNPq) under the grant 405622/2018-0.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


*Article*
