Open access peer-reviewed chapter

Chemotherapy and Mechanisms of Action of Antimicrobial Agent

Written By

Rahman Laibi Chelab

Submitted: October 3rd, 2020 Reviewed: December 12th, 2020 Published: June 9th, 2021

DOI: 10.5772/intechopen.95476

Chapter metrics overview

216 Chapter Downloads

View Full Metrics

Abstract

Pseudomonas aeruginosa is a widespread opportunistic pathogen that causes bloodstream, urinary tract, burn wounds infections and is one of the largest pathogens that infect cystic fibrosis patients’ airways and can be life-threatening for P. aeruginosa infections. In addition, P. aeruginosa remains one of the most significant and difficult nosocomial pathogens to handle. Increasingly, multi-drug resistance (MDR) strains are identified and the option of therapy is often very limited in these cases, particularly when searching for antimicrobial combinations to treat serious infections. The fact that no new antimicrobial agents are active against the MDR strains of P. aeruginosa is an additional matter of concern. In recent decades, bacterial drug resistance has increased, but the rate of discovery of new antibiotics has decreased steadily. The fight for new, powerful antibacterial agents has therefore become a top priority. This chapter illustrates and explores the current state of several innovative therapeutic methods that can be further discussed in clinical practice in the treatment of P. aeruginosa infections.

Keywords

  • P. aeruginosa
  • drug resistance
  • alternative therapies
  • vaccine
  • phage therapy

1. Introduction

We are currently facing an international crisis with many troublesome aspects: new antibiotics are no longer being detected, resistance mechanisms are developing in almost all clinical isolates of bacteria, and the effective treatment of infections is hampered by recurrent infections caused by persistent bacteria. Antibiotic failure is one of the most worrying health issues worldwide [1]. Although resistance acquisition is a natural phenomenon, it is accelerated by antibiotic misuse, inadequate inspection and poorly regulated management of antibiotics have resulted in the appearance and spread of multidrug-resistant (MDR) bacteria abroad in clinical medicine and in the livestock industry [2, 3].

Empirical antibiotic treatment requires monotherapy and combination therapy for suspected cases of P. aeruginosa and reduces mortality in patients with serious P. aeruginosa infections [4, 5]. However, because of the ability of this bacterium to avoid many of the currently available antibiotics, treatment of P. aeruginosa infections has become a great challenge [6]. Recently, the World Health Organization (WHO) has identified carbapenem-resistant P. aeruginosa as one of three bacterial species with an urgent need for new antibiotics to be developed to treat infections [7]. In addition, inappropriate treatment use of antibiotics accelerates the production of multidrug-resistant strains of P. aeruginosa, resulting in the ineffectiveness of empirical antibiotic therapy against this microorganism [8]. Resistance to a range of antibiotics, including aminoglycosides, quinolones and β-lactams, is demonstrated by P. aeruginosa [9]. Generally, the main mechanisms of P. aeruginosa used to fight antibiotic attack can be divided into intrinsic, acquired and adaptive resistance. Low external membrane permeability, the expression of efflux pumps that remove antibiotics from the cell, and the development of antibiotic inactivating enzymes are part of the intrinsic resistance of P. aeruginosa. Either horizontal transfer of resistance genes or mutational modifications will achieve the acquired resistance of P. aeruginosa [10]. P. aeruginosa ‘s adaptive resistance requires the development of biofilm in the lungs of infected patients, where the biofilm functions as a diffusion barrier to inhibit the access of antibiotics to bacterial cells [11].

The effectiveness and safety of murepavadin in the treatment of infections of the lower respiratory tract caused by P. aeruginosa (suspected or confirmed) in patients with ventilation-associated pneumonia or CF-unrelated bronchiectasis (Clinical Trials. gov identifiers NCT02096315 and NCT02096328) have been tested in two clinical trials. However, by July 17, 2019, the trials were stopped because in research participants who had obtained murepavadin, an unusually high level of renal failure had been found. This decision would not impact the production of an aerosolized formulation of murepavadin for topical use [12]. Murepavadin is a particular weapon against P. aeruginosa, which separates it from the broad pipeline of antimicrobial natural and synthetic peptides acting against multiple taxa, P. aeruginosa included. Recently, several novel peptides with broad antimicrobial activity have been identified, such as antimicrobial peptide DGL13K, Mel4 and melamine (Melimine and Mel4 are chimeric cationic peptides with broad-spectrum antimicrobial activity), Cecropin B, Lysine-based peptidomimetics (LBP-2), Truncated pseudin-2 analogs (Pse-T2), antimicrobial peptide, termed 6 K-F17 (sequence: KKKKKK-AAFAAWAAFAA-NH2), Melittin-derived peptides (MDP1, MDP2) [13, 14, 15, 16, 17, 18, 19, 20]. In addition, multidrug-tolerant persistent cells can form in the biofilm that are capable of surviving antibiotic attack; in cystic fibrosis (CF) patients, these cells are responsible for prolonged and recurrent infections [21]. For patients whose infections are resistant to traditional antibiotics, the development of new antibiotics or alternative therapeutic methods for treating P. aeruginosa infections is urgently needed. In recent years, new antibiotics with novel modes of action have been investigated, as have new routes of administration and resistance to bacterial enzyme alteration. Compared to traditional antibiotics, some of these newer antibiotics demonstrate excellent in vitro antibacterial activity against P. aeruginosa as well as lower minimum inhibitory concentration (MIC) [22, 23]. Moreover, several novel non-antibiotic therapeutic approaches that are highly successful in destroying antibiotic resistant P. aeruginosa strains have been documented in recent studies [24]. These approaches include: antimicrobial peptides, phage therapy, inhibition of quorum sensing, iron chelation, the use of nanoparticles, probiotic and vaccine strategy. In order to combat P. aeruginosa infections, these therapeutic approaches may be used either as an alternative to or in conjunction with traditional antibiotic therapies.

Advertisement

2. Mechanisms of action of antimicrobial agents

There are six basic mechanisms of antimicrobial agents presented below:

  1. Inhibition of microbial cell wall synthesis.

  2. Inhibition of microbial cell membrane function.

  3. Inhibition of microbial protein synthesis.

  4. Inhibition of microbial DNA synthesis.

  5. Inhibition of microbial RNA synthesis.

  6. Inhibition of microbial metabolic pathways.

2.1 Inhibition of microbial cell wall synthesis

An integral microbial structure responsible for the shape of the cell is the cell wall. In addition, because of the high cytoplasmic osmotic pressure, the cell wall prevents cell lysis and facilitates the anchoring of membrane components and extracellular proteins, such as adhesins [25]. Bacterial cell wall synthesis has perhaps become the target field most commonly exploited for antimicrobial production on the basis of the number of antimicrobial drugs in clinical usage. Due to the absence of equivalents in human biology, the components of the cell wall synthesis machinery are attractive antimicrobial targets, thus providing intrinsic objective selectivity. The cytoplasmic synthesis of building blocks composed of N-acetyl muramic acid (M) linked to N-acetyl glucosamine (G) with an attached pentapeptide (P) side chain (referred to as MGP subunits) comprises the sequential late steps in cell wall synthesis. The linkage of the MGP subunit to the lipid II molecule enables subsequent translocation to the outside or periplasmic space of the cell through the cytoplasmic membrane. By catalyzing glycosidic linkages between the M and G components of the MGP subunits, transglycosylase enzymes then assemble the MGP subunits into a linear backbone. An immature peptidoglycan structure is constituted by linearly connected MGP subunits. Transpeptidase enzymes then work to cross-link pentaglycine bridges to the peptide side chains, in the process, the terminal 2 D-alanines of the peptide side chain are cleaved, creating the mature, lattice-like peptidoglycan that provides the form and osmotic stability of the bacterium [26]. β-lactam antibiotics, such as penicillins and cephalosporins, are the most widely used antimicrobials that prevent cell wall biosynthesis [27]. These β-lactam antibiotics interact directly with bacterial transpeptidases and inhibit them effectively. As transpeptidase inhibitors, β-lactams thus obstruct the transition from immature to mature peptidoglycan, so these enzymes are also referred to as penicillin-binding proteins (PBPs). Due to the stereochemical similarity of the β-lactam moiety with the D-alanine-D-alanine substrate, they are capable of doing this. Transpeptidases form a lethal covalent penicilloyl enzyme complex in the presence of the drug, which helps to inhibit the usual transpeptidation reaction. This results in peptidoglycan that is weakly cross-linked, which makes the developing bacteria extremely susceptible to cell lysis and death [28].

2.2 Inhibition of microbial cell membrane function

Lipids, proteins and lipoproteins are essentially made of biological membranes. The cytoplasmic membrane for water, ions, nutrients and transport systems serves as a diffusion barrier. Most health workers now assume that membranes are a lipid matrix with uniformly distributed globular proteins to penetrate through the bilayer of the lipid. A number of antimicrobial agents may cause disorganization of the membrane. These agents can be categorized into cationic, anionic, and neutral agents. Polymyxin B and colistemethate (polymyxin E) are the best-known compounds [29]. For several antimicrobial agents, the cytoplasmic membrane forms an important barrier.

The mode of action of certain antimicrobial agents may be due to the ability of such medicines to increase membrane permeability, making it easier for them and other compounds to penetrate. Antibacterial cationic agents, increased permeability of the outer membrane to the lysozyme and hydrophobic compounds has been identified, such as polymyxin B. The initial function of these antimicrobial agents is to interrupt the structure of the outer membrane, allowing the cell to join itself and other compounds and inhibit unique metabolic processes [30]. There are several cell-damaging properties of Polymixin B: (i) the surface charge, lipid composition and membrane structure are disturbed; (ii) the K+ gradient on the cytoplasmic membrane is dissipated; and (iii) the cytoplasmic membrane is depolarized. One of the key factors regulating bacterial exposure to polymixin B is the permeability of the external membrane to lipophilic compounds. Since polymixin B is bulkier than its displacement of inorganic divalent cations, in the presence of polymixin B, the packing order of lipopolysaccharides (LPS) is changed. This results in increased permeability of a variety of molecules to the outer membrane and also promotes polymixin B uptake (“self-promoted” uptake) [31].

2.3 Inhibition of microbial protein synthesis

Microbial protein synthesis inhibition a range of groups of antimicrobial agents work by inhibiting the synthesis of bacterial proteins (ribosome function). That include aminoglycosides, macrolides, tetracyclines, ketolides, lincosamides, streptogramins, chloramphenicol and oxazolidinones [26, 32]. The synthesis of microbial proteins is led by ribosomes in conjunction with cytoplasmic factors which, during the initiation phase, elongation phase and termination phases, bind transiently to particles. Microbial ribosomes contain 70S particles consisting of two 50S and 30S subunits, which join at the initiation stage of the synthesis of proteins and split at the termination stage. In bacterial protein synthesis, antimicrobial agents block various steps by interfering with the work of either the cytoplasmic factors or the ribosomes. Inhibitors which bind to the ribosomal subunit of 30S primarily interfere with initiation, although some interfere with the pairing of the AA- tRNA anticodon with the mRNA codon, elongation is thus impaired. The steps involved in the elongation process interact with inhibitors that bind to the 50S ribosomal subunit or to elongation factors that are transiently connected to ribosomes at certain stages of the cycle.

Through binding to particular ribosomal subunits [33], aminoglycosides function. By inducing the development of aberrant, non-functional complexes as well as causing misreading, aminoglycoside-type drugs may combine with other binding sites on 30S ribosomes and destroy bacteria. Spectinomycin is an antimicrobial agent that is closely linked to the aminoglycosides of aminocylitol. It binds and is bacteriostatic but not bactericidal to a particular protein in the ribosome. Tetracyclines are other agents which bind to 30S ribosomes. These agents tend to inhibit aminoacyl tRNA binding to the A site of the bacterial ribosome. Tetracycline binding is temporary, so it’s bacteriostatic for these agents. Nevertheless, a wide range of bacteria, chlamydias and mycoplasmas are inhibited and highly helpful agents [29]. There are three major groups of medicines that inhibit the ribosomal subunit of 50S. A bacteriostatic agent that inhibits both gram-positive and gram-negative bacteria is chloramphenicol. By binding to a peptidyltransferase enzyme on the 50S ribosome, it prevents peptide bond formation. Macrolides are large compounds of the lactone ring that bind to 50S ribosomes and tend to impair the reaction or translocation of peptidyltransferase, or both. Erythromycin, which inhibits gram-positive species and a few gram-negative species, such as haemophilus, mycoplasma, chlamydia and legionella, is the most significant macrolide. Against many of these pathogens, new molecules including azithromycin and clarithromycin have greater activity than erythromycin. There is a similar activity site for lincinoids, the most important of which is clindamycin. Generally, macrolides and lincinoids are bacteriostatic and only inhibit the development of new peptide chains [29].

2.4 Inhibition of microbial DNA synthesis

The modulation of chromosomal supercoiling by topoisomerase-catalyzed strand breakage and rejoining reactions is needed for DNA synthesis, mRNA transcription and cell division [34]. Depending on whether they catalyze reactions involving transient breakage of one (type I) or both (type II) strands of DNA, DNA topoisomerase enzymes are classified into two groups, I and II [35]. The topological state of DNA inside cells is regulated by topoisomerases and is important for the vital processes of protein translation and cell replication. The enzyme that negatively super-coils DNA in the presence of ATP is DNA gyrase, a type II DNA topoisomerase [36]. Moreover, in the absence of ATP, this enzyme plays a role in the catenation and decatenation reaction of a double-stranded DNA circle, resolves knots in DNA, and also relaxes supercoiled DNA negatively. As a result, for almost all cellular procedures involving duplex DNA, including replication, recombination and transcription, the enzyme is vital. It is unique to the prokaryotic kingdom and is essential to the organism’s survival. Thus, for antibacterial drugs, DNA gyrase remains an ideal and attractive target. The most effective DNA gyrase-targeted antimicrobial agents are quinolones. Nalidixic acid, a naphthyridone inadvertently discovered as a by-product during chloroquine synthesis, was the source of the compounds [37].

Quinolones are unique DNA-gyrase inhibitors. DNA gyrase reactions such as supercoiling and relaxation involving DNA breakage and reunion are inhibited by quinolones, specifically interfering with the DNA gyrase breakage-reunion reaction by interacting with subunit A (GyrA) [38]. Relatively poor antimicrobial activity is found in first-generation quinolones, nalidixic acid and oxolinic acid. However, the synthesis and improvement over many generations of fluoroquinolones, such as norfloxacin and ciprofloxacin (second generation), levofloxacin (third generation), and moxifloxacin and gemifloxacin (fourth generation), has resulted in a variety of potent antimicrobial agents [38]. Most bacterial pathogens possess an additional essential topoisomerase, topoisomerase I (Topo I), in addition to the type II topoisomerases. Topo I is architecturally and mechanistically distinct from gyrase and topoisomerase IV, and is an attractive candidate for new antibacterial chemotypes to be discovered as such [36].

2.5 Inhibition of microbial RNA synthesis

Rifamycins inhibit DNA-dependent transcription by binding the DNA-bound and effectively transcribing RNA polymerase with a high affinity to the β-subunit (coded by rpoB). In the channel formed by the RNA polymerase-DNA complex, from which the newly synthesized RNA strand emerges, the β- subunit is located. Rifamycins clearly require that RNA synthesis has not progressed beyond two ribonucleotides being added; This is due to the drug molecule ‘s capacity to sterically inhibit the initialization of nascent RNA strands. It should be noted that rifamycins are not believed to work by blocking the RNA synthesis elongation stage, although a recently discovered class of RNA polymerase inhibitors (based on the CBR703 compound) could inhibit elongation by modifying the enzyme allosterically [34].

2.6 Inhibition of microbial metabolic pathways

By competitively blocking the biosynthesis of tetrahydrofolate, which acts as a carrier of one-carbon fragments and is required for the ultimate synthesis of DNA, RNA and bacterial cell wall proteins, trimethoprim and sulfonamides interfere with folic acid metabolism in the microbial cell. Bacteria and protozoan parasites typically lack a transport mechanism in order to extract preformed folic acid from their host, unlike mammals [29]. Most of these species, while some are capable of using exogenous thymidine, must synthesize folic acid, circumventing the need for metabolism of folic acid. The conversion of pteridine and p-aminobenzoic acid (PABA) to dihydrofolic acid by the pteridine synthetase enzyme is competitively inhibited by sulfonamides. Sulfonamides have a greater affinity for pteridine synthetase than for PABA. Trimethoprim has a huge affinity (10,000 to 100,000 times greater than that of the mammalian enzyme) for bacterial dihydrofolate reductase; it inhibits tetrahydrofolate synthesis when bound to this enzyme [29].

Advertisement

3. Mechanisms of resistance to antimicrobial agents

The ability of an organism to overcome the action of an antimicrobial agent to which it was previously susceptible is a general definition of antimicrobial resistance [39]. With the growing production of MDR strains (i.e. resistance to at least three antibiotics), nosocomial infection caused by antibiotic resistant P. aeruginosa has emerged as a major concern in clinical care settings [40]. Because of its outer membrane with low permeability (1/100 of the permeability of the outer membrane of E. coli), P. aeruginosa exhibits intrinsic resistance to various antimicrobial agents (β-lactam and penem group of antibiotics) [41, 42]. While several other processes are also responsible for their intrinsic resistance, including the efflux system that expels antibiotics from the cell’s bacteria and the production of inactivating enzyme antibiotics. This bacterium, however is a highly diverse pathogen capable of adapting to the conditions around it. When subjected to selective pressure from antibiotics, the mediated reaction encourages bacterial survival and improves resistance to antibiotics [43, 44, 45].

The development of antibiotic resistance during host colonization of patients with CF has been confirmed, with P. aeruginosa strains developing and gaining resistance during antimicrobial therapy [46]. Studies have shown a clear link between increased applications of ciprofloxacin, with a growing incidence of strains resistant to ciprofloxacin [47]. Therefore the excessive use of antimicrobial agents is another factor associated with the rise in MDR-Psedomonas aeruginosa. This acquired resistance may be attributable to the effects of the mutational event or the acquisition by horizontal gene transfer of the resistance gene and may occur during the mutational event of antibiotic therapy, leading to over-expression of endogenous β-lactamases or efflux pump, specific porin expression [48].

3.1 Resistance to β-lactam

Inhibition of the synthesis of the bacterial peptidoglycan cell wall requires β-lactam antibiotics [39]. Penicillin, cephalosporin, carbapenem and monobactam are included in this class. These classes include piperacillin and ticarcillin (penicillin), ceftazidime (cephalosporin 3rd generation), cefepime (cephalosporin 4th generation), aztreonam (monobactam), imipenem, meropenem and doripenem (carbapenems) are most powerful β-lactam widely used to treat P. aeruginosa is β-lactam [49]. These enzymes break the amide bond of the β-lactam ring through resistance to the β-lactam mediated by the action of β-lactamases, rendering the antimicrobial ineffective. The expression of endogenous β-lactamases or the expression of acquired β-lactamases may be due to this inactivation of the drug. To date, hundreds of β-lactamases have been recognized and are distinguished by their substrate specificity. There are four major groups of beta-lactamases known in P. aeruginosa on the basis of Amber’s molecular classification system: A-D [50]. Through the serine-residue catalytic activity, classes A, C and D inactivate the β-lactams, while class B or metallo- β-lactamases (MBLs) require zinc for their action [51].

3.2 AmpC β-lactamase (Cephelosporinase)

In particular, the development of endogenous β-lactamase, such as chromosomal cephalosporinase (AmpC β-lactamase). A variety of β-lactams, such as benzyl penicillin, narrow spectrum cephalosporin and imipenem, can be induced in P. aeruginosa. Naturally, P. aeruginosa is susceptible to carboxypenicillins, ceftazidime and aztreonam, but it can develop resistance through a mutation in the gene that contributes to AmpC β-lactamase hyper-production [52, 53]. The enzyme is usually produced in small amounts (‘low-level’ expression), resistance to aminopenicillins and to most early cephalosporins is determined. P. aeruginosa produces an inducible chromosome-coded AmpC β-lactamase (cephalosporinase) belonging to the Ambler-based molecular class C and the first functional group according to Bush et al. [54, 55].

However, production of chromosomal cephalosporin in P. aeruginosa, in the presence of inducing β-lactams (especially imipenem), can increase from 100 to 1000 times [56]. β-lactamase inhibitors used in clinical practice, such as clavulanic acid, sulbactam and tazobactam, do not inhibit AmpC cephalosporinase function. β-lactamase of AmpC is encoded by the gene ampC [57, 58]. Several genes, including ampR, ampG, and ampD, are involved in ampC gene induction. AmpR encodes a positive transcriptional regulator and this regulator is required for the induction of β-lactamase. AmpG, a transmembrane protein that functions as a permease for 1,6-anhydromurapeptides, which are known to be the signal molecules involved in the induction of ampC, is the second gene involved. The third gene, ampD, encodes a cytosolic amidase of N-acetyl-anhydromuramyl-L-alanine that hydrolyses 1,6-anhydromurapeptides, which functions as an ampC expression repressor. The 4th chromosome, ampE, encodes the protein of the cytoplasmic membrane that serves as the molecule of the sensory transducer necessary for induction. Except for avibactam, the activity of this AmpC β-lactamase is not inhibited by commercially available β-lactam.

3.3 Class A carbenicillin hydrolysing β-lactamases

Four β-lactamases (PSE- of Pseudomonas specific enzyme) carbenicillin hydrolyzing enzymes were identified in P. aeruginosa: PSE-1 (CARB-2), PSE-4 (CARB-1), CARB-3 and CARB-4 [59]. These enzymes belong to the group of β-lactamases of molecular class A and include carboxypenicillins, ureidopenicillins and cefsulodine in their substrate profile. These enzymes belong to functional group 2c and molecular class A [60]. Commercially available β-lactam inhibitors, such as clavulanic acid, tazobactam, and sulbactam, can inhibit the activity of this β-lactamase [61].

3.4 Resistance to aminoglycoside

Aminoglycosides are a microbial protein synthesis inhibitor which act by binding to the ribosomal subunit of the bacterial 30S and interfering with the initiation of protein synthesis. Resistance to aminoglycosides in Pseudomonas is mediated by transferable aminoglycoside modifying enzymes (AMEs), low permeability of the outer membrane, active efflux and, in rare cases, target modification [62, 63, 64].

3.5 Aminoglycoside-modifying enzymes

AMEs inactivate the aminoglycoside by adding the antibiotic molecule to a phosphate, adenyl or acetyl radical, and thus modified antibiotics minimize the binding affinity of the bacterial cell (30S ribosomal subunit) to its target [65, 66]. Aminoglycoside phosphoryl transferases (APHs), aminoglycoside adenylyl transferases (also known as nucleotidyltransferases) (AADs or ANTs) and aminoglycoside acetyltransferases (AACs) are three types of AMEs involved in aminoglycoside alteration. The following AMEs are most commonly expressed by P. aeruginosa: AAC(69)-II (resistant to gentamicin, tobramycin and netilmicin), AAC(3)-I (resistant to gentamicin), AAC(3)-II (resistant to gentamicin, tobramycin and netilmicin), (69)-I (resistant to tobramycin, netilmicin and amicacin) and ANT(29)-I (resistant to tobramicin and gentamicin) [67].

3.6 Low outer membrane permeability

Membrane impermeability or reduced permeability is a mechanism known to provide resistance to many antibiotic forms, including aminoglycosides, β-lactams and quinolones [68]. For instance, this resistance mechanism is often encountered in cystic fibrosis isolates that are continually under antibiotic attack. Several mechanisms, such as lipopolysaccharide (LPS) modifications, alteration of membranous proteins involved in substratum absorption, and inactivation of enzymatic complexes involved in the energetic membrane necessary for transport system activity, may cause membrane impermeability [69].

3.7 Active efflux pumps

The combination of low membrane permeability and active efflux pumps is partially due to the natural resistance of P. aeruginosa to many groups of antibiotics. P. aeruginosa’s efflux systems involved in antibiotic resistance belong to the family of resistance-nodulation-division (RND) [70]. In order to confer resistance to several antibiotics, four major efflux systems have been described: MexAB-OprM, MexCD-OprJ, MexEF-OprN and MexXY-OprM. These systems consist of three proteins: (1) the efflux pump protein found in the cytoplasmic membrane (MexB, MexD, MexF and MexY), (2) the pore-acting outer membrane protein (OprM, OprJ and OprN) and (3) A protein in the periplasmic space that bridges the cytoplasmic and outer membrane proteins (MexA, MexC, MexE and MexX). In both natural and acquired resistance, MexAB-OprM and MexXY-OprM are active, whereas only the other two mechanisms are observed in cumulative resistance.

Acquired resistance is observed following mutations in the regulatory systems that can be caused by antibiotic pressure and that can confer resistance to all groups of antibiotics upon over-expression of these efflux systems. Polymyxins, except [69]. Resistance to multiple groups of antibiotics that are substrates of these efflux systems can be caused by exposure to a single antibiotic. Quinolones are substrates of all efflux systems and are an important trigger factor that can generate cross-resistance to efflux systems of several major classes of antibiotics, including β-lactams and aminoglycosides, for pseudomonal therapy [71]. It is understood that efflux systems confer a moderate degree of resistance, but they typically act simultaneously with other mechanisms of resistance, thus taking part in the high-level resistance that can be observed in P. aeruginosa.

3.8 Target modification

Due to the low affinity of the drug to the bacterial ribosome, bacteria may be resistant to aminoglycosides. This can be achieved by 16S rRNA methylation by target modification. Various 16S rRNA methylases have been identified for P. aeruginosa: RmtA, first reported in clinical isolates of P. aeruginosa resistant to aminoglycosides and conferred resistance to all parenterally administered aminoglycosides, including amicacin, tobramycin, isepamicin, kanamycin, arbecacin and gentamicin, secondary 16S rRNA methylases including RmtB, ArmA and RmtD [72].

3.9 Resistance to fluoroquinolones

Resistance to fluoroquinolones arises by mutation in the DNA gyrase or topoisomerase 1 V coding bacterial chromosome gene or by successful drug transport out of the cell [73]. Topoisomerase 1 V mutations can occur in gyrA / gyrB genes within the motif of the quinolone-resistant determinative region (QRDR), which is considered to be the active site of the enzyme. This contributes to the altered amino acid sequences of the subunits A and B, and hence to the altered topoisomerase II with a low affinity for quinolone molecules. As a result of point mutations in parC and parE genes encoding the ParC and ParE enzyme subunits, modifications of a secondary target (topoisomerase IV) occur. The over-expression of efflux includes other types of fluoroquinolone tolerance in Pseudomonas. Mutations in the nalB, nfxB and nfxC genes, resulting in overexpression of MexA-MexB-OprM, MexC-MexD- OprJ and MexE- MexF- OprN fallowing efflux [74].

3.10 Biofilm-mediated resistance

A biofilm is an aggregate of microorganisms that bind to each other on a living or non-living surface and are embedded in an extracellular polymeric (EPS) matrix of self-produced substances, including exopolysaccharides, proteins, metabolites, and eDNA [75, 76]. The microbial cells grown in biofilms are less sensitive than the cells grown in free aqueous suspension to the antimicrobial agents and the host immune response [77]. Even bacteria that are deficient or lack protective mutations in their intrinsic resistance, when they grow in a biofilm, they can become less susceptible to antibiotics [78]. The general mechanisms of biofilm-mediated resistance that protect bacteria from antibiotic attack include antibiotic penetration prevention, altered microenvironment that induces slow biofilm cell growth, adaptive stress response induction, and differentiation of persistent cells [78, 79, 80].

P. aeruginosa causes chronic lung infections in CF patients and, through the production of DNA, proteins and exopolysaccharides, forms a biofilm on lung epithelial cell surfaces. The regulation of the formation of P. aeruginosa biofilm is multifactorial and mainly depends on quorum sensing systems, GacS / GacA and RetS / LadS two-component regulatory systems, exopolysaccharides and cdi- GMP [81]. Quorum sensing is a form of communication between bacterial cells and cells that regulates gene expression in response to changes in cell population density. P. aeruginosa has three major systems of quorum sensing, LasILasR, RhlI-RhlR, and PQS-MvfR, all of which contribute to mature and differentiated biofilm formation. During biofilm formation, P. aeruginosa undergoes numerous physiological and phenotypic changes [82]. For example, P. aeruginosa strains convert to a mucoid phenotype in CF chronic infection that displays upregulated production of alginate driven by the CF microenvironment, enabling the formation of colonies of biofilms. Due to its ability to show swarming and twitching motility, P. aeruginosa flagellum is important for the initiation of biofilm formation. However, P. aeruginosa significantly decreases flagellum expression after surface attachment and may also permanently lose the flagellum due to genetic mutations, reducing host immune response activation, allowing P. aeruginosa to evade immune detection and phagocytosis [83].

Advertisement

4. The global economic scenario of antibiotic resistance

It is still an immense global challenge to quantify the exact economic effect of resistant bacterial infections. Measuring the distribution of the disease associated with antibiotic resistance is a crucial prerequisite in this situation. A major economic burden for the entire world is antibiotic resistance. In the USA alone, 99,000 deaths are caused annually by antibiotic-resistant pathogen-associated hospital-acquired infections (HAIs). Approximately 50,000 Americans died in 2006 because of two popular HAIs, namely pneumonia and sepsis, costing the US economy around $8 billion [84]. Patients with antibiotic-resistant bacterial infections need to remain in the hospital for at least 13 days, creating an extra 8 million hospital days each year. There have been estimates of costs of up to $29,000 per patient infected for an antibiotic-resistant bacterial infection. In total, economic losses of approximately $ 20 billion were recorded in the US, while losses of approximately $35 billion per year were also recorded in terms of loss of productivity due to antibiotic resistance in health care systems [85].

A worst-case scenario could emerge in the coming future, according to the analysts of the Research and Development Corporation, a US non-profit global organization, where the planet could be left without any effective antimicrobial agent to treat bacterial infections. In this case, the global economic burden will be nearly $120 trillion ($3 trillion per annum), roughly equal to the entire actual annual health care budget of the United States. In general, the world population will be significantly affected: about 444 million people will succumb to infections as of 2050, and birth rates will decrease rapidly in this scenario [86, 87]. These losses are calamitous, but these estimates reflect imperfect images of the economic costs of antibiotic resistance due to data limitations such as the inclusion of total conditions and resistance-susceptible diseases. The use of antibiotics in the livestock and food industries is another very critical trait of antimicrobial resistance (AMR), that was missing from the investigation. It is an important player in the rising AMR, likely causing its own projected economic losses. There is also a misappropriation of the use of antimicrobials as growth promoters in many developing countries. This activity has been outlawed in the European Union since 2006 [88, 89].

Advertisement

5. Novel alternative antimicrobial therapy for P. aeruginosa treatment

The overuse and misuse of antibiotics, which can lead to unwanted side effects and the production of drug-resistant bacterial strains, is a growing public health issue. The production of new antibiotics, in addition, is very limited and timely. The development of innovative therapeutic approaches to the treatment of infections with P. aeruginosa is therefore highly desirable and has received further interest over the past decade. These innovative therapeutic techniques, which involve antimicrobial peptides, phage therapy, inhibition of quorum sensing as well as the use of iron chelation, nanoparticles, probiotics and vaccine strategies.

5.1 Antimicrobial peptides

A number of species, from bacteria to animals, develop antimicrobial peptides (AMPs), also called host defense peptides, and they are active against a wide range of microorganisms [90]. There is no complete understanding of the mode(s) of operation of AMPs. It is widely agreed that the cytoplasmic membrane is attacked by AMPs, leading to cell death [91]. AMPs have also been shown to possess anti-biofilm and immunomodulatory properties, in addition to antimicrobial activity, AMPs have been proposed as an alternative to traditional antibiotics to battle bacterial infections as a result of their broad-spectrum activity; AMPs exhibit rapid killing kinetics, low mediated resistance levels, and low host toxicity [92]. Many antimicrobial peptides, including GL13 K, LL-37, T9W, NLF20, cecropin P1, indolicidin, magainin II, nisin, ranalexin, melittin, and defensin, have demonstrated powerful antimicrobial effects of either direct bactericidal effects or biofilm disruption against P. aeruginosa [93]. In addition, by facilitating antibiotic absorption, disrupting biofilm formation or inhibiting bacterial quorum, some AMPs have demonstrated synergy with traditional antibiotics against several bacteria, including P. aeruginosa [94]. For instance, it has been shown that the clearance of P. aeruginosa biofilm was increased by a combination of GL13 K with tobramycin [95]. In 2017, Zheng et al. [96] observed that when combined with tetracycline in vitro, the minimum inhibitory concentration of cecropin A2 against clinical isolates of P. aeruginosa was reduced 8-fold.

5.2 Phage therapy

By inducing lysis, bacteriophages (phages) are viruses that infect and destroy bacteria [97]. In 1915, the British bacteriologist Frederick Twort first discovered phages. Two years later, Félix d’Herelle made a similar discovery independently in Paris and presented the phage therapy notion. With the advent of antibiotic therapy, phage therapy was abandoned in several countries, but has been continuously developed in Eastern European countries with facilities in Warsaw, Poland, and Tbilisi, Georgia [98]. Shotgun metagenome sequencing showed that there were antipseudomonal phages in the phage cocktails sold in pharmacies in Georgia and Russia [99]. The successful treatment of infections with MDR P. aeruginosa has been reported in a few case reports from Belgium and the US, but has not gained broad acceptance in the Western world [100]. Phage therapy has many benefits, including replication at the infection site, high precision for attacking bacteria without effects on commensal flora, less side effects than other therapies, antibiotic-resistant bacteria bactericidal activity and simple administration [101]. The use of phages as an alternative to antibiotics has been extensively studied for the treatment of P. aeruginosa infections. There are 137 different phages that have been characterized to date that target the Pseudomonas genus [102]. Many in vitro and in vivo studies have been performed to test the efficacy of phages against chronic infections of P. aeruginosa. For instance, co-incubation of phage PA709 with the clinical strain P. aeruginosa 709 has been shown to significantly reduce the viability of P. aeruginosa. Another research found that intranasal administration of P3-CHA bacteriophage to mice receiving a lethal dose of P. aeruginosa strain CHA substantially improved the rate of survival and reduced the bacterial load in the lungs [103].

Another benefit of phage therapy is that phages can be genetically modified as vehicles to transport bacteria with antimicrobial agents, thus increasing treatment efficacy [104]. While phages have been shown to be successful in vitro and in animal models against bacterial infection, only a small number of phage therapy clinical trials have been performed to date. The reasons for this include: safety issues about post-treatment phage clearance and impurity of phage preparations; poor stability of phage preparations; and lack of knowledge of the comprehensive phage mode of action and bacterial resistance to phage growth [105]. In clinical trials, the use of phages against P. aeruginosa infections has been studied in patients with venous leg ulcers, burn wounds and otitis, and no adverse reactions have been identified during these clinical trials [106].

5.3 Quorum sensing inhibition

Quorum sensing is a mechanism that enables bacteria to regulate the expression of genes in a manner based on cell density. To control virulence and biofilm formation, P. aeruginosa utilizes quorum sensing [107]. Las and Rhl are two major P. aeruginosa quorum-sensing systems responsible for the synthesis of the signal molecules of N-acyl homoserine lactone (AHL), N-(3-oxododecanoyl)-L-homoserine lactone (3O-C12-HSL) and N-butanoyl-L-homoserine lactone (C4-HSL). 3O-C12-HSL and C4-HSL bind to and activate their LasR and RhlR cognate transcription factors, respectively, inducing the formation of biofilms and the expression of various virulence factors, including elastase, proteases, pyocyanin, lectins, rhamnolipids, and toxins [108]. The third P. aeruginosa quorum-sensing system, PQSMvfR, has been reported to facilitate the formation of biofilms in addition to the LasI-LasR and RhlI-RhlR systems. This mechanism regulates the development of the Pseudomonas quinolone signal (PQS), 2-heptyl-3-hydroxy-4-quinolone, by the transcriptional regulator MvfR, also known as PqsR, by controlling the pqsABCDE operon. In addition, PqsA and PqsD proteins have been implicated in the development of biofilms [82].

A promising technique for treating P. aeruginosa infections is known to be the inhibition of quorum sensing. This approach is capable of preventing or decreasing the formation of biofilms, reducing bacterial virulence and has a low risk of bacterial resistance growth. In addition, this strategy has a small scope, such that any unwanted inhibitory effects on beneficial bacteria are impossible. For the Las and Rhl systems, quorum sensing inhibitors may be either natural or synthetic and are capable of reducing the activity of AHL synthase, inhibiting the development of AHL, degrading AHLs or competing for AHL receptor binding [109]. In recent years, the use of quorum sensing inhibitors for the treatment of infections with P. aeruginosa has been intensively studied. The carotenoid zeaxanthin, typically found in plants, algae and lichens, for example, reduced the formation of biofilms in P. aeruginosa by binding to the signal receptors for quorum sensing, lasR and RhlR, and blocking the expression of virulence genes, lasB and rhlA [110]. Flavonoids are a class of naturally developed plant metabolites that have acted as LasR and RhlR antagonists and substantially decreased their ability to bind to the P. aeruginosa promoters of quorum sensing-regulated genes [111].

5.4 Iron chelation

Iron is important for bacterial growth and is involved in a number of cellular processes, such as the production of electricity, the replication of DNA and the transport of electrons [112]. Compared to healthy people, the iron content of human sputum was found to be substantially elevated in CF patients, indicating that an increased amount of iron promotes chronic CF lung infection [113]. P. aeruginosa utilizes pyoverdine and pyochelin siderophores to obtain iron from the extracellular environment [114]. Therefore, a technique to fight P. aeruginosa infections is to limit the concentration of extracellular iron or disrupt iron uptake by P. aeruginosa. Several studies have related iron metabolism to the pathogenesis of chronic infections, indicating that iron analogues and chelators may work against P. aeruginosa as potential therapeutic agents. For example, iron chelators, 2,2′- dipyridyl (2DP), diethylenetriaminepentacetic acid (DTPA) and EDTA, have been reported to impair growth and biofilm formation of P. aeruginosa and have been more effective under anaerobic conditions [115].

Gallium is a nonredox iron III analog that disrupts the metabolism of bacterial iron by acting in several biological processes as an iron replacement, so it is a US FDA-approved medication for cancer-associated hypercalcemia treatment [116]. In 2007, Kaneko et al., reported that gallium was able to inhibit the growth of P. aeruginosa, prevent the development of biofilms, and manifest excellent bactericidal activity in vitro by reducing the uptake of bacterial iron and repressing the synthesis of pyoverdine mediated by the transcriptional regulator PvdS. In addition, in mouse infection models, gallium has also been found to remove P. aeruginosa effectively.

5.5 Nanoparticles

Currently, a variety of diseases, including cancer and bacterial infectious diseases, have received significant attention from nanoparticles to treat them. Nanoparticles are small materials that have been used in a number of chemical, biological and biomedical applications, having a size of less than 100 nm and a large surface area to mass ratio [117]. The nanoparticles used for their antimicrobial activity are highly penetrable in the bacterial membranes, may interfere with the formation of biofilms, have several antimicrobial mechanisms, and are strong antibiotic carriers [118]. For the prevention of P. aeruginosa infections, metallic and antimicrobial agent-loaded nanoparticles have been extensively examined. Silver nanoparticles, for example, are powerful antimicrobial agents that generate silver ions responsible for the inhibition, like DNA synthesis, of bacterial enzymatic systems. Silver nanoparticles have shown important antimicrobial effects on the clinical strains of P. aeruginosa, killing P. aeruginosa effectively and inhibiting its in vitro growth. In addition, silver nanoparticles have demonstrated low mammalian cell cytotoxicity, although this requires more in vivo research [119].

Nanoparticles are capable of delivering antimicrobial agents such as antibiotics to bacteria, as described earlier. Kwon et al., developed porous silicon nanoparticles with a novel antimicrobial peptide fused with a synthetic bacterial toxin, containing membrane-interacting peptides. This engineered nanoparticle was discovered in a mouse model of P. aeruginosa lung infection to increase the survival rate and bacterial clearance. Moreover, it has been found that the binding of antibiotics to nanoparticle surfaces greatly improves the effectiveness of both antibiotics and nanoparticles. In this respect, silver ampicillin-attached nanoparticles have a higher rate of in vitro killing of ampicillin-resistant P. aeruginosa isolates compared to silver ampicillin-attached nanoparticles [120].

5.6 Probiotic as an alternative antimicrobial therapy

Probiotics are living microorganisms which, when ingested in appropriate quantities, provide health benefits [121]. The majority of probiotic bacteria are gram-positive, and their primary functions are related to intestinal tract health regulation and maintenance (e.g., Lactobacillus and Bifidobacterium) [122]. The probiotics in the intestines that colonize the human host are the most numerous. The commensal intestinal microbiome leads to enhanced infection tolerance, differentiation of the host immune system, and nutrient synthesis [123]. The probiotic Pediococcus acidilactici HW01 was studied against P. aeruginosa and observed decreased P. aeruginosa motility as well as decreased pyocyanin development, decreased protease and rhamnolipid production, and decreased stainless steel surface biofilm formation. Another research conducted by Moraes et al., showed that Lactobacillus brevis and Bifidobacterium bifidum were effective against S. aureus biofilms grown on titanium discs. The findings showed a decrease in S. aureus growth on titanium discs when both probiotics were used, but L. brevis strains was shown to have the greatest inhibitory effect on biofilm formation.

Recent studies by Xu et al. [124], have indicated that probiotics can be used by patients infected by COVID 19 to prevent secondary infections. There was intestinal microbial dysbiosis in some patients with COVID-19. In all patients, nutritional and gastrointestinal functions must be measured. To control the composition of the intestinal microbiota and reduce the risk of secondary infection due to bacterial translocation, nutritional support and application of probiotics was suggested.

5.7 Vaccine strategy

The concept of a vaccine strategy is to avoid infection until it can be produced. The production of vaccines aims to prevent and decrease infections of P. aeruginosa [125]. However, no approved vaccine against this pathogen is yet available. P. aeruginosa antigens, which are responsible for pathogenesis, induce potent immune responses. LPS O-antigen, polysaccharide protein conjugates, outer membrane proteins OprF and OprI, type III secretion system portion PcrV, flagella, pili, DNA, live-attenuated P. aeruginosa and whole killed cells are possible candidates for P. aeruginosa vaccines [126]. Among the potential P. aeruginosa vaccines, phase III clinical trials in CF patients were performed only with the flagella vaccine and the recombinant vaccine IC43, containing OprF and OprI.

Related to the ability of this pathogen to undergo phenotypic changes in variable environmental conditions, the existing vaccines for P. aeruginosa demonstrate poor efficacy in clinical trials. For example, P. aeruginosa strains downregulate the expression of highly immunogenic virulence factors in CF patients’ lungs, such as LPS O-antigen, type III secretion systems, flagella and pili [127]. In addition, impaired mechanisms of host protection often reduce vaccination effectiveness. Due to the CF lungs having an altered mucus layer, impaired phagocytosis, and dysregulated inflammatory responses, including aberrant cytokine and chemokine production, and reduced phagocyte recruitment, the lung microenvironment in CF patients has become a great challenge for effective vaccination [128].

Advertisement

6. Role of combination therapy versus monotherapy

Early administration of adequate antibiotic therapy was associated with a favorable clinical outcome, especially among critically ill patients with serious P. aeruginosa infections [129]; on the other hand, delays in administering adequate antibiotic therapy were associated with a substantial increase in mortality. The progressive rise in antibiotic resistance in P. aeruginosa has been established in recent years as the key explanation for the inadequacy of antibiotics, with a negative effect on patient survival [130].

The evidence available indicates that the greatest advantage of combination therapy derives from an increased probability of selecting an appropriate agent during empirical therapy, rather than avoiding resistance during definitive therapy or benefiting from synergistic action in vitro. Therefore, researchers recommend early administration of a combination regimen when P. aeruginosa is suspected, followed by a prompt de-escalation when the antimicrobial susceptibility test becomes available, to balance between early antibiotic administration and the risk of resistance selection. An approach consisting of the prescription of an anti-pseudomonal beta-lactam (piperacillin / tazobactam, ceftolozane / tazobactam, ceftazidime, cefepime, or carbapenem) plus a second (aminoglycoside or fluoroquinolone) anti-pseudomonal agent is encouraged.

Advertisement

7. New antipseudomonal antibiotics

Related to the emergence of multidrug-resistant strains, traditional antibiotic therapies against P. aeruginosa infections have become increasingly ineffective. The use of various antibiotic combinations and the development of new antibiotics are existing therapeutic options for P. aeruginosa treatment. New antibiotics have been shown to be more effective in destroying P. aeruginosa and have a lower frequency of production of resistance compared to current antibiotics due to their novel modes of action, efficient delivery of drugs (e.g. inhaled antibiotics) and resistance to bacterial enzyme alteration. Novel antibiotics with action against P. aeruginosa have been available in Europe in recent years and others are in advanced stages of clinical development. In certain instances, indirect evidence indicates their possible superiority over standard anti-pseudomonal regimes.

7.1 Doripenem

Doripenem is a new carbapenem antibiotic with wide spectrum activity against gram-negative and gram-positive bacteria by binding to penicillin-binding proteins by inhibiting bacterial cell wall synthesis; it has been approved for the treatment of complicated intra-abdominal infection and urinary tract infection by the US Food and Drug Administration (FDA) [131]. Except for the metallo-β-lactamases of class B, doripenem is resistant to hydrolysis by several β-lactamases. Importantly, compared to other carbapenem antibiotics such as meropenem and imipenem, the in vitro antibacterial activity of doripenem against the P. aeruginosa isolates from CF patients was found to be more active [132]. In addition, the effectiveness of doripenem was tested in patients with P. aeruginosa ventilator-associated pneumonia, a phase III clinical trial of patients with P. aeruginosa ventilator-associated pneumonia found that patients treated with doripenem had higher rates of cure compared to patients treated with imipenem. Of note, headache, nausea, diarrhea, rash, and phlebitis are among the side effects of doripenem [133].

7.2 Plazomicin

Plazomicin is a semisynthetic aminoglycoside antibiotic of the next generation that is synthetically derived from the natural product sisomicin. A wide range of aminoglycoside modifying enzymes, but not 16S rRNA ribosomal methyltransferases, are able to resist plazomicin [134]. Plazomicin exhibits potent in vitro activity against both gram-negative and gram-positive bacterial pathogens and has an activity close to that of amikacin against strains of multidrug-resistant P. aeruginosa. In addition, Pankuch et al., reported in vitro synergistic activity of plazomicin against clinical isolates of P. aeruginosa in combination with cefepime, doripenem, imipenem or piperacillin-tazobactam and no antagonism was observed in this study, indicating that plazonmicin is a possible candidate for combination therapy in the treatment of infections with multidrug-resistant P. aeruginosa. Plazomicin can cause nephrotoxic and ototoxic effects that are mild to moderate [135].

7.3 POL7001

As a novel class of antibiotics against P. aeruginosa, protein epitope mimetic (PEM) molecules have emerged; some PEM molecules inhibit the transfer of LPS to the outer bacterial membrane [136]. A macrocycle molecule belonging to the PEM antibiotic family is POL7001. The efficacy of POL7001 was tested by Cigana et al., both in vitro and in murine P. aeruginosa acute and chronic pneumonia models. They observed that P. aeruginosa multidrug-resistant isolates were susceptible to POL7001 in CF patients, and that POL7001-treated mice had substantially decreased bacterial burden and decreased levels of lung inflammation during acute and chronic P. aeruginosa infection. POL7001 as a novel therapeutic agent for potential clinical trials is indicated by the new mode of action, effective pulmonary delivery and potent in vitro and in vivo activity. The side effects of POL7001 have not yet been identified [137].

7.4 Arikayce ™

Arikayce™ has been approved by the FDA for the treatment of Mycobacterium avium complex (MAC) lung disease, and is a liposomal amikacin treatment. Clinical trials for this drug and its efficacy in the treatment of P. aeruginosa in patients with cystic fibrosis have been performed. While these are early phases and some experiments would have to resolve the drawbacks of this compound in order to improve safety, some experimental clinical trials have been performed [138].

7.5 Ceftolozane-tazobactam

To resolve P. aeruginosa antimicrobial resistance mechanisms, such as changes in porine permeability and upregulation of efflux pumps, ceftolozane-tazobactam is being created. Due to a higher affinity for all essential PBPs, including PBP1b, PBP1c and PBP3, this drug has an intrinsically potent anti-pseudomonal effect [139]. Ceftolozane/ Tazobactam has been shown to have a strong in vitro activity against most strains of MDR P. aeruginosa [including strains developing extended- spectrum β-lactamase (ESBL) but not carbapenemase]. The therapeutic use of ceftolozane-tazobactam in complicated intra-abdominal and urinary tract infections has been suggested by the FDA [140]. In addition, a study is currently underway for the treatment of HAP, including ventilator-associated pneumonia (VAP). In 71 percent of patients with MDR P. aeruginosa infections, evidence from real-world trials using ceftolozane-tazobactam for the treatment of MDR P. aeruginosa infections showed promising results.

7.6 Ceftazidime-avibactam

Ceftazidime-avibactam is a novel combination of β-lactam / BLI approved for the treatment of complicated urinary tract infections (UTIs) and complicated intra-abdominal infections by the Food and Drug Administration (FDA) and European Medicines Agency (EMA). In vitro studies have shown that the combination of ceftazidime-avibactam is highly effective against KPC- producing Klebsiella pneumoniae carbapenemase (KPCs), oxacillinase (OXA), extended- spectrum β-lactamases (ESBLs) and AmpC enzymes producing Enterobacteriaceae. The drug does not, however, have any action against metallo-beta β- lactamases (MBL, VIM and NDM) and avibactam does not have any improved activity against P. aeruginosa [141]. In phase III research compared ceftazidime-avibactam to meropenem (NTC01808092), the efficacy of ceftazidime-avibactam against VAP was analyzed [142]. The predominant isolated baseline gram-negative pathogens were K. pneumoniae and P. aeruginosa, with 28% of patients possessing a non-susceptible isolate of 1% ceftazidime. 356 patients in the clinically evaluable population were treated with ceftazidime-avibactam and 370 with meropenem. The research met the non-inferiority criterion for ceftazidime-avibactam as there was no disparity in the outcome between the groups. In addition, the efficacy of ceftazidime-avibactam was close to that of ceftazidime-susceptible pathogens against ceftazidime-non-susceptible strains and was also comparable to meropenem.

7.7 Imipenem-cilastatin-relebactam

Relebactam is a β-lactamase inhibitor (BLI) diazabicyclooctane that inhibits β-lactamase class A and C activity, but has no activity against metallo-β-lactamase. It has been shown that the combination of imipenem-cilastatin with relebactam has synergistic activity against a broad range of MDR gram negative pathogens including P. aeruginosa, KPC-producing K. pneumoniae and Enterobacter spp. [143]. This medication has been tested predominantly in patients with IAI, complicated UTI, and pyelonephritis, although a trial is currently underway in patients with HAP/ VAP. Some new medications have a small effect on P. aeruginosa, such as plazomycin, meropenem-vaborbactam and aztreonam-avibactam [144].

Advertisement

8. Conclusions

Treatment of infections with P. aeruginosa continues to be significant challenging. Improving the early diagnosis and empirical treatment of serious P. aeruginosa infections is an urgent need. First, to quickly announce the detection and susceptibility results for Pseudomonas in blood cultures and other clinically important cultures, matrix-assisted-laser-desorption-ionization time-of-flight mass spectrometry (MALDI-TOF) and modern molecular techniques should be routinely introduced. However, in order to decide if such diagnostic methods have a real effect on hospitalization time and patient mortality, controlled trials would be required. Secondly, more studies are urgently required to classify patients at risk of infection with MDR P. aeruginosa (bloodstream infections, urinary tract infections) based on clinical risk factors. Ultimately, clinical response depends on factors such as underlying diseases, seriousness of infection, form of infection, adequate control of the source, and response to prior antibiotics. There is an immediate need to determine the true impact of the latest anti-Pseudomonas drugs recently approved for the treatment of these infections on patient outcomes. To date, however, due to high cost, side effects and safety issues, few of these newer methods have continued to clinical practice.

References

  1. 1. Olga P, Lucia B, Inès B, Laura FG, Mónica GB, Antón A, María L, German B, Maria T. 2020. Strategies to Combat Multidrug-Resistant and Persistent Infectious Diseases. Antibiotics.2020; 9, 65. DOI: 10.3390/antibiotics9020065
  2. 2. WHO. Worldwide Country Situation Analysis: Response to Antimicrobial Resistance; WHO Library Cataloguing-in-Publication Data;World Health Organization: Geneva, Switzerland, 2015. ISBN: 978 92 4156494 6
  3. 3. Cerceo E, Deitelzweig SB, Sherman BM, Amin AN. Multidrug-Resistant Gram-Negative Bacterial Infections in the Hospital Setting: Overview, Implications for Clinical Practice, and Emerging Treatment Options. Microbial Drug Resistance. 2016;22, 412-431. DOI: 10.1089/mdr.2015.0220
  4. 4. El Solh AA, Alhajhusain A. Update on the treatment of Pseudomonas aeruginosa pneumonia. Antimicrobial Agents and Chemotherapy. 2009; 64, 229-238. DOI: 10.1093/jac/dkp201
  5. 5. Park SY, Park HJ, Moon SM, Park KH, Chong YP. Impact of adequate empirical combination therapy on mortality from bacteremic Pseudomonas aeruginosa pneumonia. BMC Infectious Diseases. 2012; 12, 308. DOI: 10.1186/1471-2334-12-308
  6. 6. Lister PD, Wolter DJ, Hanson ND. Antibacterial-resistant Pseudomonas aeruginosa: clinical impact and complex regulation of chromosomally encoded resistance mechanisms. Clinical Microbiology Reviews. 2009; 22, 582-610.DOI: 10.1128/CMR.00040-09
  7. 7. Tacconelli, E., Magrini, N., Carmeli, Y., Harbarth, S., Kahlmeter, G., Kluytmans, J., Mendelson, M., Pulcini, C., Singh, N., Theuretzbacher, U. Global priority list of antibiotic-resistant bacteria to guide research, discovery, and development of new antibiotics. World Health Organization 1-7
  8. 8. Hirsch EB, Tam VH. Impact of multidrug-resistant P. aeruginosa infection on patient outcomes. Expert Review of Pharmacoeconomics and Outcomes Research. 2010;10, 441-451. DOI: 10.1586/erp.10.49
  9. 9. Hancock RE, Speert DP. Antibiotic resistance in Pseudomonas aeruginosa: mechanisms and impact on treatment. Drug Resistance Updates. 2000; 3, 247-255. DOI: 10.1054/drup.2000.0152
  10. 10. Breidenstein EB, Fuente-Nunez C, Hancock RE. Pseudomonas aeruginosa: all roads lead to resistance. Trends in Microbiology. 2011; 19, 419-426. DOI.10.1016/j.tim.2011.04.005
  11. 11. Drenkard E. Antimicrobial resistance of Pseudomonas aeruginosa biofilms. Microbes and Infection. 2003; 5, 1213-1219. DOI: 10.1016/j.micinf.2003.08.009
  12. 12. Burkhard TU. Emerging therapies against infections with Pseudomonas aeruginosa. 2019; 8(F1000 Faculty Rev):1371 . DOI: 10.12688/f1000research.19509.1
  13. 13. Gorr SU, Flory CM, Schumacher RJ. In vivo activity and low toxicity of the second-generation antimicrobial peptide DGL13K. PLoS One. 2019; 14(5). DOI: 10.1371/journal.pone.0216669
  14. 14. Yasir M, Dutta D, Willcox MDP. Comparative mode of action of the antimicrobial peptide melimine and its derivative Mel4 against Pseudomonas aeruginosa. Scientific Reports. 2019; 9(1): 7063. DOI: 10.1038/s41598-019-42440-2
  15. 15. Romoli O, Mukherjee S, Mohid SA. Enhanced silkworm cecropin B antimicrobial activity against Pseudomonas aeruginosa from single amino acid variation. ACS Infectious Diseases. 2019; 5(7): 1200-13. DOI: 10.1021/asinfecdis. 9b00042
  16. 16. Molchanova N, Wang H, Hansen PR. Antimicrobial activity of α- peptide/β-peptoid lysine-based peptidomimetics against colistin-resistant Pseudomonas aeruginosa isolated from Cystic Fibrosis patients. Frontiers of Microbiology. 2019; 10: 275. DOI: 10.3389/fmicb.2019.00275
  17. 17. Candido ES, Cardoso MH, Chan LY. Short cationic peptide derived from archaea with dual antibacterial properties and anti-Infective potential. ACS infectious diseases. 2019; 5(7): 1081-1086. DOI: 10.1021/asinfecdis. 9b00073
  18. 18. Kang HK, Seo CH, Luchian T. Pse-T2, an antimicrobial peptide with high-level, broad-spectrum antimicrobial potency and skin biocompatibility against multidrug-resistant P. aeruginosa infection. Antimicrobial Agents and Chemotherapy. 2018; 62(12). DOI: 10. 1128/ AAC. 01493-18
  19. 19. Beaudoin T, Stone TA, Glibowicka M. Activity of a novel antimicrobial peptide against P. aeruginosa biofilms. Scientific Reports. 2018; 8(1): 14728. DOI: 10.1038/s41598-018-33016-7
  20. 20. Akbari R, Hakemi Vala M, Hashemi A. Action mechanism of melittinderived antimicrobial peptides, MDP1 and MDP2, de novo designed against multidrug resistant bacteria. Amino Acids. 2018; 50(9): 1231-43. DOI: 10.1007/s00726-018-2596-5
  21. 21. Mulcahy LR, Burns JL, Lory S, Lewis K. Emergence of Pseudomonas aeruginosa strains producing high levels of persister cells in patients with cystic fibrosis. Journal of Bacteriology. 2010 D;192(23):6191-9. DOI: 10.1128/JB.01651-09
  22. 22. Walkty A, Adam H, Baxter M, Denisuik A, Lagace WP. In vitro activity of plazomicin against 5,015 gram-negative and gram-positive clinical isolates obtained from patients in canadian hospitals as part of the CANWARD study, 2011-2012. Antimicrobial Agents and Chemotherapy. 2014; 58(5): 2554-63. DOI: 10.1128/AAC.02744-13
  23. 23. Cigana C, Bernardini F, Facchini M, Alcala-Franco B, Riva C. Efficacy of the Novel Antibiotic POL7001 in Preclinical Models of Pseudomonas aeruginosa Pneumonia. Antimicrobial Agents and Chemotherapy. 2016; 60, 4991-5000. DOI: 10.1128/AAC.00390-16
  24. 24. Chatterjee M, Anju CP, Biswas L, Anil Kumar V, Gopi Mohan C, Biswas R. Antibiotic resistance in Pseudomonas aeruginosa and alternative therapeutic options. International Journal of Medical Microbiology. 2016;306(1):48-58. DOI: 10.1016/j.ijmm.2015.11.004
  25. 25. Guilhelmelli F, Vilela N, Albuquerque P, Derengowski LS, Silva-Pereira S, Kyaw CM. Antibiotic development challenges: the various mechanisms of action of antimicrobial peptides and of bacterial resistance. Frontiers of Microbiology. 2013;4:353. DOI: 10.3389/fmicb.2013.00353
  26. 26. Hooper DC. Mechanisms of Action of Antimicrobials: Focus on Fluoroquinolones. Clinical Infectious Diseases. 2001;32 (Suppl 1):S9-S15. DOI: 10.1086/319370
  27. 27. Kotra LP, Mobashery S. Beta-lactam antibiotics, Beta-lactamases and bacterial resistance. Bulletin of Institute Pasteur. 1998; 96: 139-150. DOI: 10.1016/S0020-2452(98)80009-2
  28. 28. Wilke MS, Lovering AL, Strynadka NCJ. Beta-Lactam antibiotic resistance: a current structural perspective. Current Opinion in Microbiology. 2005;8:525-533. DOI: 10.1016/j.mib.2005.08.016
  29. 29. Neu HC, Gootz TD. Baron S. Medical Microbiology. 4th edition. Galveston (TX): University of Texas Medical Branch at Galveston; 1996. ISBN-10: 0-9631172-1-1
  30. 30. Peterson JW, Baron S. Bacterial Pathogenesis. Medical Microbiology. 4th edition. Galveston (TX): University of Texas Medical Branch at Galveston; 1996. DOI: 10: 0-9631172-1-1
  31. 31. Krupovi_CM, Daugelavi_cius R, Bamford DH. Polymyxin B Induces Lysis of Marine Pseudoalteromonads. Antimicrobial Agents and Chemotherapy. 2007; 51(11):3908-3914. DOI: 10.1128/aac.00449-07
  32. 32. McKee EE, Ferguson M, Bentley AT, Marks TA. Inhibition of mammalian mitochondrial protein synthesis by oxazolidinones. Antimicrobial Agents and Chemotherapy. 2006;50(6):2042-9. DOI: 10.1128/AAC.01411-05
  33. 33. Cocito C, Di Giambattista M, Nyssen E, Vannuffel P. Inhibition of protein synthesis by streptogramins and related antibiotics. Journal of Antimicrobial Chemotherapy. 1997; 39:7-13. DOI:10.1093/JAC/39.SUPPL-1.7
  34. 34. Kohanski MA, Dwyer DJ, Collins JJ. How antibiotics kill bacteria: from targets to networks. Nature Reviews Microbiology, 2010; 8:423-435. DOI: 10.1038/nrmicro2333
  35. 35. Liu LF, Liu CC, Alberts BM. Type II DNA topoisomerases: enzymes that can unknot a topologically knotted DNA molecule via a reversible double-strand break. Cell. 1980;19(3):697-707. DOI: 10.1016/s0092-8674(80)80046-8
  36. 36. Ehmann DE, Lahiri SD. Novel compounds targeting bacterial DNA topoisomerase/DNA gyrase. Current Opinion in Pharmacology. 2014 ;18:76-83. DOI: 10.1016/j. coph. 2014.09.007
  37. 37. Chatterji, Unniraman MS, Mahadevan S, Nagaraja V. Effect of different classes of inhibitors on DNA gyrase from Mycobacterium smegmatis. Journal of Antimicrobial Chemotherapy. 2001;48(4):479-485. DOI: 10.1093/jac/48.4.479
  38. 38. Collin F, Karkare S, Maxwell A. Exploiting bacterial DNA gyrase as a drug target: current state and perspectives. Applied Microbiology and Biotechnology. 2011; 92(3):479-497. DOI: 10. 1007/s00253-011-3557-z
  39. 39. Bagge N, Hentzer M, Andersen JB, Ciofu O, Givskov M, Hoiby N. Dynamics and spatial distribution of Beta-lactamase expression in Pseudomonas aeruginosa biofilms. Antimicrobial Agents and Chemotherapy. 2004; 48(4): 1168-1174. DOI: 10.1128/aac.48.4. 1168-1174.2004
  40. 40. Juan C, Macia MD, Gutierrez O, Vidal C, Perez JL, Oliver A. Molecular mechanisms of beta-lactam resistance mediated by AmpC hyper production in P. aeruginosa clinical strains. Antimicrobial Agents and Chemotherapy. 2005;49(11):4733-4738. DOI: 10.1128/AAC.49.11.4733-4738.2005
  41. 41. Bush K, Jacoby GA, Medeiros AA. A functional classification scheme for beta-lactamases and its correlation with molecular structure. Antimicrobial Agents and Chemotherapy. 1995;39(6):1211-33. DOI: 10.1128/aac.39.6.1211
  42. 42. Martin-Loeches I, Deja M, Koulenti D. Potentially resistant microorganisms in intubated patients with hospital acquired pneumonia: the interaction of ecology, shock and risk factors. Intensive Care Medicine. 2013;39(4):672-681. DOI: 10.1007/s00134-012-2808-5
  43. 43. Okomoto K, Gotoh N, Nishino T. Pseudomonas aeruginosa reveals high intrinsic resistance to penem antibiotics: penem resistance mechanisms and their interplay. Antimicrobial Agents and Chemotherapy. 2001; 45(7): 1964-1971. DOI: 10. 1128/AAC.45.7.1964-1971
  44. 44. Paul M, Benuri-Silbiger I, Soares-Weiser K, Leibovici L. Beta lactam monotherapy versus beta lactam- aminoglycoside combination therapy for sepsis in immune competent patients: systematic review and meta-analysis of randomized trials. British medical journal. 2004; 328 (7441): 668. DOI: 10. 1136/ bmj. 38028.520995.63
  45. 45. Poole K. Aminoglycoside resistance in Pseudomonas aeruginosa. Antimicrobial Agents and Chemotherapy. 2005;49(2):479-487. DOI: 10.1128/AAC.49.2.479-487
  46. 46. Rello J, Allegri C, Rodriguez A. Risk factors for ventilator-associated pneumonia by Pseudomonas aeruginosa in presence of recent antibiotic exposure. Anesthesiology. 2006; 105(4):709-714.DOI: 10.1097/00000542-200610000-00016
  47. 47. Horcajada JP, Shaw E, Padilla B. Healthcare-associated, community-acquired and hospital-acquired bacteraemic urinary tract infections in hospitalized patients: a prospective multicentre cohort study in the era of antimicrobial resistance. Clinical microbiology and infection. 2013;19(10):962-8. DOI: 10.1111/1469-0691.12089
  48. 48. Honore N, Nicolas MH, Cole ST. Regulation of enterobacterial cephalosporinase production: the role of a membrane-bound sensory transducer. Molecular Microbiology. 1989; 3(8):1121-1130. DOI: 10.1111/j.1365-2958.1989.tb00262.x
  49. 49. Kock R, Becker K, Cookson B. Methicillin-resistant S. aureus (MRSA): burden of disease and control challenges in Europe. Eurosurveillance. 2010;15(41):19688. DOI: 10.2807/ese.15.41.19688-en
  50. 50. Vakulenko SB, Mobashery S. Versatility of aminoglycosides and prospects for their future. Clinical Microbiology Reviews. 2003; 16(3): 430-450. DOI: 10.1128/CMR.16.3.430-450.2003
  51. 51. Ramirez MS, Tolmasky ME. Aminoglycoside modifying enzymes. Drug Resistace Updates. 2010; 13(6): 151-171. DOI: 10.1016/j.drup.2010.08.003
  52. 52. Poole K. 2011. Pseudomonas aeruginosa: resistance to the max. Frontiers of Microbiology. 2011; 2: 65. DOI: 10.3389/fmicb.2011.00065
  53. 53. Liano-Sotelo B, Azucena EF, Kotra LP, Mobashery S, Chow CS. Aminoglycosides modified by resistance enzymes display diminished binding to the bacterial ribosomal aminoacyl- tRNA site. Chemistry&Biology.2002;9(4):455-463 DOI: 10. 1016/s1074-5521 (02) 00125-4
  54. 54. Miller GH, Sabatelli FJ, Hare RS. The Aminoglycoside Resistance Study Group. The most frequent aminoglycoside resistance mechanisms e changes with time and geographic area: a reflection of aminoglycoside usage patterns. Clinical Infectious Diseases. 1997; 24 Suppl 1:S46-62. DOI: 10.1093/clinids/24.supplement-1.s46
  55. 55. MacLeod DL, Nelson LE, Shawar RM. Aminoglycoside resistance mechanisms for cystic fibrosis Pseudomonas aeruginosa isolates are unchanged by long-term, intermittent, inhaled tobramycin treatment. The Journal of Infectious Diseases. 2000;181(3):1180-1184. DOI: 10.1086/315312
  56. 56. Yamane K, Doi Y, Yokoyama K. Genetic environments of the rmtA gene in Pseudomonas aeruginosa clinical isolates. Antimicrobial Agents and Chemotherapy 2004; 48(6): 2069-2074. DOI: 10.1128/AAC.48.6.2069-2074.2004
  57. 57. Gast Gurung M, Moon DC, Tamang MD. Emergence of 16S rRNA methylase gene armA and cocarriage of blaIMP-1 in Pseudomonas aeruginosa isolates from South Korea. Diagnostic Microbiology and Infectious Disease.2010;68:468-470. DOI: 10.1016/j.diagmicrobio.2010.07.021
  58. 58. Hooper LV, Gordon JI. Commensal host-bacterial relationships in the gut. Science. 2001;292(5519):1115-1118. DOI: 10.1126/science.1058709
  59. 59. Furtado GH, d’Azevedo PA, Santos AF, Gales AC, Pignatari AC, Medeiros EA. Intravenous polymyxin B for the treatment of nosocomial pneumonia caused by multidrug-resistant Pseudomonas aeruginosa. International Journal of Antimicrobial Agents. 2007;30(4):315-319. DOI: 10.1016/j.ijantimicag.2007.05.017
  60. 60. Kallel H, Hergafi L, Bahloul M. Safety and efficacy of colistin compared with imipenem in the treatment of ventilator-associated pneumonia: a matched case-control study. Intensive Care Medicine. 2007;33(7):1162-1167. DOI: 10.1007/s00134-007-0675-2
  61. 61. Gunderson BW, Ibrahim KH, Hovde LB, Fromm TL, Reed MD, Rotschafer JC. Synergistic activity of colistin and ceftazidime against multiantibiotic-resistant Pseudomonas aeruginosa in an in vitro pharmacodynamic model. Antimicrobial Agents and Chemotherapy.2003;47(3):905-909. DOI: 10.1128/aac.47.3.905-909.2003
  62. 62. Zavascki AP, Goldani LZ, Li J, Natio RL. Polymyxin B for the treatment of multidrug-resistant pathogens: a critical review. Antimicrobial Agents and Chemotherapy. 2007 ;60(6):1206-1215. DOI: 10.1093/jac/dkm357
  63. 63. D’Souza BB, Padmaraj SR, Rekha PD, Tellis RC, Prabhu S, Pothen P. In vitro synergistic activity of colistin and ceftazidime or ciprofloxacin against multidrug-resistant clinical strains of Pseudomonas aeruginosa. Microbial Drug Resistance. 2014;20(6):550-554. DOI: 10.1089/mdr.2014.0006
  64. 64. Bialvaei AZ, Samadi Kafil H. Colistin, mechanisms and prevalence of resistance. Current Medical Research and Opinion. 2015;31(4):707-721. DOI: 10.1185/03007995.2015.1018989
  65. 65. Falagas ME, Kastoris AC, Karageorgopoulos DE, Rafailidis PI. Fosfomycin for the treatment of infections caused by multidrug-resistant non-fermenting gram-negative bacilli: a systematic review of microbiological, animal and clinical studies. International Journal of Antimicrobial Agents. 2009;34(2):111-120. doi: 10.1016/j.ijantimicag.2009.03.009
  66. 66. Patwardhan V, Singh S. Fosfomycin for the treatment of drug resistant urinary tract infections: potential of an old drug not explored fully. International Urology and Nephrology. 2017;49(9):1637-1643. DOI: 10.1007/s11255-017-1627-6
  67. 67. Okazaki M, Suzuki K, Asano N. Effectiveness of fosfomycin combined with other antimicrobial agents against multidrug-resistant Pseudomonas aeruginosa isolates using the efficacy time index assay. Journal of Infection and Chemotherapy . 2002;8(1):37-42. DOI: 10.1007/s101560200004
  68. 68. Lambert PA. Mechanisms of antibiotic resistance in Pseudomonas aeruginosa. . Royal Society of Medicine Press. 2002;95 (Suppl 41):22-26. PMCID: PMC1308633
  69. 69. Strateva T, Yordanov D. P. aeruginosa a phenomenon of bacterial resistance. International Journal of Medical Microbiology. 2009;58(Pt 9):1133-48. DOI: 10.1099/jmm.0.009142-0
  70. 70. Livermore DM. Multiple mechanisms of antimicrobial resistance in Pseudomonas aeruginosa: our worst nightmare. Clinical Infectious Diseases. 2002;34(5):634-40. DOI: 10.1086/338782
  71. 71. Masuda N, Ohya S. Cross-resistance to meropenem, cephems, and quinolones in P. aeruginosa. Antimicrobial Agents and Chemotherapy. 1992; 36(9):1847-51. DOI: 10.1128/AAC.36.9.1847
  72. 72. Mizuta M, Linkin DR, Nachamkin I. Identification of optimal combinations for empirical dual antimicrobial therapy of Pseudomonas aeruginosa infection: potential role of a Combination Antibiogram. Infection Control and Hospital Epidemiology. 2006; 27(4):413-5. DOI: 10.1086/503175
  73. 73. Rahal JJ. Novel antibiotic combinations against infections with almost completely resistant P. aeruginosa and Acinetobacter species. Clinical Infectious Diseases. 2006;43 Suppl 2:S95-99. DOI: 10.1086/504486
  74. 74. Par SY, Park HJ, Moon SM. Impact of adequate empirical combination therapy on mortality from bacteremic P. aeruginosa pneumonia. BMC Infectious Diseases.2012; 12(1):308. DOI: 10.1186/1471-2334-12-308
  75. 75. Das T, Sehar, S, Manefield, M. The roles of extracellular DNA in the structural integrity of extracellular polymeric substance and bacterial biofilm developmen. Environmental Microbiology Reports.2013; 5(6):778-86. DOI: 10.1111/1758-2229.12085
  76. 76. Donlan, R.M., 2002. Biofilms: microbial life on surfaces. Emerging Infectious Diseases 8(9):881-90. DOI: 10.3201/eid0809.020063
  77. 77. Stewart PS, Costerton JW. Antibiotic resistance of bacteria in biofilms. Lancet. 2001;358 (9276):135-138. DOI: 10.1016/S0140-6736(01)05321-1
  78. 78. Stewart PS. Mechanisms of antibiotic resistance in bacterial biofilms. International Journal of Medical Microbiology. 2002; 292(2):107-113. DOI: 10.1078/1438-4221-00196
  79. 79. Anderl JN, Franklin MJ, Stewart PS. Role of antibiotic penetration limitation in Klebsiella pneumoniae biofilm resistance to ampicillin and ciprofloxacin. Antimicrobial Agents Chemotherapy. 2000; 44(7): 1818-1824. DOI: 10.1128/aac.44.7.1818-1824.2000
  80. 80. Walters 3rd, M.C., Roe, F., Bugnicourt, A., Franklin, M.J., Stewart, P.S., 2003. Contributions of antibiotic penetration, oxygen limitation, and low metabolic activity to tolerance of Pseudomonas aeruginosa biofilms to ciprofloxacin and tobramycin. Antimicrobial Agents Chemotherapy. 2003;47(1):317-23.DOI: 10.1128/aac.47.1.317-323.2003
  81. 81. Rasamiravaka T, Labtani Q , Duez P, El Jaziri M. The formation of biofilms by Pseudomonas aeruginosa: a review of the natural and synthetic compounds interfering with control mechanisms. BioMed Research International. 2015, 759348. 5:759348. doi: 10.1155/2015/759348
  82. 82. Kang D, Turner KE, Kirienko NV. PqsA promotes Pyoverdine production via biofilm formation. Pathogens. 2017;7(1):3. DOI: 10.3390/pathogens7010003
  83. 83. Jyot J, Sonawane A, Wu W, Ramphal R. Genetic mechanisms involved in the repression of flagellar assembly by P. aeruginosa in human mucus. Molecular Microbiology. 2007; 63(4) :1026-38 DOI: 10.1111/j. 1365-2958. 2006.05573.x
  84. 84. Guidos RJ. Combating antimicrobial resistance: policy recommendations to save lives. Clinical Infectious Diseases. 2011;52(5)S397–S428. DOI:10.1093/cid/cir153
  85. 85. Ventola CL. The antibiotic resistance crisis: part 1: causes and threats. P&T. 2015;40(4):277-283. PMC4378521
  86. 86. Gould IM, Bal AM. New antibiotic agents in the pipeline and how they can help overcome microbial resistance. Virulence. 2013;4(2):185-191. DOI: 10.4161/viru.22507
  87. 87. Bartlett JG, Gilbert DN, Spellberg B. Seven ways to preserve the miracle of antibiotics. Clinical Infectious Diseases. 2013;56(10):1445-1450. DOI; 10.1093/cid/cit070
  88. 88. Economou V, Gousia P. Agriculture and food animals as a source of antimicrobial-resistant bacteria. Infection Drug Resistance. 2015; 8: 49 61. DOI: 10.2147/IDR.S5577
  89. 89. Maron DF, Smith TJ, Nachman KE. Restrictions on antimicrobial use in food animal production: an international regulatory and economic survey. Global Health. 2013;9:48. 2013 Oct 16;9:48. DOI: 10.1186/1744-8603-9-48
  90. 90. Toke O. Antimicrobial peptides: new candidates in the fight against bacterial infections. Biopolymers. 2005;80(6):717-35. DOI: 10.1002/bip.20286
  91. 91. Park SC, Park Y, Hahm KS. The role of antimicrobial peptides in preventing multidrug-resistant bacterial infections and biofilm formation. International Journal of Molecular Sciences. 2011. 12(9):5971-5992 DOI: 10.3390/ijms12095971
  92. 92. Hancock RE, Haney EF, Gill EE. The immunology of host defence peptides: beyond antimicrobial activity. Nature reviews Immunology. 2016; 16(5):321-34.DOI: 10.1038/nri.2016.29
  93. 93. Papareddy P, Kasetty G, Kalle M, Bhongir RK, Morgelin M. NLF20: an antimicrobial peptide with therapeutic potential against invasive Pseudomonas aeruginosa infection. Journal of Antimicrobial Chemotherapy.2016;71(1):170-180, DOI: 10.1093/jac/dkv322
  94. 94. Grassi L, Maisetta G, Esin S, Batoni G. Combination strategies to enhance the efficacy of antimicrobial peptides against bacterial biofilms. Frontiers of Microbiology. 2017; 8: 2409. DOI: 10.3389/fmicb.2017.02409
  95. 95. Hirt H, Gorr SU. Antimicrobial peptide GL13K is effective in reducing biofilms of P. aeruginosa. Antimicrobial Agents Chemotherapy. 2013; 57(10): 4903-4910. DOI: 10.1128/AAC.00311-13
  96. 96. Zheng Z, Tharmalingam N, Liu Q , Jayamani E, Kim W. Synergistic efficacy of Aedes aegypti antimicrobial peptide Cecropin A2 and Tetracycline against Pseudomonas aeruginosa. Antimicrobial Agents Chemotherapy. 2017; 61(7): DOI: 10.1128/AAC.00686-17
  97. 97. Clokie MR, Millard AD, Letarov AV, Heaphy S. Phages in nature. Bacteriophage. 2011;1(1):31-45. DOI: 10.4161/bact.1.1.14942
  98. 98. Kutateladze M, Adamia R. Phage therapy experience at the Eliava Institute. Médecine et Maladies Infectieuses. 2008; 38(8): 426-430. DOI: 10.1016/j.medmal.2008.06.023
  99. 99. Villarroel J, Larsen M, Kilstrup M: Metagenomic Analysis of Therapeutic PYO Phage Cocktails from 1997 to 2014. Viruses. 2017; 9(11): pii: E328. DOI: 10.3390/v9110328
  100. 100. Jennes S, Merabishvili M, Soentjens P. Use of bacteriophages in the treatment of colistin-only-sensitive Pseudomonas aeruginosa septicaemia in a patient with acute kidney injury-a case report. Critical Care. 2017; 21(1): 129. DOI: 10. 1186/s13054-017- 1709-y
  101. 101. Ly-Chatain MH. The factors affecting effectiveness of treatment in phages therapy. Frontiers Microbiology. 2014; 5: 51. DOI: 10.3389/fmicb.2014.00051
  102. 102. Pires DP, Vilas Boas D, Sillankorva S, Azeredo J. Phage therapy: a step forward in the treatment of P. aeruginosa infections. Journal of virology. 2015;89(15):7449-56. DOI: 10.1128/JVI.00385-15
  103. 103. Morello E, Saussereau E, Maura D, Huerre M, Touqui L, Debarbieux L. Pulmonary bacteriophage therapy on Pseudomonas aeruginosa cystic fibrosis strains: first steps towards treatment and prevention. PloS one. 2011;6(2).DOI: 10.1371/journal.pone.0016963
  104. 104. Pires DP, Cleto S, Sillankorva S, Azeredo J, Lu TK. Genetically engineered phages: a review of advances over the last decade. Microbiology and Molecular Biology Reviews. 2016;80(3):523-43. DOI: 10.1128/MMBR.00069-15
  105. 105. Vandenheuvel D, Lavigne R, Brussow H. Bacteriophage therapy: advances in formulation strategies and human clinical trials. Annual Review of Virology. 2015; 2(1):599-618. DOI: 10.1146/annurev-virology-100114-054915
  106. 106. Wright A, Hawkins CH, Anggard EE, Harper DR. A controlled clinical trial of a therapeutic bacteriophage preparation in chronic otitis due to antibiotic-resistant P aeruginosa; a preliminary report of efficacy. Clinical Otolaryngology. 2009;34(4):349-57. DOI: 10.1111/j.1749-4486.2009.01973.x
  107. 107. Christiaen SE, Matthijs N, Zhang XH, Nelis HJ, Bossier P, Coenye T. Bacteria that inhibit quorum sensing decrease biofilm formation and virulence in Pseudomonas aeruginosa PAO1. Pathogens and Disease. 2014; 70(3)217-279. DOI: 10.1111/2049-632X.12124
  108. 108. Rutherford ST, Bassler BL. Bacterial quorum sensing: its role in virulence and possibilities for its control. Cold Spring Harb Perspective in Medicine. 2012; 2(11): a012427. DOI: 10.1101/cshperspect.a012427
  109. 109. Kalia VC. Quorum sensing inhibitors: an overview. Biotechnology advances . 2013;31(2):224-245. DOI: 10.1016/j.biotechadv.2012.10.004
  110. 110. Gokalsin B, Aksoydan B, Erman B, Sesal NC. Reducing virulence and biofilm of Pseudomonas aeruginosa by potential Quorum Sensing Inhibitor Carotenoid: Zeaxanthin. Microbial ecology. 2017 Aug;74(2):466-473.DOI: 10.1007/s00248-017-0949-3
  111. 111. Paczkowski JE, Mukherjee S, McCready AR, Cong JP, Aquino CJ. Flavonoids suppress Pseudomonas aeruginosa virulence through Allosteric inhibition of Quorum-sensing receptors. The Journal of biological chemistry. 2017 ;292(10):4064-4076. DOI: 10.1074/jbc.M116.770552
  112. 112. Ma L, Terwilliger A, Maresso AW. Iron and zinc exploitation during bacterial pathogenesis. Metallomics. 2015;7(12):1541-1554. DOI: 10.1039/c5mt00170f
  113. 113. Reid DW, Carroll V, O’May C, Champion A, Kirov SM. Increased airway iron as a potential factor in the persistence of Pseudomonas aeruginosa infection in cystic fibrosis. The European Respiratory Journal. 2007;30(2):286-92. DOI: 10.1183/09031936.00154006
  114. 114. Cornelis P, Dingemans J. Pseudomonas aeruginosa adapts its iron uptake strategies in function of the type of infections. Frontiers in Cellular and Infection Microbiology. 2013; 3:75. DOI: 10. 3389/fcimb. 2013.00075
  115. 115. O’May CY, Sanderson K, Roddam LF, Kirov SM, Reid DW. Iron-binding compounds impair Pseudomonas aeruginosa biofilm formation, especially under anaerobic conditions. Journal of Medical Microbiology. 2009; 58(Pt 6):765-73. DOI: 10.1099/jmm.0.004416-0
  116. 116. Minandri F, Bonchi C, Frangipani E, Imperi F, Visca P. Promises and failures of gallium as an antibacterial agent. Future Microbiology. 2014;9(3):379-97. DOI: 10.2217/fmb.14.3
  117. 117. Jeevanandam J, Barhoum A, Chan YS, Dufresne A, Danquah MK. Review on nanoparticles and nanostructured materials: history, sources, toxicity and regulations. Beilstein Journal of Nanotechnology. 2018;9:1050-1074. DOI: 10.3762/bjnano.9.98. eCollection 2018
  118. 118. Wang L, Hu C, Shao L. The antimicrobial activity of nanoparticles: present situation and prospects for the future. International Journal of Nanomedicine. 2017; 12: 1227-1249. Doi: 10. 2147/IJN.S121956
  119. 119. Salomoni R, Leo P, Montemor AF, Rinaldi BG, Rodrigues M. Antibacterial effect of silver nanoparticles in P. aeruginosa. Nanotechnology Science and Applications. 2017;10:115-121. DOI: 10.2147/NSA.S133415
  120. 120. Brown AN, Smith K, Samuels TA, Lu J, Obare SO, Scott ME. Nanoparticles functionalized with ampicillin destroy multiple-antibiotic-resistant isolates of Pseudomonas aeruginosa and Enterobacter aerogenes and methicillin-resistant Staphylococcus aureus. Applied and Environmental Microbiology. 2012; 78(8): 2768-2774. DOI: 10.1128/AEM.06513-11
  121. 121. FAO/WHO. Health and nutritional properties of probiotics in food including powder milk with live lactic acid bacteria. pp. 1-4. Food and Agriculture Organization of the United Nations
  122. 122. Marco ML, Pavan S, Kleerebezem M. Towards understanding molecular modes of probiotic action. Current Opinion in Biotechnology. 2006;17(2):204-10. DOI: 10.1016/j.copbio.2006.02.005
  123. 123. Ubeda C, Pamer EG. Antibiotics, microbiota, and immune defense. Trends in Immunology. 2012;33(9):459-466. DOI: 10.1016/j.it.2012.05.003
  124. 124. Xu K, Cai H, Shen Y, Ni Q , Chen Y, Hu S, Li L. [Management of corona virus disease-19 (COVID-19): the Zhejiang experience]. Journal of Zhejiang University. Medical Sciences. 2020;49(1):147-157. DOI: 10.3785/j.issn.1008-9292.2020.02.02
  125. 125. Priebe, G.P., Goldberg JB. Vaccines for Pseudomonas aeruginosa: a long and winding road. Expert Review of Vaccines. 2014;13(4):507-519. DOI: 10.1586/14760584.2014.890053
  126. 126. Doring G, Pier GB. Vaccines and immunotherapy against Pseudomonas aeruginosa. Vaccine. 2008;26(8):1011-1024. DOI: 10.1016/j.vaccine.2007.12.007
  127. 127. Gellatly SL, Hancock RE. Pseudomonas aeruginosa: new insights into pathogenesis and host defenses. Pathogens and disease. 2013;67(3):159-73. DOI: 10.1111/2049-632X.12033
  128. 128. Grimwood K., Kyd JM, Owen SJ, Massa HM, Cripps AW. Vaccination against respiratory P. aeruginosa infection. Human Vaccines & Immunotherapeutics. 2015;11(1): 14-20. DOI: 10. 4161/hv.34296
  129. 129. Pena C, Suarez C, Tubau F, Dominguez A, Sora M, Pujol M, Gudiol F, Ariza J. Carbapenem-resistant P. aeruginosa: factors influencing multidrug-resistant acquisition in non-critically ill patients. European Journal of Clinical Microbiology & Infectious Diseases. 2009;28(5):519-22. DOI: 10.1007/s10096-008-0645-9
  130. 130. Garnacho-Montero J, Sa-Borges M, Sole-Violan J, Barcenilla F, Escoresca-Ortega A, Ochoa M, Cayuela A, Rello J. Optimal management therapy for P. aeruginosa ventilator-associated pneumonia: an observational, multicenter study comparing monotherapy with combination antibiotic therapy. Critical Care Medicine. 2007;35(8):1888-95. DOI: 10. 1097/01. CCM. 0000275389. 31974. 22
  131. 131. Greer, N.D. Doripenem (Doribax): the newest addition to the carbapenems. Proceedings / Baylor University Medical Center. 2008;21(3):337-341. DOI: 10.1080/08998280.2008.11928422
  132. 132. Riera E, Cabot G, Mulet X, Garcia-Castillo M, del Campo R. P. aeruginosa carbapenem resistance mechanisms in Spain: impact on the activity of imipenem, meropenem and doripenem. The Journal of Antimicrobial Chemotherapy. 2011;66(9):2022-2027. doi: 10.1093/jac/dkr232
  133. 133. Hilas O, Ezzo DC, Jodlowski TZ. Doripenem (doribax), a new carbapenem antibacterial agent. P&T. 2008. 33(3):134-180. PMCID: PMC2730083
  134. 134. Cox G, Ejim L, Stogios, PJ, Koteva K, Bordeleau E. Plazomicin retains antibiotic activity against most aminoglycoside modifying enzymes. ACS infectious diseases. 2018;4(6):980-987. DOI: 10.1021/acsinfecdis.8b00001
  135. 135. Karaiskos I, Souli M, Giamarellou H. Plazomicin: an investigational therapy for the treatment of urinary tract infections. Expert Opinion on Investigational Drugs. 2015;24(11):1501-11. DOI: 10.1517/13543784.2015.1095180
  136. 136. Srinivas N, Jetter P, Ueberbacher BJ, Werneburg M, Zerbe K. Peptidomimetic antibiotics target outer-membrane biogenesis in Pseudomonas aeruginosa. Science. 2010;327(5968):1010-1013. DOI: 10.1126/science.1182749
  137. 137. Zheng Panga, Renee Raudonisb, Bernard R. Glickc, Tong-Jun Lina bd, Zhenyu Cheng. Antibiotic resistance in Pseudomonas aeruginosa: mechanisms and alternative therapeutic strategies. Biotechnology Advances. 2019;37(1):177-192. DOI: 10.1016/j.biotechadv.2018.11.013
  138. 138. Michael EC, Margaret RD, Alina MH, Monica CG. Novel Therapeutic Strategies Applied to P. aeruginosa Infections in Cystic Fibrosis. Materials 2019, 12, 4093; DOI:10.3390/ma12244093
  139. 139. Zhanel GG, Chung P, Adam H, Zelenitsky S, Denisuik A, Schweizer F, Lagace-Wiens PR, Rubinstein E, Gin AS, Walkty A, Hoban DJ, Lynch JP III, Karlowsky JA. Ceftolozane/tazobactam: a novel cephalosporin/beta-lactamase inhibitor combination with activity against multidrug-resistant gram-negative bacilli. Drugs. 2014;74(1):31-51. DOI: 10.1007/s40265-013-0168-2
  140. 140. Solomkin J, Hershberger E, Miller B, Popejoy M, Friedland I, Steenbergen J, Yoon M, Collins S, Yuan G, Barie PS, Eckmann C. Ceftolozane/tazobactam plus metronidazole for complicated intra-abdominal infections in an era of multidrug resistance: results from a randomized, double-blind, phase 3 trial (ASPECT-cIAI). Clinical Infectious Diseases. 2015; 60(10): 1462-1471. DOI: 10. 1093/cid/civ097
  141. 141. Aktas Z, Kayacan C, Oncul O. In vitro activity of avibactam (NXL104) in combination with beta-lactams against Gram-negative bacteria, including OXA-48 beta-lactamase-producing K. pneumoniae. International Journal of Antimicrobial Agents. 2012;39(1):86-9. DOI: 10.1016/j.ijantimicag.2011.09.012
  142. 142. Torres A, Zhong N, Pachl J, Timsit JF, Kollef M, Chen Z, Song J, Taylor D, Laud PJ, Stone GG, Chow JW. Ceftazidime-avibactam versus meropenem in nosocomial pneumonia, including ventilator-associated pneumonia (REPROVE): a randomised, double-blind, phase 3 non-inferiority trial. The Lancet. Infectious Diseases. 2018;18(3):285-95. DOI: 10.1016/S1473-3099(17)30747-8
  143. 143. Lapuebla A, Abdallah M, Olafisoye O, Cortes C, Urban C, Landman D, Quale J. Activity of imipenem with relebactam against gram-negative pathogens from New York City. Antimicrobial Agents and Chemotherapy. 2015; 59 (8): 5029-31. DOI: 10.1128/AAC. 00830-15
  144. 144. Wright H, Bonomo RA, Paterson DL. New agents for the treatment of infections with Gram-negative bacteria: restoring the miracle or false dawn. Clinical Microbial Infection. 2017;23(10):704-12. DOI: 10.1016/j.cmi.2017.09.001

Written By

Rahman Laibi Chelab

Submitted: October 3rd, 2020 Reviewed: December 12th, 2020 Published: June 9th, 2021