Bacteriophage-Delivering Hydrogels: Comparison
View Latest Version
Please note this is a comparison between Version 2 by Camila Xu and Version 1 by Rachel Yoon Chang.

Hydrogels are non-toxic polymeric materials exhibiting three-dimensional networks along with their hydrophilic characteristics playing an essential role in containing large water content, which serves as a biocompatible environment suited for formulation and delivery of bacteriophages.

  • bacteriophage (phage)
  • hydrogel
  • multidrug-resistant bacteria

1. Introduction

Bacteria are developing resistance against commercial antibiotics at an alarming rate, and antibiotic resistance is now one of the biggest threats to global health, contributing to rise in morbidity and mortality [1,2][1][2]. Indiscriminate use of antibiotics has led to the emergence of multidrug-resistant (MDR) bacteria with increased pathogenicity [3]. Thus, novel therapeutics are urgently needed to address the consequence of MDR infections. Bacteriophage (phage) therapy is being reconsidered as a potential alternative or adjunctive therapy to conventional antibiotics due to its ability to treat infections associated with MDR bacteria [4]. Phages are viruses that specifically target bacteria without hindering commensal microbiome [1,5][1][5]. Obligatory lytic (virulent) phages are utilized in phage therapy. Lytic phages inject their genetic material into the bacteria upon receptor recognition, self-replicate and then burst release their progenies during bacteriolysis [1,6][1][6]. Unlike some antibiotics, which are known to elicit more severe side effects in the patient than the infection itself [7[7][8],8], phage therapy is considered generally safe with no severe side effects reported in humans [1]. Other advantages include the ability to co-administer phages with antibiotics to induce synergistic antimicrobial effect [9,10,11][9][10][11] and to penetrate bacterial biofilms both in vitro and in vivo [12[12][13][14],13,14], further expanding the role of phage therapy. Biofilm formation on medical devices has posed significant problems to healthcare systems not only due to the emergence of MDR pathogens but also the antibacterial shielding effect of extracellular polymeric substances [15]. Phages produce enzymes which degrade extracellular polymeric substances, thereby being able to target persistent bacteria that are difficult to kill with antibiotics [16]. Therefore, phage therapy holds promising potential to help ease the burden of MDR bacterial infections. Considering its favorable antibacterial effects, phage therapy has been developed and approved as a standard medical application in Russia, the Republic of Georgia and other Former Soviet Union countries for many decades [17]. Moreover, phage therapy has undergone at least four Phase 1 and 2 clinical trials in the last 10 years to further expand its application to reach the market in the West [6]. Clinical case studies have investigated direct application in liquid formulation of phages to sinuses, wound and ear infection sites [18,19,20,21][18][19][20][21]. Although promising, liquid formulations lacking controlled delivery of phages can be therapeutically limited by the sudden release and rapid dispersion and/or elimination of phages from the desired microenvironment.

Formulation of phages involve a dual challenge of ensuring phage biostability and physical stability of the formulation (solution, suspension, gel or powder). Phages are, in a sense, large protein complexes enclosing genetic materials (DNA or RNA) and are only partially stable in solution, like most proteins. Naturally, protein stabilization strategies need to be considered, and they have in fact been applied in formulating phage therapeutics [22]. To become a viable therapeutic product, both native structure and biological activity of phages in the formulations must be retained during production and storage. Individual phages may have different stability profiles even in the same formulation [23,24[23][24][25],25], adding further complexity. To date, the development of stable phage formulations for therapeutic purposes is still an underexplored area of research. Hydrogels have been used as a vehicle to enable controlled delivery or administration of biologics such as phages to the target site of interest, including wounds [26][27][28] [26,27,28] and implants [29,30,31][29][30][31]. Hydrogels are non-toxic polymeric materials exhibiting three-dimensional networks along with their hydrophilic characteristics playing an essential role in containing large water content, which serves as a biocompatible environment suited for biological molecules [32,33][32][33]. Moreover, hydrogels resemble living tissues by holding a high proportion of water content within its matrix, providing optimal environment for accommodating proteins, living cells and other biomolecules, hence expanding its application in biomedical field [33,34][33][34]. In addition, hydrogel system enables controlled release of drugs through their tunable physical properties and biodegradability [35], which is also applicable to biomolecules delivery [36]. Considering the favorable characteristics for incorporating biological agents, hydrogels are a promising vehicle for the delivery of phages.

Exploiting the benefits of both phages and hydrogels, phage hydrogels have been utilized to treat and/or prevent MDR bacterial infections. A growing number of in vitro and in vivo preclinical studies indicate that hydrogels could be an ideal phage delivery system.

2. Stability of Phages in Hydrogels

Phage stability is an important factor to consider, as this corresponds to the viability of phages over time, which ultimately affects their bactericidal effect. The stability of phages is dependent on many factors, including pH, temperature, formulation composition and light exposure [61]. Phages, being the active pharmaceutical ingredient, must remain biologically stable in the developed formulation over the desired storage time. However, only a few studies have investigated the storage stability of phage hydrogels. In two separate studies, Kumari et al. have shown that Klebsiella phage Kpn5 remained stable in a 3% HPMC hydrogel when stored at 37 °C for seven days [27,28][27][28]. Similarly, phage LM99 in Alg and Alg-nanoHA hydrogel was biologically stable for seven days (storage temperature unreported) [29]. In another study, Klebsiella phage Kpn5, Pseudomonas phage PA5 and Staphylococcus phage MR10 remained viable in PVA-SA hydrogel when stored at room temperature for 28 days [26]. However, complete inactivation of some phages has been reported in PVA hydrogel, which was thought to be attributed to highly damaging radical species [48].

The field lacks well-controlled long-term storage stability studies of phage hydrogels. In general, neutral polymers are recommended for formulating phages in hydrogels to minimize charge-induced inactivation of phages [62]. Phage capsid has an overall net negative charge and tail a net positive charge at physiological pH. Anionic polymers may unfavorably interact with positively charged tails through electrostatic interaction and block the tail fiber proteins responsible for bacteria binding, compromising phage infectivity. For example, phage formulated in Carbomer (anionic polymer) hydrogel resulted in 99.95% titer reduction within four weeks when stored at 4 °C, whereas those in hydroxyethylcellulose gel (non-ionic) hydrogel remained biologically stable [63]. As only a limited number of hydrogels have been tested for phage stability, in the future, studies on non-ionic hydrogels such as guar gum, agarose, polyethylene oxide, polyvinyl pyrrolidone, polyacrylamide, polycarbophil, poly(hydroxyethyl) methacrylate and hydroxypropyl cellulose could be done.

3. Efficacy of Phage-Delivering Hydrogels

3.1. Orthopedic Implant-Associated Infection

Orthopedic implant-associated infections are commonly observed among implanted surgical devices and can cause significant patient morbidity along with financial burden [30]. Virtually all materials used in implantable devices are readily colonized by bacteria and result in biofilm formation with increased resistance to the host immune system and antibiotics [64]. Current treatment options for bone infection are limited to the use of antibiotics and surgical debridement of affected tissue, often followed by implant removal [30]. The use of systematic antibiotics in bone infection is often associated with poor delivery to the site of infection with nephrotoxic and hepatotoxic adverse effects at high dose [65]. To locally deliver MDR bacteria-killing phages, injectable hydrogels have been explored in vivo and in vitro to treat orthopedic implant-associated infections (Figure 2).

Antibiotics 10 00130 g002 550

Figure 2. Topical application of phage-delivering hydrogel formulation as a wound dressing (A), injectable phage-delivering hydrogel formulation via syringe (B) and phage-delivering formulation via a hydrogel-coated catheter (C).

Wroe et al. demonstrated that ΦPaer14-encapsulating PEG-4-MAL hydrogel significantly reduced formation of P. aeruginosa biofilm (17-fold reduction in colony-forming unit (CFU) counts) as compared with control gels in vitro. Fluorescence staining confirmed that control gels showed a higher load of live bacteria and biofilm-associated proteins, whereas the phage hydrogel-treated group showed higher levels of dead bacteria. Furthermore, phage hydrogel formulated with protease-degradable peptide linkers exhibited rapid killing of planktonic cells in vitro [30] [30]. Antimicrobial efficacy of phage hydrogels can be modulated by utilizing faster degrading peptide sequence, which leads to rapid release of phages for potentially faster elimination of bacteria. In animals with radial segmental defects infected with P. aeruginosa, treatment with PEG-4-MAL hydrogel containing a cocktail of four Pseudomonas phages exhibited 4.7-fold reduction in bacterial counts (Table 2). Similarly, Johnson et al. applied PEG-4-MAL hydrogel containing both BMP-2 and phages on a mouse radial defect model for 8 weeks, resulting in a significant reduction (>1 log reduction) in the target P aeruginosa (PsAer-9) bacteria [41].

Table 2. Phage-delivering hydrogels for the prevention of orthopedic implant-associated bone infections.

Hydrogel Target Bacteria Phages Study Key Findings References
Alginate hydrogel, alginate nanohydroxyapatite hydrogel E. faecalis vB_EfaS_LM99 In vitro and in vivo (rabbit)

and P. mirabilis (Curtin/Donlan et al., Fu et al., Carson et al., Lehman/Donlan) can be incorporated into a hydrogel coating on a catheter and significantly reduce biofilm formation [53,54,55,56][53][54][55][56]. Over 2-log reduction in P. aeruginosa biofilm viability was observed upon pre-treatment of the catheter hydrogel with phage M4 [53]. Emergence of phage-resistant biofilm isolate was observed between 24 h and 48 h, but the use of a five-phage cocktail significantly suppressed this phenomenon. A study by Curtin and Donlan has shown that phage 456 incorporating a hydrogel-coated catheter can reduce S. epidermidis biofilm formation over a 24 h exposure with log CFU/cm2 reduction of 4.5 (Table 3). Interestingly, supplemental divalent cations along with phage 456 fostered further reduction in S. epidermidis cell attachment with log CFU/cm2 reduction of 2.3 as compared with phage-free controls [55]. Divalent cations such as Ca2+ or Mg 2+ are known for promoting growth or enhancing the antibacterial activity of phage [55,57,67][55][57][67]. Although phage-dependent, divalent cations can aid in phage attachment to host bacteria [49]. In gel formulations, divalent cations are used as crosslinking agents, yet these cations may not be freely available for interaction with phages. Hence, supplemental addition of cations after hydrogel formulation could be considered to enhance the antibacterial activity of phages.

Table 3. Summary of phage-delivering hydrogels in catheter-associated urinary tract infection.

Therapeutic Use Hydrogel Target Bacteria Phages Study Design Findings References
Preventing catheter-associated

UTI		Inhibited the attachment and colonization of MDR E. faecalis in femoral tissues. Inhibited growth (99.9%) of planktonic cells after 48 h. [29]
PEG-4-MAL hydrogel P. aeruginosa ΦPaer4,

ΦPaer14,

ΦPaer22,

ΦW2005A		 In vitro and in vivo (mouse) Reduced P. aeruginosa growth (4.7-fold) at the infection site after 7 days of implantation in mice [30]
Alginate hydrogel S. aureus Genetically modified phage In vitro and in vivo (rat) No significant reduction in bone infection. [31]

Note: PEG-4-MAL, polyethylene glycol-4-maleimide; MDR, multidrug-resistance.

In the study conducted by Barros et al., the efficacy of phages LM99 formulated in alginate hydrogel was examined in vitro and ex vivo against MDR Enterococcus faecalis isolated from orthopedic implant-associated infections [29]. Treatment with phage hydrogel reduced planktonic cells by 99% and bacterial attachment on hydrogels by 98% after 24 h of incubation. Antibacterial effect was also observed ex vivo with 99.9% reduction in CFU counts after 48 h of treatment with LM99 hydrogel formulation. Additionally, phage-free alginate-nanoHA hydrogel had osteogenic and mineralization response, suggesting that phage hydrogels are a promising multifunctional approach for controlling bacterial infection during implant and bone integration.

Despite promising in vitro and ex vivo data, only limited studies explored the efficacy of phage hydrogels for treatment of orthopedic implant-associated infections in vivo [30,31,41][30][31][41]. Treatment with fosfomycin and/or phage delivered using alginate hydrogel exhibited minimal or no significant antibacterial effect (<1 log reduction in all groups) in soft tissue and bone infection rat models, respectively. Lack of in vitro and in vivo correlation could be due to insufficient delivery and/or release of phages to exert therapeutic effect at the site of infection. The final titer of phage hydrogel was 3.0 × 107 PFU/mL and only 10 μL could fit into the small defect site (i.e., 3.0 × 105 PFU/mL). Hence, a higher initial dose of phages in hydrogels accompanied with controlled release from the gel matrix seems to be essential for in vivo efficacy. Moreover, the use of pig or sheep animal models that best mimic human bones may be more beneficial. In vivo studies often utilized co-administration of both bacteria and phage hydrogels, probably due to the complexity of surgical procedures. A more natural model would be to first inoculate bacteria at the site of interest to create a physical carrier (e.g., biofilms), followed by treatment with phage hydrogels. Further work is required to determine the antimicrobial effect in vivo of phage hydrogels in such chronic orthopedic implant-associated infection.

3.2. Catheter-Associated Urinary Tract Infection (CAUTI)

Catheters are commonly used indwelling device in healthcare facilities and are susceptible to biofilm formation, triggering catheter-associated urinary tract infection (CAUTI) [66]. To overcome CAUTI often associated with MDR pathogens, phage-delivering hydrogels are rising as one of the strategies to prevent and eradicate biofilms (Figure 2). Several in vitro studies have demonstrated that phages active against S. epidermidis, P. aeruginosa, E. coli

PEG-polyurethane hydrogel P. aeruginosa Pseudomonas phage cocktail: ΦPaer4, ΦPaer14, M4, 109, ΦE2005-A and ΦE2005-C In vitro Reduced formation of multi-species biofilm comprising P. aeruginosa (4 log) and P. mirabilis (2 log) in an artificial urine medium with 72 and 96 h exposure [56]
P. mirabilis Proteus phage cocktail: ΦPmir1, ΦPmir32, ΦPmir34, ΦPmir37
    S. epidermidis Phage 456 In vitro Reduced biofilm formation over a 24 h exposure with a log reduction of 4.47 [55]
    P. mirabilis

E. coli T4

Coli-proteus phage		 In vitro Reduced biofilm formation by approximately 90% [54]
    P. aeruginosa Phage M4 In vitro Reduced biofilm formation and bacterial attachment to catheter.

Phage cocktails on hydrogel-coated catheter reduction of 99.9% on biofilms composed of 11 variants after 48 h.		 [53]
Preventing encrustation of catheter lumen and catheter-associated

UTI		 PVA-

Eudragit		® S 100 hydrogel P. mirabilis Phage isolated from crude sewage In vitro Phage-delivering hydrogel reduced the P. mirabilis biofilm formation by 6 log reduction [43]

Note: PEG-polyurethane, polyethylene glycol-polyurethane; PVA, polyvinyl alcohol; UTI, urinary tract infection.

 

Note: PEG-polyurethane, polyethylene glycol-polyurethane; PVA, polyvinyl alcohol; UTI, urinary tract infection.

Multispecies biofilm can form on a catheter [68,69] and act as a stable reservoir of various pathogens that are difficult to eliminate. Lehman and Donlan [56] used a phage cocktail comprising six P. aeruginosa phages (1.0 × 10

Multispecies biofilm can form on a catheter [68][69] and act as a stable reservoir of various pathogens that are difficult to eliminate. Lehman and Donlan [56] used a phage cocktail comprising six P. aeruginosa phages (1.0 × 10

9

PFU/mL each) and/or four P. mirabilis phages (3.0 × 10

8 PFU/mL each) (Table 3) on a hydrogel-coated catheter to target single and multispecies biofilms. Treatment with Pseudomonas phage cocktail reduced single-species biofilm levels by 2.5 log after 24 h as compared with buffer-treated control, followed by regrowth at 48 h (1.5 log reduction). Antibiofilm activity of the cocktail was more pronounced against two-species biofilm with 3 log and 4 log reductions at 24 h and 48 h, respectively. Similarly, treatment with a Proteus phage cocktail reduced P. mirabilis populations in both single and multispecies biofilms. Interestingly, increased pH due to P. mirabilis urease activity caused elimination of P. aeruginosa by 72 h regardless of phage treatment, suggesting that observed efficacy against multispecies biofilms could be due to interplay of multiple factors. Increased pH caused by bacterial urease leads to supersaturation and precipitation of struvite and apatite crystals, forming crystalline biofilms that can block urinary catheters. A simple PVA-based phage hydrogel further formulated with pH-responsive polymer delayed the catheter blockage time (26 h) caused by P. mirabilis biofilms as compared with non-treated control (13 h) [43]. The majority of these studies used 24 h old biofilms, which are considered quite young. The efficacy against mature biofilms (≥48 h) with more complex and persistent extracellular matrix should be investigated in the future. Furthermore, adjunctive use of other anti-biofilm agents, such as antibiotics [70], nitric oxide [71,72] and amino acids [73,74], could be considered for a synergistic antibacterial effect for treating chronic infections.

PFU/mL each) (Table 3) on a hydrogel-coated catheter to target single and multispecies biofilms. Treatment with Pseudomonas phage cocktail reduced single-species biofilm levels by 2.5 log after 24 h as compared with buffer-treated control, followed by regrowth at 48 h (1.5 log reduction). Antibiofilm activity of the cocktail was more pronounced against two-species biofilm with 3 log and 4 log reductions at 24 h and 48 h, respectively. Similarly, treatment with a Proteus phage cocktail reduced P. mirabilis populations in both single and multispecies biofilms. Interestingly, increased pH due to P. mirabilis urease activity caused elimination of P. aeruginosa by 72 h regardless of phage treatment, suggesting that observed efficacy against multispecies biofilms could be due to interplay of multiple factors. Increased pH caused by bacterial urease leads to supersaturation and precipitation of struvite and apatite crystals, forming crystalline biofilms that can block urinary catheters. A simple PVA-based phage hydrogel further formulated with pH-responsive polymer delayed the catheter blockage time (26 h) caused by P. mirabilis biofilms as compared with non-treated control (13 h) [43]. The majority of these studies used 24 h old biofilms, which are considered quite young. The efficacy against mature biofilms (≥48 h) with more complex and persistent extracellular matrix should be investigated in the future. Furthermore, adjunctive use of other anti-biofilm agents, such as antibiotics [70], nitric oxide [71][72] and amino acids [73][74], could be considered for a synergistic antibacterial effect for treating chronic infections.

3.3. Trauma-Associated Skin and Soft Tissue Infection

Wounds from burn injuries provide a favorable environment for the growth of bacteria [27]. Burn injuries contribute to the fourth leading devastating form of trauma worldwide and may lead to death if untreated [26]. Topical application of antimicrobial agents is preferred over systemic antibiotics due to high bioavailability at the site of infection [28]. Hydrogels are commonly used in wound care products as they promote wound healing, maintain a hydrated environment essential for self-healing and clear debridement while absorbing the exudates [75]. Phages have been incorporated in hydrogels to treat bacterial wound infections (see Figure 2; their topical application has been extensively reviewed by Chang et al. [62]).

Phage K formulated in HAMA/agarose hydrogel system exhibited a clear zone of inhibition on S. aureus bacterial lawn [48]. Furthermore, temperature-sensitive PNIPAM-co-ALA hydrogel attached to non-woven polypropylene could deliver infective phage K and form a bacterial zone of clearance, indicating in vitro efficacy of the formulation (Table 4) [46]. In another study, treatment with PVA-SA hydrogel-based membrane containing phages MR10, Kpn5 and PA5 (all at a reported MOI of 10) resulted in 6 log reduction in S. aureus, 6.37 log reduction in Klebsiella pneumoniae and 4.6 log reduction in P. aeruginosa biomass, respectively, after 6 h in vitro (Table 4) [26]. Treatment with MR10 hydrogel resulted in fast wound healing in S. aureus burn wound infection model in mice as compared with non-treated control, particularly when minocycline was used in combination. On day 14, those receiving dual-agent treatment showed complete regeneration of skin layers, sweat glands and hair follicles similar to normal mouse skin. In addition to rapid wound healing, phage hydrogels significantly reduced mortality in mice burn wound infection model [28]. A study by Kumari et al. demonstrated that treatment with phage Kpn5 in HPMC hydrogel (MOI of 200) increased the survival rate of Klebisella-infected animals as compared with other antimicrobials, including silver nitrate and gentamicin (Table 4). All mice in the phage hydrogel treated group survived, while 87% survived in the non-treated group on day 1. On day 7, the phage-treated group showed the highest level of protection (63%) as compared with the untreated group (0%). The survival rate may be further increased by the combined use of phages and antibiotics.

Table 4. Summary of phage-delivering hydrogels in trauma-associated skin and soft tissue infection.

Therapeutic Use Hydrogel Target Bacteria Phages Study Design Findings References
Treating wound associated with burn injury HPMC hydrogel K. pneumoniae Kpn5 In vivo (mice) The highest survival rate compared to silver nitrate and gentamicin after 7 days [28]
PVA-SA hydrogel S. aureus

P. aeruginosa

K. pneumoniae MR10

PA5

Kpn5		 In vitro and in vivo (mice) Reduced resistant burn wound infection significantly (>1 log reduction) and exhibited reduction in inflammation with wound contraction. [26]
Treating skin infection Agarose-HAMA hydrogel S. aureus Phage K In vitro Triggered release of phage K under the presence of hyaluronidase, degrading the HAMA layer, and thus, inhibiting bacterial growth [48]
Treating skin and soft tissue infection PNIPAM-co-ALA hydrogel S. aureus Phage K In vitro PNIPAM-co-ALA nanogels attached to phage K exhibited thermally triggered bacterial lysis of S. aureus at 37 °C. [46]
Treating orthopedic implant-associated soft tissue infection Alginate hydrogel S. aureus Genetically modified phage In vitro and in vivo

(rat)		 Reduced soft tissue infection significantly

(>0.5 log reduction)		 [31]

Note: PVA-SA, polyvinyl alcohol-sodium alginate; HPMC, hydroxypropyl methylcellulose; PNIPAM-co-ALA, N-isopropylacrylamide-co-allylamine; HAMA, hyaluronic acid methacrylate.

 

Note: PVA-SA, polyvinyl alcohol-sodium alginate; HPMC, hydroxypropyl methylcellulose; PNIPAM-co-ALA, N-isopropylacrylamide-co-allylamine; HAMA, hyaluronic acid methacrylate.

Despite demonstrating prominent antibacterial activity, phage-delivering hydrogels were limited to targeting only one species in the context of combating wound infections. As wound infections are often polymicrobial [76], future studies should include phage cocktail consisting of prominent bacterial isolates in wound infections, such as P. aeruginosa, methicillin-resistant S. aureus and K. pneumoniae. Subsequent multispecies wound infection model should be developed and utilized to better reflect the clinical setting. Furthermore, formulation of hydrogels containing both phage and antibiotic should be considered to maximize bactericidal activity, while minimizing phage- and antibiotic-resistant isolates. Overall, most of the phage-delivering hydrogels demonstrated significant efficacy against MDR bacterial infection in vivo and in vitro. Aligned with efficacy, safety of phage-delivering hydrogels needs to be considered to further progress into human clinical trials.

References

  1. Górski, A.; Międzybrodzki, R.; Węgrzyn, G.; Jończyk-Matysiak, E.; Borysowski, J.; Weber-Dąbrowska, B. Phage therapy: cur-rent status and perspectives. Medicinal Research Reviews 2020, 40, 459-463, doi:https://doi.org/10.1002/med.21593.
  2. Balcão, V.M.; Moreira, A.R.; Moutinho, C.G.; Chaud, M.V.; Tubino, M.; Vila, M.M.D.C. Structural and functional stabiliza-tion of phage particles in carbohydrate matrices for bacterial biosensing. Enzyme and Microbial Technology 2013, 53, 55-69, doi:https://doi.org/10.1016/j.enzmictec.2013.03.001.
  3. Ventola, C.L. The antibiotic resistance crisis: part 1: causes and threats. P T 2015, 40, 277-283.
  4. Chang, R.Y.K.; Wallin, M.; Lin, Y.; Leung, S.S.Y.; Wang, H.; Morales, S.; Chan, H.K. Phage therapy for respiratory infections. Advanced Drug Delivery Reviews 2018, 133, 76-86, doi:10.1016/j.addr.2018.08.001.
  5. Huh, H.; Wong, S.; St Jean, J.; Slavcev, R. Bacteriophage interactions with mammalian tissue: therapeutic applications. Ad-vanced Drug Delivery Reviews 2019, 145, 4-17, doi:10.1016/j.addr.2019.01.003.
  6. Melo, L.D.R.; Oliveira, H.; Pires, D.P.; Dabrowska, K.; Azeredo, J. Phage therapy efficacy: a review of the last 10 years of preclinical studies. Critical Reviews in Microbiology 2020, 46, 78-99, doi:10.1080/1040841x.2020.1729695.
  7. Grill, M.F.; Maganti, R.K. Neurotoxic effects associated with antibiotic use: management considerations. Br J Clin Pharmacol 2011, 72, 381-393, doi:10.1111/j.1365-2125.2011.03991.x.
  8. Spapen, H.; Jacobs, R.; Van Gorp, V.; Troubleyn, J.; Honoré, P.M. Renal and neurological side effects of colistin in critically ill patients. Ann Intensive Care 2011, 1, 14-14, doi:10.1186/2110-5820-1-14.
  9. Lin, Y.; Chang, R.Y.K.; Britton, W.J.; Morales, S.; Kutter, E.; Chan, H.-K. Synergy of nebulized phage PEV20 and ciprofloxa-cin combination against Pseudomonas aeruginosa. International Journal of Pharmaceutics 2018, 551, 158-165, doi:10.1016/j.ijpharm.2018.09.024.
  10. Lin, Y.; Chang, R.Y.K.; Britton, W.J.; Morales, S.; Kutter, E.; Li, J.; Chan, H.K. Inhalable combination powder formulations of phage and ciprofloxacin for P. aeruginosa respiratory infections. European Journal of Pharmaceutics and Biopharmaceutics 2019, 142, 543-552, doi:10.1016/j.ejpb.2019.08.004.
  11. Lin, Y.; Quan, D.; Yoon Kyung Chang, R.; Chow, M.Y.T.; Wang, Y.; Li, M.; Morales, S.; Britton, W.J.; Kutter, E.; Li, J., et al. Synergistic activity of phage PEV20-ciprofloxacin combination powder formulation-A proof-of-principle study in a P. aeru-ginosa lung infection model. European Journal of Pharmaceutics and Biopharmaceutics 2021, 158, 166-171, doi:10.1016/j.ejpb.2020.11.019.
  12. Dakheel, K.H.; Rahim, R.A.; Neela, V.K.; Al-Obaidi, J.R.; Hun, T.G.; Isa, M.N.M.; Yusoff, K. Genomic analyses of two novel biofilm-degrading methicillin-resistant Staphylococcus aureus phages. BMC Microbiology 2019, 19, 114, doi:10.1186/s12866-019-1484-9.
  13. Fong, S.A.; Drilling, A.; Morales, S.; Cornet, M.E.; Woodworth, B.A.; Fokkens, W.J.; Psaltis, A.J.; Vreugde, S.; Wormald, P.-J. Activity of bacteriophages in removing biofilms of Pseudomonas aeruginosa isolates from chronic rhinosinusitis patients. Front Cell Infect Microbiol 2017, 7, 418-418, doi:10.3389/fcimb.2017.00418.
  14. Chang, R.Y.K.; Das, T.; Manos, J.; Kutter, E.; Morales, S.; Chan, H.K. Bacteriophage PEV20 and ciprofloxacin combination treatment enhances removal of Pseudomonas aeruginosa biofilm isolated from cystic fibrosis and wound patients. AAPS Journal 2019, 21, 49, doi:10.1208/s12248-019-0315-0.
  15. Gebreyohannes, G.; Nyerere, A.; Bii, C.; Sbhatu, D.B. Challenges of intervention, treatment, and antibiotic resistance of bio-film-forming microorganisms. Heliyon 2019, 5, e02192-e02192, doi:10.1016/j.heliyon.2019.e02192.
  16. Harper, D.R.; Parracho, H.M.R.T.; Walker, J.; Sharp, R.; Hughes, G.; Werthén, M.; Lehman, S.; Morales, S. Bacteriophages and biofilms. Antibiotics (Basel) 2014, 3, 270-284, doi:10.3390/antibiotics3030270.
  17. Abedon, S.T.; Kuhl, S.J.; Blasdel, B.G.; Kutter, E.M. Phage treatment of human infections. Bacteriophage 2011, 1, 66-85, doi:10.4161/bact.1.2.15845.
  18. Merabishvili, M.; Pirnay, J.P.; Verbeken, G.; Chanishvili, N.; Tediashvili, M.; Lashkhi, N.; Glonti, T.; Krylov, V.; Mast, J.; Van Parys, L., et al. Quality-controlled small-scale production of a well-defined bacteriophage cocktail for use in human clinical trials. PLoS One 2009, 4, e4944, doi:10.1371/journal.pone.0004944.
  19. Rhoads, D.D.; Wolcott, R.D.; Kuskowski, M.A.; Wolcott, B.M.; Ward, L.S.; Sulakvelidze, A. Bacteriophage therapy of venous leg ulcers in humans: results of a phase I safety trial. Journal of Wound Care 2009, 18, 237-238, 240-233, doi:10.12968/jowc.2009.18.6.42801.
  20. Wright, A.; Hawkins, C.H.; Anggård, E.E.; Harper, D.R. A controlled clinical trial of a therapeutic bacteriophage preparation in chronic otitis due to antibiotic-resistant Pseudomonas aeruginosa; a preliminary report of efficacy. Clinical Otolaryngology 2009, 34, 349-357, doi:10.1111/j.1749-4486.2009.01973.x.
  21. Ooi, M.L.; Drilling, A.J.; Morales, S.; Fong, S.; Moraitis, S.; Macias-Valle, L.; Vreugde, S.; Psaltis, A.J.; Wormald, P.J. Safety and tolerability of bacteriophage therapy for chronic rhinosinusitis due to Staphylococcus aureus. JAMA Otolaryngol Head Neck Surg 2019, 145, 723-729, doi:10.1001/jamaoto.2019.1191.
  22. Chang, R.Y.K.; Kwok, P.C.L.; Khanal, D.; Morales, S.; Kutter, E.; Li, J.; Chan, H.K. Inhalable bacteriophage powders: glass transition temperature and bioactivity stabilization. Bioengineering & Translational Medicine 2020, 5, e10159, doi:10.1002/btm2.10159.
  23. Clark, W.A. Comparison of several methods for preserving bacteriophages. Journal of Applied Microbiology 1962, 10, 466-471.
  24. Chang, R.Y.; Wong, J.; Mathai, A.; Morales, S.; Kutter, E.; Britton, W.; Li, J.; Chan, H.K. Production of highly stable spray dried phage formulations for treatment of Pseudomonas aeruginosa lung infection. European Journal of Pharmaceutics and Bio-pharmaceutics 2017, 121, 1-13, doi:10.1016/j.ejpb.2017.09.002.
  25. Chang, R.Y.K.; Wallin, M.; Kutter, E.; Morales, S.; Britton, W.; Li, J.; Chan, H.K. Storage stability of inhalable phage powders containing lactose at ambient conditions. International Journal of Pharmaceutics 2019, 560, 11-18, doi:10.1016/j.ijpharm.2019.01.050.
  26. Kaur, P.; Gondil, V.S.; Chhibber, S. A novel wound dressing consisting of PVA-SA hybrid hydrogel membrane for topical delivery of bacteriophages and antibiotics. International Journal of Pharmaceutics 2019, 572, 118779, doi:https://doi.org/10.1016/j.ijpharm.2019.118779.
  27. Kumari, S.; Harjai, K.; Chhibber, S. Topical treatment of Klebsiella pneumoniae B5055 induced burn wound infection in mice using natural products. Journal of Infection in Developing Countries 2010, 4, 367-377.
  28. Kumari, S.; Harjai, K.; Chhibber, S. Bacteriophage versus antimicrobial agents for the treatment of murine burn wound infec-tion caused by Klebsiella pneumoniae B5055. Journal of Medical Microbiology 2011, 60, 205-210, doi:10.1099/jmm.0.018580-0.
  29. Barros, J.A.R.; Melo, L.D.R.d.; Silva, R.A.R.d.; Ferraz, M.P.; Azeredo, J.C.V.d.R.; Pinheiro, V.M.d.C.; Colaço, B.J.A.; Fer-nandes, M.H.R.; Gomes, P.d.S.; Monteiro, F.J. Encapsulated bacteriophages in alginate-nanohydroxyapatite hydrogel as a novel delivery system to prevent orthopedic implant-associated infections. Nanomedicine: Nanotechnology, Biology and Medicine 2020, 24, 102145, doi:https://doi.org/10.1016/j.nano.2019.102145.
  30. Wroe, J.A.; Johnson, C.T.; García, A.J. Bacteriophage delivering hydrogels reduce biofilm formation in vitro and infection in vivo. Journal of Biomedical Materials Research Part A 2020, 108, 39-49, doi:10.1002/jbm.a.36790.
  31. Cobb, L.H.; Park, J.; Swanson, E.A.; Beard, M.C.; McCabe, E.M.; Rourke, A.S.; Seo, K.S.; Olivier, A.K.; Priddy, L.B. CRISPR-Cas9 modified bacteriophage for treatment of Staphylococcus aureus induced osteomyelitis and soft tissue infection. PLOS ONE 2019, 14, e0220421, doi:10.1371/journal.pone.0220421.
  32. Ahmed, E.M. Hydrogel: preparation, characterization, and applications: a review. Journal of Advanced Research 2015, 6, 105-121, doi:https://doi.org/10.1016/j.jare.2013.07.006.
  33. Pérez-Luna, V.H.; González-Reynoso, O. Encapsulation of biological agents in hydrogels for therapeutic applications. Gels 2018, 4, 61, doi:10.3390/gels4030061.
  34. Caló, E.; Khutoryanskiy, V.V. Biomedical applications of hydrogels: a review of patents and commercial products. European Polymer Journal 2015, 65, 252-267, doi:https://doi.org/10.1016/j.eurpolymj.2014.11.024.
  35. Narayanaswamy, R.; Torchilin, V.P. Hydrogels and their applications in targeted drug delivery. Molecules 2019, 24, 603, doi:10.3390/molecules24030603.
  36. Li, J.; Mooney, D.J. Designing hydrogels for controlled drug delivery. Nature Reviews Materials 2016, 1, 16071, doi:10.1038/natrevmats.2016.71.
  37. Lee, K.Y.; Mooney, D.J. Alginate: properties and biomedical applications. Progress in Polymer Science 2012, 37, 106-126, doi:https://doi.org/10.1016/j.progpolymsci.2011.06.003.
  38. Marrella, A.; Lagazzo, A.; Dellacasa, E.; Pasquini, C.; Finocchio, E.; Barberis, F.; Pastorino, L.; Giannoni, P.; Scaglione, S. 3D porous gelatin/PVA hydrogel as meniscus substitute using alginate micro-particles as porogens. Polymers (Basel) 2018, 10, 380, doi:10.3390/polym10040380.
  39. Cheng, Y.; Shi, X.; Jiang, X.; Wang, X.; Qin, H. Printability of a cellulose derivative for extrusion-based 3D printing: the ap-plication on a biodegradable support material. Frontiers in Materials 2020, 7, doi:10.3389/fmats.2020.00086.
  40. Jansen, L.E.; Negrón-Piñeiro, L.J.; Galarza, S.; Peyton, S.R. Control of thiol-maleimide reaction kinetics in PEG hydrogel net-works. Acta Biomaterialia 2018, 70, 120-128, doi:https://doi.org/10.1016/j.actbio.2018.01.043.
  41. Johnson, C.; Dinjaski, N.; Prieto, M.; García, A. Bacteriophage encapsulation in poly (ethylene glycol) hydrogels significantly reduces bacteria numbers in an implant-associated infection model of bone repair. Igarss 2014, 281.
  42. Liu, Y.; Vrana, N.E.; Cahill, P.A.; McGuinness, G.B. Physically crosslinked composite hydrogels of PVA with natural macro-molecules: structure, mechanical properties, and endothelial cell compatibility. J Biomed Mater Res B Appl Biomater 2009, 90, 492-502, doi:10.1002/jbm.b.31310.
  43. Milo, S.; Hathaway, H.; Nzakizwanayo, J.; Alves, D.R.; Esteban, P.P.; Jones, B.V.; Jenkins, A.T.A. Prevention of encrustation and blockage of urinary catheters by Proteus mirabilis via pH-triggered release of bacteriophage. Journal of Materials Chemistry B 2017, 5, 5403-5411, doi:10.1039/C7TB01302G.
  44. Elton, R.K. Flexible lubricious organic coatings. US5179174A, 12 January 1993.
  45. Zhou, G.; Ma, C.; Zhang, G. Synthesis of polyurethane-g-poly(ethylene glycol) copolymers by macroiniferter and their protein resistance. Polymer Chemistry 2011, 2, 1409-1414, doi:10.1039/C1PY00016K.
  46. Hathaway, H.; Alves, D.R.; Bean, J.; Esteban, P.P.; Ouadi, K.; Sutton, J.M.; Jenkins, A.T. Poly(N-isopropylacrylamide-co-allylamine) (PNIPAM-co-ALA) nanospheres for the thermally triggered release of bacterio-phage K. European Journal of Pharmaceutics and Biopharmaceutics 2015, 96, 437-441, doi:10.1016/j.ejpb.2015.09.013.
  47. Roach, B.; Nover, A.; Ateshian, G.; Hung, C. Agarose hydrogel characterization for regenerative medicine applications: focus on engineering cartilage. 2016; 10.1002/9781119126218.ch16pp. 258-273.
  48. Bean, J.E.; Alves, D.R.; Laabei, M.; Esteban, P.P.; Thet, N.T.; Enright, M.C.; Jenkins, A.T.A. Triggered release of bacteriophage K from Sagarose/hyaluronan hydrogel matrixes by Staphylococcus aureus virulence factors. Chemistry of Materials 2014, 26, 7201-7208, doi:10.1021/cm503974g.
  49. Chhibber, S.; Kaur, T.; Kaur, S. Essential role of calcium in the infection process of broad-spectrum methicillin-resistant Staphylococcus aureus bacteriophage. Journal of Basic Microbiology 2014, 54, 775-780, doi:10.1002/jobm.201300051.
  50. Bourdin, G.; Schmitt, B.; Marvin Guy, L.; Germond, J.-E.; Zuber, S.; Michot, L.; Reuteler, G.; Brüssow, H. Amplification and purification of T4-like escherichia coli phages for phage therapy: from laboratory to pilot scale. Appl Environ Microbiol 2014, 80, 1469-1476, doi:10.1128/AEM.03357-13.
  51. Phelps, E.A.; Enemchukwu, N.O.; Fiore, V.F.; Sy, J.C.; Murthy, N.; Sulchek, T.A.; Barker, T.H.; García, A.J. Maleimide cross-linked bioactive PEG hydrogel exhibits improved reaction kinetics and cross-linking for cell encapsulation and in situ delivery. Advanced Materials 2012, 24, 64-70, doi:https://doi.org/10.1002/adma.201103574.
  52. Foster, G.A.; Headen, D.M.; González-García, C.; Salmerón-Sánchez, M.; Shirwan, H.; García, A.J. Protease-degradable mi-crogels for protein delivery for vascularization. Biomaterials 2017, 113, 170-175, doi:10.1016/j.biomaterials.2016.10.044.
  53. Fu, W.; Forster, T.; Mayer, O.; Curtin, J.J.; Lehman, S.M.; Donlan, R.M. Bacteriophage cocktail for the prevention of biofilm formation by Pseudomonas aeruginosa on catheters in an in vitro model system. Antimicrobial Agents and Chemotherapy 2010, 54, 397-404, doi:10.1128/aac.00669-09.
  54. Carson, L.; Gorman, S.P.; Gilmore, B.F. The use of lytic bacteriophages in the prevention and eradication of biofilms of Proteus mirabilis and Escherichia coli. FEMS Immunology and Medical Microbiology 2010, 59, 447-455, doi:10.1111/j.1574-695X.2010.00696.x.
  55. Curtin, J.J.; Donlan, R.M. Using bacteriophages to reduce formation of catheter-associated biofilms by Staphylococcus epider-midis. Antimicrobial Agents and Chemotherapy 2006, 50, 1268-1275, doi:10.1128/aac.50.4.1268-1275.2006.
  56. Lehman, S.M.; Donlan, R.M. Bacteriophage-mediated control of a two-species biofilm formed by microorganisms causing catheter-associated urinary tract infections in an in vitro urinary catheter model. Antimicrobial Agents and Chemotherapy 2015, 59, 1127-1137, doi:10.1128/aac.03786-14.
  57. Rodney M. Donlan, S.M.L., Andres J. Garcia Controlled covalent attachment of biactive bacteriophage for regulating biofilm development. US9457132B2, 4 October 2016.
  58. Malik, D.J.; Sokolov, I.J.; Vinner, G.K.; Mancuso, F.; Cinquerrui, S.; Vladisavljevic, G.T.; Clokie, M.R.J.; Garton, N.J.; Stapley, A.G.F.; Kirpichnikova, A. Formulation, stabilisation and encapsulation of bacteriophage for phage therapy. Advances in Col-loid and Interface Science 2017, 249, 100-133, doi:10.1016/j.cis.2017.05.014.
  59. Merabishvili, M.; Vervaet, C.; Pirnay, J.P.; De Vos, D.; Verbeken, G.; Mast, J.; Chanishvili, N.; Vaneechoutte, M. Stability of Staphylococcus aureus phage ISP after freeze-drying (lyophilization). PLoS One 2013, 8, e68797, doi:10.1371/journal.pone.0068797.
  60. Miguel, S.P.; Ribeiro, M.P.; Brancal, H.; Coutinho, P.; Correia, I.J. Thermoresponsive chitosan–agarose hydrogel for skin re-generation. Carbohydrate Polymers 2014, 111, 366-373, doi:https://doi.org/10.1016/j.carbpol.2014.04.093.
  61. Jończyk-Matysiak, E.; Łodej, N.; Kula, D.; Owczarek, B.; Orwat, F.; Międzybrodzki, R.; Neuberg, J.; Bagińska, N.; We-ber-Dąbrowska, B.; Górski, A. Factors determining phage stability/activity: challenges in practical phage application. Expert Review of Anti-infective Therapy 2019, 17, 583-606, doi:10.1080/14787210.2019.1646126.
  62. Chang, R.Y.K.; Morales, S.; Okamoto, Y.; Chan, H.K. Topical application of bacteriophages for treatment of wound infec-tions. Translational research 2020, 220, 153-166, doi:10.1016/j.trsl.2020.03.010.
  63. Carbol, J.; Tan, P.; Varma, Y.; Osborne, D. Formulating topical products containing live microorganisms as the active ingre-dient. Pharmaceutical Technology Europe 2018, 42, 24-27.
  64. McConoughey, S.J.; Howlin, R.; Granger, J.F.; Manring, M.M.; Calhoun, J.H.; Shirtliff, M.; Kathju, S.; Stoodley, P. Biofilms in periprosthetic orthopedic infections. Future Microbiology 2014, 9, 987-1007, doi:10.2217/fmb.14.64.
  65. Caplin, J.D.; García, A.J. Implantable antimicrobial biomaterials for local drug delivery in bone infection models. Acta Bio-materialia 2019, 93, 2-11, doi:10.1016/j.actbio.2019.01.015.
  66. Nicolle, L.E. Catheter associated urinary tract infections. Antimicrob Resist Infect Control 2014, 3, 23-23, doi:10.1186/2047-2994-3-23.
  67. Ma, L.; Green, S.I.; Trautner, B.W.; Ramig, R.F.; Maresso, A.W. Metals enhance the killing of bacteria by bacteriophage in human blood. Scientific Reports 2018, 8, 2326, doi:10.1038/s41598-018-20698-2.
  68. Garibaldi, R.A.; Mooney, B.R.; Epstein, B.J.; Britt, M.R. An evaluation of daily bacteriologic monitoring to identify preventable episodes of catheter-associated urinary tract infection. Infection Control 1982, 3, 466-470, doi:10.1017/s0195941700056599.
  69. Tambyah, P.A.; Halvorson, K.T.; Maki, D.G. A prospective study of pathogenesis of catheter-associated urinary tract infec-tions. Mayo Clinic Proceedings 1999, 74, 131-136, doi:10.4065/74.2.131.
  70. Algburi, A.; Comito, N.; Kashtanov, D.; Dicks, L.M.T.; Chikindas, M.L. Control of biofilm formation: antibiotics and beyond. Appl Environ Microbiol 2017, 83, e02508-02516, doi:10.1128/aem.02508-16.
  71. Howlin, R.P.; Cathie, K.; Hall-Stoodley, L.; Cornelius, V.; Duignan, C.; Allan, R.N.; Fernandez, B.O.; Barraud, N.; Bruce, K.D.; Jefferies, J., et al. Low-dose nitric oxide as targeted anti-biofilm adjunctive therapy to treat chronic Pseudomonas aeru-ginosa infection in cystic fibrosis. Mol Ther 2017, 25, 2104-2116, doi:10.1016/j.ymthe.2017.06.021.
  72. Rouillard, K.R.; Markovetz, M.R.; Bacudio, L.G.; Hill, D.B.; Schoenfisch, M.H. Pseudomonas aeruginosa biofilm eradication via nitric oxide-releasing cyclodextrins. ACS Infectious Diseases 2020, 6, 1940-1950, doi:10.1021/acsinfecdis.0c00246.
  73. Kolodkin-Gal, I.; Romero, D.; Cao, S.; Clardy, J.; Kolter, R.; Losick, R. D-amino acids trigger biofilm disassembly. Science 2010, 328, 627-629, doi:10.1126/science.1188628.
  74. Hochbaum, A.I.; Kolodkin-Gal, I.; Foulston, L.; Kolter, R.; Aizenberg, J.; Losick, R. Inhibitory effects of D-amino acids on Staphylococcus aureus biofilm development. Journal of Bacteriology 2011, 193, 5616-5622, doi:10.1128/jb.05534-11.
  75. Merabishvili, M.; Monserez, R.; van Belleghem, J.; Rose, T.; Jennes, S.; De Vos, D.; Verbeken, G.; Vaneechoutte, M.; Pirnay, J.P. Stability of bacteriophages in burn wound care products. PLoS One 2017, 12, e0182121, doi:10.1371/journal.pone.0182121.
  76. Bessa, L.J.; Fazii, P.; Di Giulio, M.; Cellini, L. Bacterial isolates from infected wounds and their antibiotic susceptibility pattern: some remarks about wound infection. International Wound Journal 2015, 12, 47-52, doi:10.1111/iwj.12049.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/)
Video Production Service
Feedback