1. Introduction
Skin wounds are considered a serious public health problem, resulting in social, psychological and economic consequences [1,2]. Since wounds impair the anatomical continuity of the skin, they substantially increase the risk of microbial contamination, as the lesions constitute a gateway for microorganisms [3,4,5]. In fact, wounds induced by prolonged hospitalizations and surgical interventions have a strong association with healthcare-related infections [6,7].
Once the tissue integrity is impaired, a cascade of biochemical reactions, known as the healing process, is activated to repair the damage [8,9,10]. The healing pathway consists of distinct and overlapping phases comprising homeostasis, inflammation, proliferation, re-epithelialization and tissue remodeling [9,11]. The presence of pathogenic microorganisms extends the inflammation period which is characterized by the exacerbated release of inflammatory mediators in response to bacterial persistence, closely associated to biofilm formation [12,13,14].
Moreover, the cytotoxic action of bacterial virulence determinants results in cell damage and this may amplify the inflammation [15,16,17,18]. The prolongation of the inflammatory phase results in an impairment of the healing process [12,14,19]. In this sense, microbial infections are highlighted as the most important causes of chronic wounds and are usually associated with biofilm formation, which are notoriously recalcitrant to conventional antibiotics [20,21]. The class of microorganisms known as ESKAPE (Enterococcus faecium, Staphylococcus aureus, Klebsiella pneumoniae, Acinetobacter baumannii, Pseudomonas aeruginosa and Enterobacter sp.) are among the most prevalent bacteria in cutaneous infections [22,23,24]. The dynamics of bacterial infection of the skin are illustrated in Figure 1.
Despite the development of several antimicrobial formulations (containing silver derivatives, mupirocin, fusidic acid, mafenide, gentamicin, bacitracin, neomycin and polymyxin B), the treatment of ESKAPE-related skin infections is a huge challenge [25,26,27]. This scenario is due the ability of ESKAPE bacteria to acquire profiles of multidrug-resistance (MDR), extensive drug-resistance (XDR) and pandrug-resistance (PDR) [18,28,29]. Indeed, resistance determinants and plasmids mediating resistance towards topical antibiotics such as mupirocin, fusidic acid and neomycin [30,31,32] and silver have been detected in clinical isolates of ESKAPE bacteria [33,34].
This descriptive review aims to discuss the involvement of ESKAPE pathogens in wound infections, highlighting the major virulence factors involved in colonization and tissue damage and the most frequently detected antimicrobial resistance genes. We also provide an overview of skin physiology and the participation of resident cells and professional immune cells in pathogen detection and the healing process. 2. Fundamentals of Skin Physiology and Immunology
The skin constitutes a physical barrier formed by juxtaposed cells that cover the whole body and protect it from environmental variations and traumas [35,36]. The current conception describes the skin as an organ actively involved with the metabolism of macromolecules, and as part of the immune, nervous and endocrine systems. Two distinct and tightly joined layers make up the skin: the epidermis (more superficial) and the dermis (deeper). A third layer, called the hypodermis, is located deeper and consists mainly of adipose tissue [37,38].
The proper functioning of the skin requires close communication and collaboration between various cell types including the stromal cells (keratinocytes, fibroblasts, endothelial cells and adipocytes) as well as those derived from bone marrow (dendritic cells, macrophages, natural killer cells, mast cells, T cells and others) [39,40]. Therefore, this complex organ has a variety of resident cells that play critical roles in detecting invasive organisms (or preventing infections) [41,42].
Molecular signals called damage-associated molecular patterns (examples of DAMPs include ATP, nucleic acids and HMGB1 (high mobility group box 1 protein)) are released from damaged cells. DAMPs can be detected through molecular pattern receptors present in the skin-resident cells [43,44,45,46,47]. These receptors can also identify pathogen-associated molecular patterns (PAMPs), such as peptidoglycans [41,48,49]. The main structures involved in the recognition of DAMPs and PAMPs are the so-called Toll-type receptors. The detection of these receptors by DAMPs and PAMPs activates several mechanisms resulting in the release of pro-inflammatory mediators (such as cytokines, nitric oxide) and the secretion of antimicrobial compounds [48,50,51,52].
Altogether, the skin cells and immune cells form the concept of the skin-associated lymphoid tissue (SALT), which acts as a tertiary lymphoid organ [53,54]. Another recent concept is the inducible SALT (iSALT), which denotes that the leukocytes (such as perivascular macrophages, dermal dendritic cells and T cells) involved in this structure are activated by local inflammatory stimuli [55,56]. iSALT formation is associated with activation of perivascular macrophages by IL-1a, a cytokine produced by keratinocytes in response to inflammatory agents. The activated macrophages produce CXCL2 (chemokine (C-X-C motif) ligand 2), a chemoattractant chemokine that recruits dermal dendritic cells and promotes effective antigen presentation and activation of T cells in the skin [54,55,56,57].
It is also important to emphasize the immunological relevance of the skin-associated microbiota [58,59]. These microbes cooperate with the skin and immune cells in order to maintain tissue homeostasis, for example, contributing to the effective development of innate and adaptive immune responses [60,61]. The microorganisms residing in the skin can interact through antagonistic or synergistic relationships [61]. For instance, the presence of microorganisms that metabolize host proteins and lipids results in the production of bioactive substances that act by inhibiting the proliferation of invading pathogens. They do this through the induction of immune mediators, such as IL-1 and IFN-γ, released from keratinocytes and resident T cells, respectively [58,62]. In addition, some studies show that several commensal species act to inhibit the proliferation of other pathogenic (or opportunists) bacteria, such as the relationship between Corynebacterium sp./S. pneumoniae, S. epidermidis/S. aureus and S. epidermidis/Propionibacterium acnes [61].
2.1. Immune Cells in the Epidermis
The epidermis mainly consists of keratinocytes that are closely linked to each other, forming a barrier and limiting access to the internal environment [63]. The keratinocytes play essential roles in the inflammatory response due to the secretion of cytokines (TNF-α, IL-1, IL-6) and chemokines (TARC/CCL17) [64,65], modulating the functions of T lymphocytes [66]. The Langerhans cells (LCs) are presented in the upper layer and combine features of macrophages (self-renewal, embryonic origin) and dendritic cells (antigen presentation, dendrites) [67,68]. The antigens in the lower layer of the human epidermis are captured by inflammatory dendritic epidermal cells (IDECs) [69].
The human epidermal resident T cells participate in immune surveillance and quickly respond to antigens from pathogens or damaged host cells. These cells participate in wound healing by expression of insulin-like growth factor 1 (IGF-1) [70,71]. Further, the sweat glands (epidermal appendages) are able to secrete antimicrobial peptides [36]. Macrophages and memory B cells can be also found in the epidermis, while neutrophils are recruited in response to tissue damage [41,42,72,73,74].
The macrophages assume different phenotypes that play distinct roles in skin physiology [75,76,77]. Differentiation to each phenotype is dependent on the cytokine involved in each situation [78]. The classical macrophages (also called M1), activated by INF-γ, are involved in phagocytosis and the release of inflammatory mediators such as cytokines (IL-1β, IL-6, IL-12 and TNF-α) and reactive species (nitric oxide and superoxide). M1 macrophages are crucial for the antimicrobial response and are characterized by the expression of MHC (major histocompatibility complex) class II receptor called HLA-DR (Human Leukocyte Antigen–DR isotype) [76,78,79,80].
The alternative activation of macrophages (M2) is trigged by IL-4, IL-5 and IL-13 (Th2 characteristic cytokines) and these macrophages express CD163 and arginase. This phenotype produces IL-10 and TGF-β, which are involved in the later phase of healing, promoting tissue repair [75,76,78,81]. In addition to their involvement in the reverse migration of neutrophils, the M2 macrophages induce fibroblast proliferation and collagen production [77,82,83]. Finally, the recently described M4 macrophages (induced by the chemokine CXCL4) are involved in the pathogenesis of skin lesions induced by Mycobacterium leprae [84].
Neutrophils are the first cells attracted to the wound site in response to chemotactic factors released by the skin resident cells (macrophages and keratinocytes), DAMPs and lipid mediators (such as leukotriene B4) [47,83,85]. These cells are responsible for the processes of sterilization and the degradation of cell debris through phagocytosis, neutrophil extracellular traps (NET) and the secretion of antimicrobial peptides and inflammatory mediators (reactive species and cytokines) [9,10,46,83].
Neutrophils also produce serine proteases and matrix metalloproteinases (MMPs); enzymes that are crucial for the correct progress of the healing process [86,87]. However, the high activity of these enzymes, along with the exacerbated release of inflammatory mediators, can promote tissue damage and contribute to the formation of chronic wounds [47,88,89]. Neutrophils are essential for effective wound healing by influencing M2 polarization through the release of cytokines and soluble factors (azurocidin, cathepsin G, colony stimulating factor 1, IL-13) [90,91]. The uptake of dead neutrophils and the secretion of microvesicles can also trigger the release of anti-inflammatory cytokines (such as TGF-β) by macrophages and promote wound repair [82,91].
Regarding B cells, recent data indicate that skin subpopulations are different from lymph node B cells, and that they are important in the regulation of inflammation and wound healing [73,92]. Immunosuppressive functions are attributed to a subset called B regulatory cells (Bregs) that produce anti-inflammatory cytokines IL-10, IL-35 and transforming growth factor-β (TGF-β), thus limiting the activation of inflammatory cells [93,94,95]. Bregs are involved in the differentiation of regulatory T cells [95].
2.2. Immune Cells in the Dermis
The dermis is composed of extracellular matrix proteins that give structure and elasticity to the skin. This structure provides nutrients and circulatory support to the epidermis [41,53]. Fibroblasts are the main cell types of the dermis that perform the synthesis of collagen, elastin and amorphous—fundamental substances for extracellular matrix formation [41]. These cells repair injured skin by providing structural support and guiding the migration of immune cells, allowing important cell–cell contact [96,97].
Fibroblasts are also able to produce cytokines (such as IL-1β, IL-6) and chemokines (such as CXCL8 and CXCL11) [98,99]. These mediators can actively recruit leukocyte subpopulations of the innate immune system, such as plasmacytoid dendritic cells (pDC), neutrophils, mast cells, and macrophages. This step is a crucial for the inflammatory events that follow the course of chronic inflammation, such as in the atopic dermatitis [100,101,102,103].
Other cells present in the dermis include natural killer cells, B lymphocytes and T lymphocytes [104]. T regulatory cells (Treg) are attracted by chemokine CCL20, which is produced in response to commensal microorganisms present in the skin after the birth [39]. Treg cells interact with Langerhans cells and are involved in the resolution of skin inflammation, promoting the proper healing process [105,106]. Langerhans cells also inhibit Treg during microbial invasion in order to promote inflammatory defense, which also includes the proliferation of memory cells [106].
3. ESKAPE and Wound Infections
As mentioned earlier, pathogenic microorganisms significantly slow the healing process due to tissue destruction that leads to an exacerbated immune response condition, characterizing chronic wounds [48,50]. In the following sections, some virulence determinants directly related to the action of ESKAPE pathogens in skin infections are discussed. We also address the antibiotic resistance genes that are more prevalent in the isolates related to skin infections.
3.1. Enterococcus faecium and Related Species
Enterococcus (Enterococcaceae family) species are predominantly non-pathogenic gastrointestinal commensal bacteria that, in certain circumstances, cause infections. However, some species of the genus have shown clinical relevance in the last decade, such as Enterococcus faecalis and Enterococcus faecium, both involved in wound infections [107,108,109]. Additionally, Enterococcus gallinarum and Enterococcus casseliflavus/flavescens (with intrinsic resistant towards vancomycin) have also gained attention due their involvement in surgical wound infections [110,111,112,113].
E. faecium is the representative member of Enterococcus genus in the ESKAPE group [114]. It has been isolated in surgical site infections and diabetic foot [115,116]. For example, a study conducted in India showed that E. faecium was the most commonly observed Enterococcus species in traumatic skin wounds; followed by E. faecalis [111]. Similarly, another study reported that E. faecium was the main etiologic agent in skin and soft-tissue infections (SSTI) related to combat casualties [117]. E. faecium is also involved in polymicrobial infections with E. coli [116] and E. faecalis [111]. Specifically, the association of E. faecium in diabetic foot ulcers was shown to be related with limb loss [116].
In addition, a recent study showed that some diabetic patients with wounds infected by Enterococcus presented evolution to osteomyelitis. This was a retrospective study of 275 patients with diabetic foot admitted at a tertiary care hospital in the UK in 2015. Enterococcus species, including vancomycin-resistant enterococci (VRE) strains accounted for 17% of Gram-positive isolates [118].
3.1.1. Main Genes Involved in Enterococcus faecium Resistance
E. faecium covers highly virulent strains, such as those of the clonal complex 17 (CC17), which have a multiple antimicrobial resistance profile due to the presence of several genes, such as vanA (vancomycin resistance) and poxtA [119,120,121]. This latter gene comes from mobile elements and has been frequently reported to confer resistance to phenicols, tetracycline and even linezolid, the last drug of choice for VRE strains [122,123,124]. E. faecium has the ability to survive in highly hostile and nutrient-poor environments [125,126]. These characteristics are also observed for E. faecalis strains [127].
Both E. faecium and E. faecalis have intrinsic resistance to cephalosporins, aminoglycosides, clindamycin, and trimethoprim/sulfamethoxazole [128,129,130]. VRE strains are recognized as major issues and it is estimated that these strains were responsible for infections in over 16,000 people with over a thousand deaths (1081) in European countries in 2015 [131]. Moreover, MDR enterococci are also present in coastal and fluvial waters, which can become a major public health problem due to the possibility of their transfer and arrival in the clinic [132].
Vancomycin resistance in E. faecium is related to van gene clusters that comprise the operons vanA, vanB, vanC, vanD, vanE, vanG, vanL, vanM and vanN [133,134,135]. Some vanA and vanB accessory genes (vanY, vanZ and vanW) have also been described and the entire cassettes can be carried by transposons (Tn1546, Tn1547 or Tn1549) [136,137]. All these genes are easily deposited on plasmids or inserted directly into the major chromosome [138,139]. The van operons are also involved in resistance to other drugs, as they have genetic variability that enables a variety of resistance phenotypes [133].
Further, specific resistance to aminoglycosides, especially gentamicin, is provided by aminoglycoside modifying enzymes (AMEs) which are encoded by genes such as aac(6’)-Ie-aph(2”)-Ia, aph(3’)-IIIa and ant(4’)-Ia, which make it impossible to bind the drug to bacterial ribosomes and consequently, the protein synthesis is inhibited [140]. AMEs are also involved in resistance towards erythromycin, tetracycline and ciprofloxacin [141].
Recently, the prevalence of E. faecalis was determined in a study involving 200 surgical wound samples obtained from patients of Minia University hospital, Egypt. A frequency of 24 (12%) was reported for wound samples. All E. faecalis isolates were classified as MDR. Specifically, the rates of resistance towards erythromycin, vancomycin and linezolid were 100, 58.28 and 23.1% of the isolates from wound samples, respectively. In addition, the vanA gene was detected in 71.4% of vancomycin-resistant isolates. Similarly, the majority of the strains harbored resistance genes ere(B) and erm(B)—83.3% and 70.8%, respectively—responsible for the production of esterase enzymes for erythromycin [142].
The combination of existing drugs is an important strategy towards drug resistant strains of E. faecium and related species. The use of daptomycin with β-lactams has been well documented against VRE strains, including in murine models of infection [143,144,145]. Combinatory treatment with linezolid, quinupristin-dalfopristin, tigecycline and, more recently, oritavancin and dalbavancin also showed excellent results against resistant strains [146]. The combination of retapamulin with erythromycin, quinupristin/dalfopristin and quinupristine also demonstrated synergistic activity against E. faecalis [147]. Furthermore, the use of new drug candidates has also been explored. Compounds such as 1,2,4-oxadiazoles are considered to have therapeutic potential for the treatment of E. faecium MDR strains [148]. Similarly, 1,2,4-triazolo[1,5-a] pyrimidines were able to prevent the cell wall biosynthesis of E. faecium [149]
3.1.2. Main Genes Involved in Enterococcus faecium Virulence
E. faecium also display a vast repertoire of virulence determinants that are involved in tissue adhesion and cytotoxicity. The production of biofilms by E. faecium is associated with multiple factors, such as esp (suggested as the major virulence determinant) which encodes the Enterococcus surface protein (Esp) which is related to adhesion to epithelial cells, as well as the secretion of aggregating substances [132,150]. Recent studies on the N-terminal region of Esp suggested that this protein acts by a mechanism involving amyloid-type aggregation to build the biofilm matrix in an acid environment [151].
Other virulence factors related to Enterococci adhesion and colonization include collagen-binding adhesin (encoded by ace), adhesin (efaAfm), cytolysin A (cylA), gelatinase (gelE), hyaluronidase (hyl) and emp pilus and aggregation substance (asa1) [132,152,153,154]. The collagen-binding proteins and cytolysins expressed by E. faecium compromise the bonds between collagen fibers and the balance between keratinocytes and fibroblasts [155]. Gelatinase and hyaluronidase are responsible for the hydrolysis of collagen fibers and the cutaneous extracellular matrix. The presence of an emp pilus, especially EmpA and EmpB subunits, is essential for the architecture of the pilus, formation and extension of biofilms, in addition to adhesion to fibrinogen and type I collagen [153]. Aggregating substances, on the other hand, facilitate the attachment to the skin epithelium and favor the bacterial aggregative behavior during plasmid conjugation [156].
A research study aiming to evaluate the genes associated with virulence and drug resistance determinants was performed with Enterococcus clinical isolates from burn patients. The authors reported a predominance of E. faecalis (80.7%) among the obtained enterococci (n = 57), while only two isolates were identified as E. faecium. The E. faecium strains were positive for asa1, ace and gelE [157]. Another study, also involving burn patients, reported a higher presence of E. faecalis (62.5%) and E. faecium (37.5%) among enterococcal isolates. These isolates had gelE and asa as the most detected virulence genes, while the esp and cyl showed a low level of detection. Only the E. faecium isolates exhibited resistance towards vancomycin and teicoplanin (24%). In general, higher levels of antibiotic resistance were observed in E. faecium [158].
The Table 1 illustrate some types of genes associated in resistance and virulence in E. faecium.
3.2. Staphylococcus aureus
Staphylococcus aureus naturally occurs in the microbiota of skin and other body tissues [159], facilitating the opportunistic infection of wounds [160,161]. In fact, S. aureus is one of the pathogens commonly isolated from skin lesions [162,163], with a high number of strains exhibiting complex combinations of virulence and resistance genes [3,164] (Table 2). It is an important causative agent of SSTIs, presenting high rates of morbidity and mortality, in addition to recurrent infections [165,166]. This species is also been related to the progression of diabetic foot to osteomyelitis [118].
The genomic variation in S. aureus is discontinuous, with distinct subdivisions called clonal complexes. The multifactorial forces that shape the variable structure in S. aureus are likely to include bacterial competition and barriers to genetic exchange [167,168]. Clones with high resistance to antibiotics and/or multiple virulence factors quickly emerge due to the acquisition of genes (by several routes) from other strains of S. aureus or even from other genera [169]. This plasticity allows S. aureus adaptation to different types of stress, enabling survival in different niches [170,171,172].
3.2.1. Main Genes Involved in Staphylococcus aureus Resistance
An increasing number of S. aureus strains have been found to be resistant to antimicrobial agents. This genetic variability is mediated by a diverse set of mobile genetic elements (MGEs) that include plasmids, transposons, integrons, genomic islands, S. aureus pathogenicity islands (SaPIs), integrative conjugative elements, staphylococcal chromosome cassettes (SCC) and phages [173,174].
The first resistance episode by S. aureus was reported in the 1940s for penicillin (PRSA), a period close to its own discovery and use [175]. This type of resistance has been attributed to the blaZ gene, which encodes a specific type of β-lactamase, able to cleave penicillin through the hydrolysis of its β-lactam ring [176]. Thereafter, the rate of emergence of methicillin-resistant S. aureus (MRSA) and multidrug-resistant S. aureus (MDRSA) strains has been high in SSTI in hospitalized individuals [177,178]. Although traditionally linked to the hospital environment, some MRSA and MDRSA strains have emerged in the community and have caused severe cases of skin infections [179]. This phenomenon has increased the frequency and the severity of infection by this microorganism [180].
The high prevalence of MRSA in hospitals and community settings has been a major public health challenge worldwide. An American study reported that at least 72,000 cases of invasive MRSA infections were recorded in US health systems in 2014 [181]. The acquisition of methicillin resistance occurs through the presence of mecA, a gene encoding a penicillin-binding protein (PBP2a) [176].
The discovery of mecA was only possible twenty years after the appearance of the first cases of MRSA [182]. This gene is transported by mobile elements called Staphylococcal Chromosomes Cassette mec (SCCmec) [183]. At least eleven SCCmec types have been described and correlations between more virulent strains of MRSA and SSCmec types III and IV have been observed [184,185]. SCCmec can be carried by phages [186,187]. In addition, a French study that investigated the prevalence of fluoroquinolone-resistant staphylococci (FQR) in hospitalized and healthy patients showed that this type of resistance is also associated with MRSA strains [188].
Alarming levels of resistance are already detected in isolates of S. aureus for drugs considered as the last choice for treatment, such as vancomycin [189,190]. Originating from a conjugative plasmid, resistance to vancomycin is conferred by the VRE operon vanA (previously mentioned), where the entire original enterococcal plasmid is conjugated or only the transposon Tn1546 is assigned to a resident plasmid of S. aureus [176].
The mechanism of action of vancomycin is based on the inhibition peptidoglycan polymerization, an important structural component of the bacterial cell wall [191,192]. The vanA operon—composed of the vanA, vanH, vanX, vanS, vanR, vanY and vanZ genes—is responsible for inhibiting the binding of vancomycin to peptidoglycan precursors, by either not synthesizing them or hydrolyzing those that already exist. This is regulated by a two-component system encoded by the vanS and vanR genes that activate the transcription of the operon [176].
Some other frequently reported phenotypes include borderline oxacillin-resistant S. aureus (BORSA) and vancomycin-resistant S. aureus (VRSA) strains. BORSA isolates are susceptible to cefoxitin and do not carry the mecA gene, but they are able to produce excessive amounts of β-lactamases, resulting in antimicrobial resistance [193,194]. It was also observed that resistance to oxacillin even alters the primary characteristics of the S. aureus biofilm and its virulence [195]. Strains resistant to vancomycin—one of the standard treatments for infections caused by MRSA—are emerging, and their behavior is attributed mainly to the vanA operon present in a plasmid derived from enterococci [176]. VRSA isolates have already been identified in skin lesions and diabetic foot ulcers [196].
Other MDRSA strains described harbored several resistance genes, such as: blaR1, blaIe, lmrS, vraR, mrgA, qacA and qacB for oxacillin and ciprofloxacin; NorA, which belongs to the major facilitator superfamily (MFS), and MepA, which belongs to the multidrug and toxic compound extrusion (MATE), for ciprofloxacin and norfloxacin; MdeA (MFS), related to resistance to novobiocin, mupirocin and fusidic acid; and LmrS, which encodes multiple drug efflux pumps, associated with trimethoprim and chloramphenicol [197,198,199,200].
As an alternative to the high rates of resistance and emergence of MDR strains, several new drugs have been used to treat infections caused by S. aureus. Recently, several agents have been approved to treat MRSA-infected skin lesions, including the lipoglycopeptides dalbavancin, oritavancin and telavancin, ceftaroline and tedizolid [201]. Other examples of new drugs include tannic acid, ivermectin and quinupristin/dalfopristin, which demonstrate success in combating S. aureus strains that are resistant to methicillin, erythromycin, ciprofloxacin, rifampicin and gentamicin [202,203,204].
Similarly, preliminary studies have shown that peptides such as nisin, AP7121, CSαβ-DLP2 and DLP4 demonstrate antibacterial effects against S. aureus (including MDRSA and VRSA strains). These compounds act by interrupting the molecular synthesis and microbial cell cycle [205,206,207]. Several natural products are also reported to have promising in vivo antimicrobial activity against S. aureus, including in models of wound infections [208,209,210,211].
3.2.2. Main Genes Involved in Staphylococcus aureus Virulence
As previously mentioned, S. aureus can express a variety of virulence factors that facilitate cell adhesion, mediate evasion from the immune system and induce damage to host cells [161,173,212]. Adhesion to host cells is ensured by proteins that bind to fibronectin (FnbA and FnbB), collagen (Cna), fibrinogen (Fib), laminin (Eno) and elastin (EbpS). These proteins can be referred as microbial surface components recognizing adhesive matrix molecules (MSCRAMMs) that play important roles in the evasion of immune defenses and biofilm formation [213,214].
Indeed, great capacities for both biofilm formation and intracellular survival are described for S. aureus [165,166]. These properties are related to the firm and recalcitrant polysaccharide matrices that increase its virulence and resistance to antibiotics and may even contribute to bacterial survival in phagocytes (neutrophils and macrophages). Taken together, these factors contribute for the spreading of S. aureus and predispose an infected individual to chronic and persistent infection [215,216].
Other virulence factors extremely relevant to skin infections are the toxins secreted by S. aureus that provoke tissue damage and abscess formation [217]. Among them is α-toxin, a 33 kDa pore-forming cytolytic protein which affects a wide range of human cell types, including epithelial cells, endothelial cells, T cells, monocytes and macrophages [218]. In this sense, in addition to tissue damage, this toxin is able to neutralize the protective immune response [217].
Exfoliative toxins (ETs) and leukocidins—including leukocidin ED (LukED) and Panton-Valentine leukocidin (PVL)—also play important roles in the pathogenesis of S. aureus as they destroy cell membranes by creating β-barrel-like pores that lead to cell lysis. Additionally, they impair the activation of resident immune cells [219,220]. ETs selectively cleave peptide bonds in the extracellular region of human desmoglein-1, which acts as an adhesion molecule between keratinocytes. ETs are related to staphylococcal scalded skin syndrome and bullous impetigo [221,222]. The three ETs are encoded by different genetic regions: eta (found in a phage), etb (located on plasmids), etd (located on genomic islands) [221].
LukED (encoded by the lukED gene) is described as a major cause of blood and skin infections, such as impetigo [220]. In turn, the pvl gene is commonly detected in S. aureus strains isolated from SSTI and its product is responsible for the destruction of resident immune cells and tissue necrosis [219,221]. Several reports have shown a high prevalence of MRSA carrying the pvl gene in community-acquired SSTI, where some of the wounds required surgical procedures for incision or drainage, and many of these strains were also resistant to erythromycin, clindamycin and tetracycline [223,224,225].
The frequency of detection of the pvl gene may vary according to the region and clonal group of S. aureus—related to community-associated methicillin-resistant Staphylococcus aureus (CA-MRSA) strains in many countries. For instance, in a study carried out in Nigeria, a high frequency of pvl gene detection was observed for SSTI and wounds, with rates 83.3 and 79.2%, respectively [226]. However, another study carried out in Iran reported that the frequency of the pvl gene was 33.3% for CA-MRSA isolates obtained from infected wounds [227].
Phenol-soluble modulins (PSMs) have gained attention for their involvement in the inflammatory response, inducing the production of cytokines and neutrophil migration [228,229,230]. During skin infection, PSMα released by S. aureus, has also been shown to influence the levels of IL-17 produced by keratinocytes [229]. It is believed that serious SSTI associated with CA-MRSA strains, may be related to the cytotoxic and membrane-disturbing PSMα [231]. PSMα, secreted by CA-MRSA, can induce the rapid formation of a type of NET that is related to the destruction of phagocytic cells, rather than contributing to the death of pathogens [231].
A recent study evaluated the presence of virulence and resistance-related genes in isolates of S. aureus from samples of skin infections (n = 200). A total of thirty-six (18%) isolates with the MDR profile carried the mupA gene, the predominant determinants of virulence included PSMα (61.5%), pvl (2.5%), eta (2.5%) and etb (1%) [232].
In addition, S. aureus has mechanisms that are directly involved in processes related to immune response modulation, such as: SSL3 (staphylococcal superantigen-like protein 3), an inhibitor of neutrophils and other TLR2-expressing cells [233]; proteases with several targets (complement system, LL-37) [234,235]; staphylokinase protein (SAK), an inhibitor of LL-37, and α-defensins [236,237].
Some examples of genes involved in virulence and resistance of S. aureus are summarized in Table 2.
3.3. Klebsiella pneumoniae
K. pneumoniae is an opportunistic, Gram-negative, encapsulated and cosmopolitan pathogen that usually causes skin infections in burned and/or immunocompromised individuals, often forming thick biofilms [238,239]. It is considered to be one of the main causes of health-associated infections [240]. For instance, studies conducted at the US Army Surgical Research Institute (Burn Center), showed K. pneumoniae as one of the four major pathogens isolated from infected wounds in hospitalized burn patients [241,242]. Similarly, an epidemiological analysis showed that 15.1% of hospital isolates of K. pneumoniae from Turkey came from cutaneous lesions [243].
3.3.1. Main Genes Involved in Klebsiella pneumoniae Resistance
The emergence of K. pneumoniae strains with hypervirulent phenotypes (hvKp) and more aggressive capsular serotypes, such as K1 and K2 present in CC23 (clonal complex 23), has been observed to be more frequent in severe skin and soft tissue infections [244,245]. Even the reduced use of antibiotics such as aminoglycosides proved to be sufficient for the emergence of resistance phenotypes, such as the expression of the armA gene, which encodes the 16SrRNA methylase enzyme responsible for blocking binding to the bacterial ribosome [246]. Other aminoglycoside resistance genes that have already been reported include aacA4, aacC2 and aadA1 [247] (Table 3).
In addition, the appearance of carbapenem-resistant K. pneumoniae (CRKP) strains has made it very difficult to treat burn wounds that are infected with this pathogen [248]. CRKP strains express carbapenemases, a type of enzyme (encoded by genes including blaKPC-2 and blaKPC-3) that is able to cleave the β-lactam drugs and exhibits low susceptibility to the action of beta-lactamase inhibitors (clavulanic acid and tazobactam) [249,250,251].
K. pneumoniae strains also show resistance to third generation cephalosporins and fluoroquinolones, mediated by extended spectrum β-lactamases (ESBLs) [245,252]. Allied to this, changes in cell permeability represent the main mechanism involved in resistance to quinolones, through the expression of the acrAB gene—responsible for efflux pumps [246].
A Chinese survey evaluated the prevalence of carbapenem-resistant Enterobacteriaceae in various types of infection. The study showed that carbapenem-resistant K. pneumoniae was the most common etiologic agent of deep wound infections (85.7%); while it was detected in 18.8% cases of superficial wound infections. Considering all types of infections, a high prevalence of nosocomial carbapenem-resistant K. pneumoniae producing IMP-4 carbapenemase (84%) and IMP-8 carbapenemase (50%) was detected [253].
Recently, research was performed that phenotypically and molecularly characterized K. pneumoniae isolates that were obtained from wounds of hospitalized patients in Tehran, Iran. The authors reported that 45.1 and 22.5% were producers of extended-spectrum β-lactamases (ESBL) and carbapenemase, respectively [254]. The isolates simultaneously carried the genes encoding ESBL (78.4%), AMEs (aac(6’)-Ib; 65.7%), carbapenemase (50%) and quinolone resistance determinants (QDRs; 49%). The authors of this study highlighted that four isolates carried the genes for carbapenemases (blaTEM, blaSHV, blaCTX-M), QDRs (qnrB and qnrS) and aac(6’)-Ib [254].
Some strains of K. pneumoniae have even shown resistance to last generation antibiotics, such as polymyxin. Reductions in negative ions hinder the binding of the drug to the bacterial surface. This occurs due a chromosomal system of modifications in the lipopolysaccharides (LPS). These changes are attributed to central genes involved in lipid A synthesis, such as lpxM [246]. A study carried out in Korean hospitals also showed that 16% of the samples of K. pneumoniae collected were resistant to tigecycline, a drug used to treat MDR strains, with mutations in the ramR and rpsJ genes and massive expression of the tetA gene also being documented [255].
In view of this great resistance problem and the appearance of MDR strains, the treatment of K. pneumoniae becomes quite challenging. An ideal therapeutic protocol for infections caused by Multidrug-resistant K. pneumoniae (MDR-KP) has not yet been well defined, but the use of high-dose meropenem, fosfomycin, tigecycline, aminoglycosides and polymyxins is widespread [256]. The combination of colistin with niclosamide has even shown good results against strains that are already resistant to colistin itself [257].
In addition, several drugs are in the clinical stages of testing for MDR-KP, with good results having been produced so far. The association of these new candidates with marketed drugs is seen as having great potential in the fight against these pathogens. These include: ceftazidime-avibactam, meropenem-vaborbactam, imipenem-relebactam, plazomicin, cefiderocol, aztreonam-avibactam, ceftaroline-avibactam, cefepime-zidebactam and nacubactam [256]. Further, immunotherapies have been evaluated, such as the investigation of cystatins 9 and C as a solution for K. pneumoniae strains producer of metallo-β-lactamase-1 [258].
3.3.2. Main Genes Involved in Klebsiella pneumoniae Virulence
Currently, four factors have been well described as contributing to K. pneumoniae virulence: fimbriae (important for the formation and installation of biofilms), capsule, lipopolysaccharide and iron uptake [240]. For the effective formation of a biofilm, it is necessary that the microorganisms involved become close enough to the target surface, fixing themselves to it with the aid of fimbriae and/or a flagella [259]. In K. pneumoniae, fimbriae type 1 and 3 are encoded by the operon mrkABCDF, and the subunits MrkA and MrkD are directly involved in binding to collagen [260,261].
The polysaccharide capsule (encoded by the capsular polysaccharide synthesis locus—cps gene) is another important virulence factor for the establishment of skin infections, since it prevents phagocytosis and complement-mediated killing [262,263]. For K. pneumoniae, 78 capsular serotypes have been reported. In particular, K1 and K2 confer hypervirulence through the excessive production of hypermucoviscous capsular material [240]. The rmpA gene—located in plasmids of K. pneumoniae—has also been observed to be an important virulence factor and is responsible for the synthesis of capsular compounds [246].
Candan et al. (2015) also described the importance of various genes for the production of the K. pneumoniae capsule (magA, k2A and wcaG) and its lipopolysaccharides (wabG, uge and ycfM), such as LPS. The products of these genes are essential for the formation of the thick and consistent biofilms that are frequently found in the skin and soft tissue infections of burned or immunocompromised patients [264].
The outermost parts of the LPS, named O-antigens, are also used for the serotyping of K. pneumoniae. At least, nine O-antigen serotypes have been described and these structures are considered as representing potential targets for vaccination [265,266]. A study using a global collection of K. pneumoniae showed that the serotypes O1, O2 and O3 were most prevalent in all types of infections [267]. The O-antigen in the O1 serotype is composed of D-galactan I and D-galactan II, with the latter being recognized as the immunodominant antigen. The synthesis of D-galactan II is performed by glycosyltransferases WbbY and WbbZ. Interesting, D-Gal II is more prevalent in community-acquired pyogenic liver abscess (PLA) strains than in non-tissue-invasive strains [268]. In addition, some strains of K. pneumoniae can modify the composition of LPS, avoiding recognition by the TLR4 receptors [240].
As a crucial factor for the growth and infection process of K. pneumoniae, the production of enterobactin, mediated mainly by the entS gene, is directly related to the ability to chelate iron molecules from the host (siderophores). The presence of enterobactin is observed in normal and hypervirulent strains, while other molecules also involved in the iron absorption process, such as aerobactin, yersiniabactin and salmoquelin are more common only in hypervirulent strains [240,269].
Some examples of virulence and drug resistance-related genes reported for K. pneumoniae are provided in Table 3.
3.4. Acinetobacter baumannii
A. baumannii is an important opportunistic nosocomial pathogen that causes severe infections associated with ventilation and blood flow in critically ill patients, but also serious infections in patients with skin lesions, especially burns [270,271,272]. MDR A. baumannii (MDR-AB) has been associated with severe and fatal cases of SSTI [273,274,275].
A recent study highlighted that 15% of patients admitted to a particular hospital acquired nosocomial infections due to MDR-AB, a condition associated with prolonged hospitalization and increased risk of death [275]. In addition, a one-day cross-section showed the presence of A. baumannii DNA in 10% of the individuals tested [276].
A. baumannii is often found in skin and soft tissue infections resulting from burns, mechanical trauma or the wounds of soldiers from war situations. There have been reports of osteomyelitis and exposed tibial fractures caused by MDR-AB during military combat operations [277,278]. Another study that evaluated a burn care center registered that the presence of A. baumannii correlated with the worsening of health status of the infected patients. In this case, higher levels of morbidity and mortality and longer hospital stays being documented in comparison to patients with uncontaminated wounds [279].
3.4.1. Main Genes Involved in Acinetobacter baumannii Resistance
The resistance of A. baumannii to β-lactam antibiotics is mainly mediated by enzymatic hydrolysis [18]. High cleavage capacities have been observed in all known penicillins and cephalosporins (except cephamycin) by the following β-lactamases: CTX-M (encoded by blactx-mgene), GES (blages), PER (blaper), SCO (blasco), SHV (blashv), TEM (blatem) and VEB (blaveb) [280,281,282,283] (Table 4). It was also observed that A. baumannii has an extreme tolerance to free radicals (such as hydrogen peroxide), as it has ample genomic flexibility and contains the element ISAba1 upstream of the catalase katG gene, which is responsible for improving resistance [284].
On the other hand, as in most Gram-negative bacteria, the emergence of resistance to tetracycline in A. baumannii occurs through efflux pumps [285]. In this case, these are Tet type pumps, encoded by the tetA gene; whereas those encoded by the tetB gene confer resistance to tetracyclines and also minocycline and doxycycline. Resistant strains of A. baumannii carrying the ribosomal defense gene tetM have already been identified [18]. For fluoroquinolones-resistant A. baumannii strains, mutations have been observed in specific drug targets, especially in the gyrA of DNA gyrase and parC of topoisomerase IV genes [286]. Episodes of resistance by light chromosomal efflux pumps have also been reported [287].
Regarding resistance to aminoglycosides, different types of AMEs are synthetized by A. baumannii strains such as phosphotransferases, acetyltransferases and adenyltransferases. These enzymes are encoded by genes such as aac(3′)-Ia, ant(2’)-Ia and ant(3″) In addition, A. baumannii strains can harbor genes for several types of AMEs at the same time [288,289]. Resistance to aminoglycosides can also occur through the expression of the armA, rmtA, rmtB, rmtC and rmtD genes, that alter the binding to bacterial ribosomes [18]. Specifically, two efflux pumps can also affect the action of gentamicin: AdeABC and AbeM [290].
Alterations in the pmrCAB operon of A. baumannii are directly related to resistance to polymyxins, such as colistin. The pmrC gene encodes a modifying phosphoethanolamine transferase, while pmrA and prmB regulate a two-system regulatory mechanism. Mutations in these last two systems favor the expression of pmrC that is responsible for modifying lipid A [291]. More recently, it has also been reported that the lpsB, lptD and vacJ genes reduce fluidity and increase the osmotic resistance of the outer membrane of A. baumannii, inducing resistance to polymyxin A [292].
Confirming these facts, a cross-sectional descriptive study carried out in five hospitals in Medellin over a period of 2 years detected 32 patients with infections caused by MDR strains of A. baumannii. The incidence of SSTI and osteomyelitis was 21.9 and 18.7%, respectively. The authors reported a high rate of antibiotic resistance among most isolates (80%), with genes for carbapenemases oxa-23 and oxa-51 detected in all strains of skin lesions [293]. A Brazilian analysis on osteomyelitis also showed that A. baumannii was present in 21% of cases, with 40% of these being resistant to carbapenems [294].
Phage-based therapies have displayed promising results against MDR-AB strains. In vivo models showed that phage-based therapy was successful in containing infections caused by multidrug-resistant A. baumannii [295,296,297]. Ultraviolet C light, blue light, and pimenta oil have also showed interesting activities in mice models of wound infections caused by A. baumannii [298,299,300].
Moreover, the combination of existing drugs is widely adopted for the treatment of injuries caused by MDR-AB. For instance, combined therapy using colistin and niclosamide has proven to be effective against strains that are already resistant to colistin [257]. Similarly, nisin was also shown to potentiate the action of polymyxin B against resistant A. baumannii [301]. Another alternative is the association of protegrin-1 with colistin, fosfomycin, levofloxacin, meropenem, tigecycline and rifampicin in the treatment of surgical wounds colonized by A. baumannii [302].
3.4.2. Main Genes Involved in Acinetobacter baumannii Virulence
Currently, the literature converges in affirming that A. baumannii has about 16 gene islands associated with virulence factors, thus directing a good part of its genome to pathogenic processes [303]. The main virulence factors associated with A. baumannii include systems of protein secretion, phospholipases, LPS, elements attached to the outer membrane, quorum sensing for biofilms and metal absorption [304].
Protein secretion systems are very effective in the virulence of A. baumannii. OmpA (encoded by the ompA gene) is one of the most studied proteins, as it is involved in the adhesion of epithelial cells and plays essential roles in the regulation of aggregation and biofilm formation in SSTIs [305,306]. Therefore, this protein represents a target for new antivirulence approaches against this pathogen [306].
OmpA is directly related to the mechanisms of cellular invasion and apoptosis, with an essential function in penetration of small solutes, and being classified as a single integral membrane protein anchored in the outer membrane [306,307]. The fixation and formation of biofilms by A. baumannii can occur in two ways: reversibly, where there is a strong physical–chemical attraction force, fundamental for the interaction between the strains and the contact surface; and irreversibly, as a result of the production of a matrix rich in exopolysaccharides, that is responsible for the permanent and coordinated adhesion of pathogens [308,309].
Phospholipases are other virulence factors widely described in A. baumannii. They are characterized as very important lipolytic enzymes for the cleavage of phospholipids that are present in cell membranes [310]. An example is phospholipase C that contributes to the cytolytic activity, allowing entry into epithelial cells [311,312]. Compounds coupled to the outer membrane of A. baumannii, such as the LPS antigenic O-polysaccharide, Csu pili (encoded by the csu gene) and biofilm-associated proteins (BapAb, encoded by the bap gene) can further promote adherence to skin epithelial cells as an initial stage of the colonization process [18,313].
A. baumannii is also known for having an external capsule with a high water-holding capacity, characterized by a dense polysaccharide that covers the entire surface of the bacterial cell and protects it against hostile environments, for example, dryness, disinfection and phagocytosis [314,315,316]. In addition, the synthesis of acinetobactin in a murine model of infection has been described, with an aggressive virulence factor of A. baumannii being noted in SSTIs [317]. A. baumannii also has sophisticated systems for metal acquisition. For example, in response to zinc (Zn), the pathogen can activate the expression of Zig A (a Zn-binding GTPase) encoded by zigA gene. The Zn uptake sytem is also composed by the ABC transporter and TonB, proteins presented in the inner and outer membranes, respectively) [318,319].
The genes associated in resistance and virulence in A. baumannii are represented in Table 4.
3.5. Pseudomonas aeruginosa
Undoubtedly, P. aeruginosa is one of the main pathogenic bacteria present in skin wounds. This microorganism belongs to the family Pseudomonadaceae, a Gram-negative bacterium that has the ability to develop in most natural and artificial environments [320]. This opportunistic pathogen is often isolated from samples of soil, water, plants and animals and can easily become resistant to antibiotics [321].
P. aeruginosa causes localized and systemic infections (e.g., ventilator-associated pneumonia, urinary tract infections or wound infections), especially in patients with severe burns, bet ulcers, and seriously ill and immunosuppressed subjects. Estimates indicate that this pathogen is involved in 10–15% of nosocomial infections, with a high prevalence of pulmonary complications in patients with cystic fibrosis [322,323]. P. aeruginosa is estimated to be present in at least one third of all skin infections worldwide, colonizing traumatic wounds, pressure and chronic ulcers and acantholytic or exudative dermatoses [324].
3.5.1. Main Genes Involved in Pseudomonas aeruginosa Resistance
P. aeruginosa has several resistance mechanisms, which can be classified as intrinsic (e.g., decreased permeability, expression of efflux systems and changes in the target; acquired, through gene transfer and mutations) and adaptive (transient in the presence or absence of stressors) [322,325,326].
In intrinsic and acquired forms, P. aeruginosa limits the entry of antibiotics into its cytoplasm by reducing the amount of non-specific porins in the membrane and replacing them with more specific ones for essential nutrients; for example, the mutation of the porin OprD (OprD gene), that reduces permeability to carbapenems [325] (Table 5). Even when some harmful substances can penetrate the bacterial cell, P. aeruginosa is able to activate its highly complex multi-drug efflux pump systems. The four best described are: MexAB-OprM, encoded by the mexAB-oprM genes; MexXY/OprM (OprA), by the expression of the mexXY-(oprA) genes; MexCD-OprJ, by the mexCD-oprJ genes; and MexEF-OprN, by mexEF-oprN [326].
These mutations are so frequent that in Europe, the report of European Centre for Disease Prevention and Control (ECDC), published in 2016, showed that 33.9% of P. aeruginosa isolates were resistant to at least one of the currently used antimicrobial groups [322]. Resistance to the most commonly used classes of drugs—such as fluoroquinolones—by strains of P. aeruginosa can also be observed through mutations of the targets of these antibiotics, more specifically, mutations in the gyrA and gyrB genes of DNA gyrase and the parC and parE genes of topoisomerase IV [327].
The resistance of P. aeruginosa to potent polymyxins has also been reported, occurring via chromosomal mutations [328,329]. However, recently acquired resistance in these strains was also detected by means of plasmids, through the conjugation of the genes mcr-1 and blaNDM-1, from E. coli and K. pneumoniae, respectively, both conferring resistance to colistin [330,331].
The adaptive mechanisms of resistance of P. aeruginosa have not yet been clarified. It is only known that this system depends on changes in defense gene expression in the presence of aggressive agents, and its withdrawal after a reduction in stress levels [332,333]. A shared characteristic for P. aeruginosa adaptive mutants is that they exhibit high levels of AmpC, due to the inactivation of ampD (ampC repressor) and other isolated ampR mutations, which assist in the coding of essential regulatory proteins in the induction of the ampC gene [326].
Thus, patients with burn infections caused by multidrug-resistant strains of P. aeruginosa, are generally affected by sepsis and suffer from high morbidity and mortality [334,335]. In a recent study, where 93 samples of P. aeruginosa collected from burn wound infections were isolated, 100% were resistant to one or more antimicrobials and 94.6% were multidrug-resistant [323].
Another major public health problem, resulting from infections by multidrug-resistant strains of P. aeruginosa, relates to the complications associated with diabetic patients [336,337]. In these patients, P. aeruginosa MDR has become an issue in the treatment of infections in diabetic foot ulcers (DFU) [338]. High rates of MBL-producing P. aeruginosa have been observed in many patients hospitalized with DFU, with this leading to lower limb amputation [337,338]. It has been described that the presence of the exoS and exoU genes is closely and directly related to the phenomena of antimicrobial resistance to multiple drugs and increased hospital stay length, making the individual more susceptible to pressure ulcers [339].
Some drug formulations have exhibited promising results towards multidrug-resistant P. aeruginosa strains in clinical trials. The associations between ceftazidime/avibactam and ceftolozane/tazobactam have shown excellent responses, including in phase III clinical studies [340,341,342,343,344]. The synergistic actions of these drugs with other drugs that are already used, such as meropenem, amikacin, aztreonam, colistin and fosfomycin, also demonstrated good results [345].
Additionally, the development of cefiderocol, a new siderophore Cephalosporin, represents a great hope for the treatment of injuries caused by MDR-PA [346,347]. The use of relebactam, imipenem and cilastatin, and some antibacterial peptides (such as ZY4) have also been demonstrated as alternatives in the fight against P. aeruginosa MDR [348,349]. Plant-derived compounds and probiotics have been suggested as emergent candidates for the treatment of P. aeruginosa-wound infections [350,351,352,353]. Finally, some studies have revealed the efficacy of some experimental vaccines for the prevention of skin infections by P. aeruginosa [354,355].
3.5.2. Main Genes Involved in Pseudomonas aeruginosa Virulence
Collectively, the virulence factors of P. aeruginosa ensure the process of invasion, tissue colonization and damage, and dissemination in the bloodstream [323]. The virulence factors associated with bacterial cells include the flagella, lipopolysaccharide, pili type III system—effector proteins that include ExoS (exoS), ExoT (exoT), ExoY (exoY) and ExoU (exoU)—and alginate. The extracellular determinants include hydrogen cyanide, metalloprotease zinc (LasB), alkaline protease, elastase (LASA), phospholipases (PLCH and PlcN), exotoxins and pyocyanin [323,339]. For the establishment of chronic infections, P. aeruginosa assumes a more aggressive behavior due adaptive mechanisms that involve the loss of fimbriae and flagella to isolate the host immune system and form biofilms. This state is associated with persistent inflammation, derived from the secretion of extracellular virulence factors [356].
P. aeruginosa possesses two different types of control systems that control the expression of the majority the virulence factors: the transcription regulatory system and two-component detection system-quorum. The two-component system (TCS) detects external signals by means of phosphotransferase, which activates specific transcriptional regulators, allowing cells to modulate gene expression in response to environmental conditions [320]. The expression of several virulence factors (including lipases, elastases, the skin and the production of many protease cytotoxins) is controlled by the mechanism of quorum sensing. This system has self-regulation dependent on the cell density. This mechanism favors the formation of aggressive and difficult to remove biofilms [320].
For example, it is known that elastase and alkaline protease (phzI, phzII, phzH, phzM, phzS, plcHa and plcN genes) deteriorate various components of the tissue—such as protein elements of connective tissue—and cleave the cell surface receptors of leukocytes, hindering the healing process of the skin [357]. It has also been observed that P. aeruginosa inhibits the degranulation of eosinophils that are present in the injured region, which ends up being an important inhibitory factor for the immune system, favoring constant tissue infection [358].
The expression of pili (pilA and pilB genes) participates in bacterial adhesion and the colonization of epithelial surfaces, as does the expression of the flagellum [357]. This induces an inflammatory response resulting in the production of IL-8, IL-6 and mucin [356,357]. Alginate plays a role in mediating mucin adhesion and promoting resistance to the defense mechanisms of the immune system, by inhibiting antibody binding and phagocytosis. The type III secretion system (TTS) injects various toxins directly into the cytosol of the host cells—with ExoU and ExoT known as being the most virulent [339,359]. Other extracellular virulence factors include phospholipase C, which destroys the host cell membrane, and exotoxin A (oxA gene), which contributes to both tissue damage in the early stages of infection, and to the uptake of important nutrients for its growth [357].
The Table 5 provides the examples of genes related for virulence and drug resistance in Pseudomonas aeruginosa.
3.6. Enterobacter spp.
Species from the genus Enterobacter are often associated with opportunistic skin infections in immunocompromised patients and demonstrate widespread resistance to antibiotics [22,360]. The most pathogenic species are usually referred to as Enterobacter cloacae complex (ECC), with the most commonly associated species being E. cloacae and E. hormaechei, in addition to E. aerogenes. Enterobacter is among the five most common Enterobacteriaceae involved in wound infections and SSTIs [361,362,363,364]. Some studies also point out the emergence of EEC clones with high epidemic potential [361,365,366]. Infection with E. cloacae or E. aerogenes results in mortality rates of up to 40% [361,367].
Despite its high prevalence, little is known about the virulence mechanisms of this genus of Enterobacteriaceae in SSTIs, but many mechanisms of antimicrobial resistance acquired by these microorganisms have already been reported [364] (Table 6).
Main Genes Involved in Enterobacter Resistance
Genetic analysis proved that ECC are producers of ESBL [220]. Several ECC strains have an MDR profile due the presence of enzymes that prevent the action of systemic and topical antibiotics that are used in the treatment of infected skin lesions, for example, TEM-1 β-lactamase [22,368,369]. The blaTEM-1 gene and its variants have high mutation rates which results in diversification of the enzymatic subtypes of resistance [22].
Alonzo et al. (2012) showed that 41.5% of the samples obtained by Enterobacter spp. were positive for the blaCTX-M resistance gene, which showed greater activity against the cephalosporins cefotaxime and ceftazidime [220]. Another study showed Enterobacter spp. as the third most common microorganism found in the evaluation of patients with mild to extreme severe burn injuries with signs of infection in the skin [370].
Other Enterobacter species have also been considered as being highly pathogenic. For instance, an MDR strain of E. asburiae was detected that expressed resistance genes to aminoglycosides, β-lactams, fluoroquinolones, fosfomycin, macrolides, phenicols, rifampicin and sulfonamides. The gene blaIMP-8 was located in the IncFIB plasmid, while blaCTX-M-3 and qnrS1 were both in the IncP1 plasmid. A non-typeable plasmid harbored blaCTX-M-14, blaTEM-1B, blaOXA-1, catB3 (phenicols resistance) and sul1 (sulfonamide resistance) [371]. Subsequently, E. cancerogenus was reported as a seriously aggressive infectious agent in skin wounds caused by mechanical trauma [372].
A larger survey involving 110 patients with skin ulcers infected by different microorganisms, showed that E. cloacae was present in approximately 7% of cases [373]. More specifically, all ten E. cloacae isolates obtained from a Turkish hospital were resistant to all available carbapenems; nine showed resistance to cefoperazone/sulbactam, trimethoprim and sulfamethoxazole, and 50–70% were resistant to other classes, such as aminoglycosides (gentamicin and amikacin) and fluoroquinolones (ciprofloxacin). The main resistance genes found in these samples were blaNDM (an unprecedented finding for this species), blaVIM and blaIMP [374]. Some strains of Enterobacter spp. also expressed blaKPC-2, blaKPC-3, blaKPC-4 and blaNDM-1 carbapenemic resistance genes [375].
Even with the high rate of emergence of resistant species in the Enterobacter genus, the clinical use of aztreonam has still shown good results in cases of severe infection by MDR clones, with no episodes of resistance reported to date [29]. The combination of colistin and imipenem drugs has also shown excellent results in in vivo models of infection [376]. Satisfactory results have also been achieved with the application of phage-based therapy (such as pyophages and multiple cocktails) in experimental models of infections induced by Enterobacter MDR strains [377,378]. The genes discussed in this section are summarized in Table 6.
4. Conclusions This work discusses the main immunological resources involved in the skin’s defense against pathogens and highlights the importance of ESKAPE bacteria as etiologic agents of cutaneous infections. The high incidence of antimicrobial resistance and hypervirulent profiles observed for ESKAPE pathogens are associated with the difficulties in the treatment of the infections provoked by them. In addition, these bacteria are prevalent in hospital settings where they can affect immunocompromised patients. Knowledge about the virulence and resistance markers of these species is important in order to develop new strategies to detect and treat their associated infections.
Enlarge this image.Figure 1. A schematic view of bacterial skin infection, derived from a loss in epidermis integrity. (1) An injury provokes a skin lesion that constitutes a gateway for microbial contamination. (2) Bacteria colonize the skin and produce a biofilm. (3) Bacteria secrete toxins that extend the tissue degradation, reaching dermis layer. (4) Resident immune cells recognize the bacteria and secrete immune mediators to neutralize the pathogens and recruit other immune cells. (5) Cell debris (damage-associated molecular patterns (DAMPS) and lipid mediators) activate immune cells and serve as chemoattractors. (6) Blood leukocytes are recruited to combat the pathogens. (7) Effector substances released by immune cells also promote tissue damage and amplify the inflammation.
| Genes | Product Function | Reference |
|---|---|---|
| Esp | Product is Enterococcus surface protein (Esp) which is responsible for epithelial cell adhesion and increased binding between the polysaccharide matrix and collagen binding proteins. | [132,150] |
| ace;efaAfm; cylA | Encode collagen binding adhesin and cytolysins that compromise the bonds between collagen fibers and the balance between keratinocytes and fibroblasts. | [155] |
| gelE; hyl | Responsible for the hydrolysis of collagen fibers and the cutaneous extracellular matrix. | [132,152,153,154] |
| Asa | Encodes aggregating substances, which facilitate the attachment to the skin epithelium and favor the bacterial aggregative behavior during plasmid conjugation. | [156] |
| vanA; vanB; vanC; vanD; vanE; vanG; vanL; vanM; vanN | Vancomycin resistance. | [133,134,135] |
| poxtA | Phenicols, tetracycline and linezolid resistance. | [122,123,124] |
| aac(6’)-Ie; aph(2’’); aph(3’)-IIIa; ant(4’)-Ia | Encode aminoglycoside modifying enzymes (AMEs) that confer resistance to drugs. | [140] |
| ere(B); erm(B) | Responsible for the production of esterase enzymes for erythromycin. | [142] |
| Genes | Product Function | Reference |
|---|---|---|
| etA; etB; etD | Encode the exfoliative toxins A, B and D that selectively bind and cleave a desmoglein-1 peptide bond. | [221] |
| lukED | Encodes leukocidin ED (LukED), a toxin related to blood and skin infections. | [220] |
| pvl | Encodes the Panton-Valentine leukocidin (PVL) which is associated to the destruction of resident immune cells and tissue necrosis. | [219,221] |
| blaZ | Involved in penicillin resistance, through the hydrolysis of its β-lactam ring. | [176] |
| mecA | Its product confers methicillin resistance, through a penicillin-binding protein. | [176] |
| vanA; vanH; vanX; vanS; vanR; vanY; vanZ; blaR1; blaIe; lmrS; vraR; mrgA; qacA; qacB; norA; mepA; mdeA; lmrS; mupA | These genes are involved in Multi-drug resistance—vancomycin, oxacillin, ciprofloxacin, norfloxacin, novobiocin, mupirocin, fusidic acid, trimethoprim and chloramphenicol. | [197,198,199,200] |
| Genes | Product Function | Reference |
|---|---|---|
| mrkABCDF | Encodes fimbriae type 1 and 3; binding to collagen. | [260,261] |
| Cps | Encodes polysaccharide capsule. | [262,263] |
| rmpA | Synthesis of capsular compounds. | [246] |
| magA, k2A; wcaG; wabG; uge; ycfM | Formation of capsule and its lipopolysaccharides (LPS). | [264] |
| wbbY; wbbZ | Modify LPS composition. | [267] |
| entS | Production of enterobactin. | [240,269] |
| armA; aacA4; aacC2; aadA1; aac(6’)-Ib | Aminoglycosides resistance. | [246,247] |
| blaKPC-2; blaKPC-3 | Carbapenem, clavulanic acid and tazobactam resistance. | [249,250,251] |
| acrAB, qnrB; qnrS | Quinolones resistance. | [246,254] |
| blaSHV; blaTEM; blaCTX-M | Carbapenems resistance. | [254] |
| lpxM | Polymyxin resistance. | [246] |
| ramR; rpsJ; tetA | Tigecycline resistance. | [255] |
| Genes | Product Function | Reference |
|---|---|---|
| ompA | Encodes OmpA protein, involved in the adhesion of epithelial cells and plays essential roles in the regulation of aggressiveness and biofilm formation. | [305,306] |
| csu; bap | Encodes Csu pili and biofilm-associated proteins that promote adherence to skin epithelial cells during initial stage of the colonization process. | [18,313] |
| zigA | Metal elimination system essential for its metabolism. | [318,319] |
| blactx-m; blages; blaper; blasco; blashv; blatem; blaveb | Penicillin and cephalosporin (except cephamycin) resistance. | [280,281,282,283] |
| katG | Hydrogen peroxide resistance. | [284] |
| tetA; tetB; tetM | Tetracyclines, minocycline and doxycycline resistance. | [18] |
| gyrA; parC | Fluoroquinolones resistance. | [286] |
| aac(3′)-Ia; ant(2’)-Ia; ant(3″); armA; rmtA; rmtB; rmtC; rmtD | Aminoglycosides resistance. | [18,288,289] |
| adeABC and adeM | Efflux pumps (gentamicin resistance) | [290] |
| pmrC; pmrA; prmB; lpsB; lptD; vacJ | Polymyxins resistance. | [291,292] |
| oxa-23; oxa-51 | Carbapenems resistance. | [293] |
| Genes | Product Function | Reference |
|---|---|---|
| exoS; exoT; exoY; exoU | Encode ExoS, ExoT, ExoY and ExoU proteins. | [323,339] |
| phzI; phzII; phzH; phzM; phzS; plcHa; plcN | Products are elastase and alkaline protease. | [357] |
| pilA; pilB | Expression of pili; participates in bacterial adhesion and the colonization of epithelial surfaces. | [357] |
| oxA | Exotoxin A; contributes to tissue damage in the early stages of infection, in addition to the uptake of important nutrients for its growth. | [357] |
| OprD | Carbapenems resistance. | [325] |
| mexAB-oprM; mexXY-(oprA); mexCD-oprJ; mexEF-oprN | Multi-drug resistance. | [326] |
| gyrA; gyrB; parC; parE | Fluoroquinolones resistance. | [327] |
| mcr-1; bl;M-1 | Polymyxins resistance. | [330,331] |
| exoS; exoU | Multi-drug resistance. | [339] |
| Genes | Product Function | References |
|---|---|---|
| blaTEM-1 | Multi-drug resistance. | [22] |
| blaCTX-M | Cephalosporins resistance. | [220] |
| blaIMP-8; blaCTX-M-3; qnrS1; blaCTX-M-14; blaTEM-1B; blaOXA-1; catB3; sul1 | Multi-drug resistance—aminoglycosides, β-lactams, fluoroquinolones, fosfomycin, macrolides, phenicols, rifampicin and sulfonamides. | [371] |
| blaNDM; blaVIM; blaIMP | Multi-drug resistance—carbapenems, cefoperazone, sulbactam, trimethoprim, sulfamethoxazole, aminoglycosides (gentamicin and amikacin) and fluoroquinolones (ciprofloxacin). | [374] |
| blaKPC-2, blaKPC-3, blaKPC-4 and blaNDM-1 | Carbapenems resistance. | [375] |
Author Contributions
G.H.R.V.d.M., E.R.O., G.V.D. and L.C.N.d.S. conceived the study and participated in its design and coordination. L.C.N.d.S., A.Z., V.L.C., L.d.S.S., J.M.N.B., R.d.C.M.d.M. were responsible for the sections about skin physiology and wound healing. G.H.R.V.d.M., E.R.O., G.V.D., G.D.E.C., J.S.P.M., A.d.S.M., A.S.d.S.A. were responsible for the sections about ESKAPE. G.H.R.V.d.M., E.R.O., A.d.S.M. and L.C.N.d.S. drafted the manuscript. All authors have read and agreed to the published version of the manuscript.
Funding
This work was funded by Fundação de Amparo à Pesquisa e Desenvolvimento Científico do Maranhão (Processes numbers: UNIVERSAL-01008/18, BEPP-02241/18, BIC-00682/19, BM-02341/19, BM-02666/19, BIC-02857/20) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (Process number: 426950/2018-6).
Institutional Review Board Statement
Not applicable.
Informed Consent Statement
Not applicable.
Data Availability Statement
Not applicable.
Conflicts of Interest
The authors declare no conflict of interest.
1. Guest, J.F.; Ayoub, N.; McIlwraith, T.; Uchegbu, I.; Gerrish, A.; Weidlich, D.; Vowden, K.; Vowden, P. Health economic burden that different wound types impose on the UK's National Health Service. Int. Wound J. 2017, 14, 322-330.
2. Mitchell, R.J.; Curtis, K.; Braithwaite, J. Health outcomes and costs for injured young people hospitalised with and without chronic health conditions. Injury 2017, 48, 1776-1783.
3. Jacquet, R.; LaBauve, A.E.; Akoolo, L.; Patel, S.; Alqarzaee, A.A.; Wong Fok Lung, T.; Poorey, K.; Stinear, T.P.; Thomas, V.C.; Meagher, R.J.; et al. Dual Gene Expression Analysis Identifies Factors Associated with Staphylococcus aureus Virulence in Diabetic Mice. Infect. Immun. 2019, 87, e00163-19.
4. Geisinger, E.; Isberg, R.R. Interplay Between Antibiotic Resistance and Virulence During Disease Promoted by Multidrug-Resistant Bacteria. J. Infect. Dis. 2017, 215, S9-S17.
5. Buch, P.J.; Chai, Y.; Goluch, E.D. Treating Polymicrobial Infections in Chronic Diabetic Wounds. Clin. Microbiol. Rev. 2019, 32.
6. Hsu, J.T.; Chen, Y.W.; Ho, T.W.; Tai, H.C.; Wu, J.M.; Sun, H.Y.; Hung, C.S.; Zeng, Y.C.; Kuo, S.Y.; Lai, F. Chronic wound assessment and infection detection method. BMC Med. Inform. Decis. Mak. 2019, 19, 99.
7. Ziwa, M.; Jovic, G.; Ngwisha, C.L.T.; Molnar, J.A.; Kwenda, G.; Samutela, M.; Mulowa, M.; Kalumbi, M.M. Common hydrotherapy practices and the prevalence of burn wound bacterial colonisation at the University Teaching Hospital in Lusaka, Zambia. Burns 2019, 45, 983-989.
8. Carvalho, A.R., Jr.; Diniz, R.M.; Suarez, M.A.M.; Figueiredo, C.; Zagmignan, A.; Grisotto, M.A.G.; Fernandes, E.S.; da Silva, L.C.N. Use of Some Asteraceae Plants for the Treatment of Wounds: From Ethnopharmacological Studies to Scientific Evidences. Front. Pharmacol. 2018, 9, 784.
9. Rodrigues, M.; Kosaric, N.; Bonham, C.A.; Gurtner, G.C. Wound Healing: A Cellular Perspective. Physiol. Rev. 2019, 99, 665-706.
10. Wang, P.H.; Huang, B.S.; Horng, H.C.; Yeh, C.C.; Chen, Y.J. Wound healing. J. Chin. Med. Assoc. 2018, 81, 94-101.
11. Zomer, H.D.; Trentin, A.G. Skin wound healing in humans and mice: Challenges in translational research. J. Dermatol. Sci. 2018, 90, 3-12.
12. Rahim, K.; Saleha, S.; Zhu, X.; Huo, L.; Basit, A.; Franco, O.L. Bacterial Contribution in Chronicity of Wounds. Microb. Ecol. 2017, 73, 710-721.
13. Cooper, R.A.; Bjarnsholt, T.; Alhede, M. Biofilms in wounds: A review of present knowledge. J. Wound Care 2014, 23, 570-582.
14. Vestby, L.K.; Gronseth, T.; Simm, R.; Nesse, L.L. Bacterial Biofilm and its Role in the Pathogenesis of Disease. Antibiotics 2020, 9, 59.
15. Bhattacharya, M.; Berends, E.T.M.; Chan, R.; Schwab, E.; Roy, S.; Sen, C.K.; Torres, V.J.; Wozniak, D.J. Staphylococcus aureus biofilms release leukocidins to elicit extracellular trap formation and evade neutrophil-mediated killing. Proc. Natl. Acad. Sci. USA 2018, 115, 7416-7421.
16. Garcia-Perez, A.N.; de Jong, A.; Junker, S.; Becher, D.; Chlebowicz, M.A.; Duipmans, J.C.; Jonkman, M.F.; van Dijl, J.M. From the wound to the bench: Exoproteome interplay between wound-colonizing Staphylococcus aureus strains and co-existing bacteria. Virulence 2018, 9, 363-378.
17. Srivastava, P.; Sivashanmugam, K. Combinatorial Drug Therapy for Controlling Pseudomonas aeruginosa and Its Association with Chronic Condition of Diabetic Foot Ulcer. Int. J. Low Extrem. Wounds 2020, 19, 7-20.
18. Ayoub Moubareck, C.; Hammoudi Halat, D. Insights into Acinetobacter baumannii: A Review of Microbiological, Virulence, and Resistance Traits in a Threatening Nosocomial Pathogen. Antibiotics 2020, 9, 119.
19. Wu, Y.K.; Cheng, N.C.; Cheng, C.M. Biofilms in Chronic Wounds: Pathogenesis and Diagnosis. Trends Biotechnol. 2019, 37, 505-517.
20. Kadam, S.; Shai, S.; Shahane, A.; Kaushik, K.S. Recent Advances in Non-Conventional Antimicrobial Approaches for Chronic Wound Biofilms: Have We Found the 'Chink in the Armor'? Biomedicines 2019, 7, 35.
21. Morgan, S.J.; Lippman, S.I.; Bautista, G.E.; Harrison, J.J.; Harding, C.L.; Gallagher, L.A.; Cheng, A.C.; Siehnel, R.; Ravishankar, S.; Usui, M.L.; et al. Bacterial fitness in chronic wounds appears to be mediated by the capacity for high-density growth, not virulence or biofilm functions. PLoS Pathog. 2019, 15, e1007511.
22. Santajit, S.; Indrawattana, N. Mechanisms of Antimicrobial Resistance in ESKAPE Pathogens. Biomed. Res. Int. 2016, 2016, 2475067.
23. Serra, R.; Grande, R.; Butrico, L.; Rossi, A.; Settimio, U.F.; Caroleo, B.; Amato, B.; Gallelli, L.; de Franciscis, S. Chronic wound infections: The role of Pseudomonas aeruginosa and Staphylococcus aureus. Expert Rev. Anti Infect. Ther. 2015, 13, 605-613.
24. Heitkamp, R.A.; Li, P.; Mende, K.; Demons, S.T.; Tribble, D.R.; Tyner, S.D. Association of Enterococcus spp. with Severe Combat Extremity Injury, Intensive Care, and Polymicrobial Wound Infection. Surg Infect. 2018, 19, 95-103.
25. Trookman, N.S.; Rizer, R.L.; Weber, T. Treatment of minor wounds from dermatologic procedures: A comparison of three topical wound care ointments using a laser wound model. J. Am. Acad. Dermatol. 2011, 64, S8-S15.
26. Punjataewakupt, A.; Napavichayanun, S.; Aramwit, P. The downside of antimicrobial agents for wound healing. Eur. J. Clin. Microbiol. Infect. Dis. 2019, 38, 39-54.
27. Rahimi, M.; Noruzi, E.B.; Sheykhsaran, E.; Ebadi, B.; Kariminezhad, Z.; Molaparast, M.; Mehrabani, M.G.; Mehramouz, B.; Yousefi, M.; Ahmadi, R.; et al. Carbohydrate polymer-based silver nanocomposites: Recent progress in the antimicrobial wound dressings. Carbohydr. Polym. 2020, 231, 115696.
28. Lupo, A.; Haenni, M.; Madec, J.Y. Antimicrobial Resistance in Acinetobacter spp. and Pseudomonas spp. Microbiol. Spectr. 2018, 6, 377-393.
29. Mulani, M.S.; Kamble, E.E.; Kumkar, S.N.; Tawre, M.S.; Pardesi, K.R. Emerging Strategies to Combat ESKAPE Pathogens in the Era of Antimicrobial Resistance: A Review. Front. Microbiol. 2019, 10, 539.
30. Mendoza, N.; Tyring, S.K. Emerging drugs for complicated skin and skin-structure infections. Expert Opin. Emerg. Drugs 2010, 15, 509-520.
31. McNeil, J.C.; Hulten, K.G.; Kaplan, S.L.; Mason, E.O. Mupirocin resistance in Staphylococcus aureus causing recurrent skin and soft tissue infections in children. Antimicrob. Agents Chemother. 2011, 55, 2431-2433.
32. Singer, H.M.; Levin, L.E.; Garzon, M.C.; Lauren, C.T.; Planet, P.J.; Kittler, N.W.; Whittier, S.; Morel, K.D. Wound culture isolated antibiograms and caregiver-reported skin care practices in children with epidermolysis bullosa. Pediatr. Dermatol. 2018, 35, 92-96.
33. Hosny, A.E.M.; Rasmy, S.A.; Aboul-Magd, D.S.; Kashef, M.T.; El-Bazza, Z.E. The increasing threat of silver-resistance in clinical isolates from wounds and burns. Infect. Drug Resist. 2019, 12, 1985-2001.
34. Andrade, L.N.; Siqueira, T.E.S.; Martinez, R.; Darini, A.L.C. Multidrug-Resistant CTX-M-(15, 9, 2)- and KPC-2-Producing Enterobacter hormaechei and Enterobacter asburiae Isolates Possessed a Set of Acquired Heavy Metal Tolerance Genes Including a Chromosomal sil Operon (for Acquired Silver Resistance). Front. Microbiol. 2018, 9, 539.
35. Visscher, M.O.; Adam, R.; Brink, S.; Odio, M. Newborn infant skin: Physiology, development, and care. Clin. Dermatol. 2015, 33, 271-280.
36. Fore, J. A review of skin and the effects of aging on skin structure and function. Ostomy Wound Manag. 2006, 52, 24-35, quiz 36-27.
37. Jia, Y.; Gan, Y.; He, C.; Chen, Z.; Zhou, C. The mechanism of skin lipids influencing skin status. J. Dermatol. Sci. 2018, 89, 112-119.
38. Hsu, Y.C.; Li, L.; Fuchs, E. Emerging interactions between skin stem cells and their niches. Nat. Med. 2014, 20, 847-856.
39. Ali, N.; Rosenblum, M.D. Regulatory T cells in skin. Immunology 2017, 152, 372-381.
40. Malissen, B.; Tamoutounour, S.; Henri, S. The origins and functions of dendritic cells and macrophages in the skin. Nat. Rev. Immunol. 2014, 14, 417-428.
41. Richmond, J.M.; Harris, J.E. Immunology and skin in health and disease. Cold Spring Harb. Perspect. Med. 2014, 4, a015339.
42. Kabashima, K.; Honda, T.; Ginhoux, F.; Egawa, G. The immunological anatomy of the skin. Nat. Rev. Immunol. 2019, 19, 19-30.
43. Gong, T.; Liu, L.; Jiang, W.; Zhou, R. DAMP-sensing receptors in sterile inflammation and inflammatory diseases. Nat. Rev. Immunol. 2020, 20, 95-112.
44. Pandolfi, F.; Altamura, S.; Frosali, S.; Conti, P. Key Role of DAMP in Inflammation, Cancer, and Tissue Repair. Clin. Ther. 2016, 38, 1017-1028.
45. Fischer, S. Pattern Recognition Receptors and Control of Innate Immunity: Role of Nucleic Acids. Curr. Pharm. Biotechnol. 2018, 19, 1203-1209.
46. Westman, J.; Grinstein, S.; Marques, P.E. Phagocytosis of Necrotic Debris at Sites of Injury and Inflammation. Front. Immunol. 2019, 10, 3030.
47. Wang, J. Neutrophils in tissue injury and repair. Cell Tissue Res. 2018, 371, 531-539.
48. Portou, M.J.; Baker, D.; Abraham, D.; Tsui, J. The innate immune system, toll-like receptors and dermal wound healing: A review. Vascul. Pharmacol. 2015, 71, 31-36.
49. Wolf, A.J.; Underhill, D.M. Peptidoglycan recognition by the innate immune system. Nat. Rev. Immunol. 2018, 18, 243-254.
50. Chen, L.; DiPietro, L.A. Toll-Like Receptor Function in Acute Wounds. Adv. Wound Care 2017, 6, 344-355.
51. Egert, M.; Simmering, R.; Riedel, C.U. The Association of the Skin Microbiota with Health, Immunity, and Disease. Clin. Pharmacol. Ther. 2017, 102, 62-69.
52. Marongiu, L.; Gornati, L.; Artuso, I.; Zanoni, I.; Granucci, F. Below the surface: The inner lives of TLR4 and TLR9. J. Leukoc. Biol. 2019, 106, 147-160.
53. Quaresma, J.A.S. Organization of the Skin Immune System and Compartmentalized Immune Responses in Infectious Diseases. Clin. Microbiol. Rev. 2019, 32.
54. Ono, S.; Kabashima, K. Novel insights into the role of immune cells in skin and inducible skin-associated lymphoid tissue (iSALT). Allergo J. Int. 2015, 24, 170-179.
55. Kogame, T.; Yamashita, R.; Hirata, M.; Kataoka, T.R.; Kamido, H.; Ueshima, C.; Matsui, M.; Nomura, T.; Kabashima, K. Analysis of possible structures of inducible skin-associated lymphoid tissue in lupus erythematosus profundus. J. Dermatol. 2018, 45, 1117-1121.
56. Ono, S.; Kabashima, K. Proposal of inducible skin-associated lymphoid tissue (iSALT). Exp. Dermatol. 2015, 24, 630-631.
57. Honda, T.; Egawa, G.; Kabashima, K. Antigen presentation and adaptive immune responses in skin. Int. Immunol. 2019, 31, 423-429.
58. Chen, Y.E.; Fischbach, M.A.; Belkaid, Y. Skin microbiota-host interactions. Nature 2018, 553, 427-436.
59. Ruff, W.E.; Greiling, T.M.; Kriegel, M.A. Host-microbiota interactions in immune-mediated diseases. Nat. Rev. Microbiol. 2020, 18, 521-538.
60. Belkaid, Y.; Segre, J.A. Dialogue between skin microbiota and immunity. Science 2014, 346, 954-959.
61. Byrd, A.L.; Belkaid, Y.; Segre, J.A. The human skin microbiome. Nat. Rev. Microbiol. 2018, 16, 143-155.
62. Nakamizo, S.; Egawa, G.; Honda, T.; Nakajima, S.; Belkaid, Y.; Kabashima, K. Commensal bacteria and cutaneous immunity. Semin. Immunopathol. 2015, 37, 73-80.
63. Yousef, H.; Alhajj, M.; Sharma, S. Anatomy, Skin (Integument), Epidermis. In StatPearls; StatPearls Publishing: Treasure Island, FL, USA, 2019.
64. Grone, A. Keratinocytes and cytokines. Vet. Immunol. Immunopathol. 2002, 88, 1-12.
65. Asahina, R.; Maeda, S. A review of the roles of keratinocyte-derived cytokines and chemokines in the pathogenesis of atopic dermatitis in humans and dogs. Vet. Dermatol. 2017, 28, 16-e15.
66. Banerjee, G.; Damodaran, A.; Devi, N.; Dharmalingam, K.; Raman, G. Role of keratinocytes in antigen presentation and polarization of human T lymphocytes. Scand. J. Immunol. 2004, 59, 385-394.
67. Deckers, J.; Hammad, H.; Hoste, E. Langerhans Cells: Sensing the Environment in Health and Disease. Front. Immunol. 2018, 9, 93.
68. Doebel, T.; Voisin, B.; Nagao, K. Langerhans Cells-The Macrophage in Dendritic Cell Clothing. Trends Immunol. 2017, 38, 817-828.
69. Otsuka, M.; Egawa, G.; Kabashima, K. Uncovering the Mysteries of Langerhans Cells, Inflammatory Dendritic Epidermal Cells, and Monocyte-Derived Langerhans Cell-Like Cells in the Epidermis. Front. Immunol. 2018, 9, 1768.
70. Toulon, A.; Breton, L.; Taylor, K.R.; Tenenhaus, M.; Bhavsar, D.; Lanigan, C.; Rudolph, R.; Jameson, J.; Havran, W.L. A role for human skin-resident T cells in wound healing. J. Exp. Med. 2009, 206, 743-750.
71. Dijkgraaf, F.E.; Matos, T.R.; Hoogenboezem, M.; Toebes, M.; Vredevoogd, D.W.; Mertz, M.; van den Broek, B.; Song, J.Y.; Teunissen, M.B.M.; Luiten, R.M.; et al. Tissue patrol by resident memory CD8(+) T cells in human skin. Nat. Immunol. 2019, 20, 756-764.
72. Li, Y.H.; Liu, Y.; Huang, L.; Xu, Y.F.; Zhu, H.; Li, T.; Deng, W.; Qin, C. Dynamic Changes of the Quantitative Distribution, Apoptosis and Proliferation of T and B Cells in the Skin of KM Mutant Mice. Zhongguo Yi Xue Ke Xue Yuan Xue Bao 2015, 37, 489-495.
73. Debes, G.F.; McGettigan, S.E. Skin-Associated B Cells in Health and Inflammation. J. Immunol. 2019, 202, 1659-1666.
74. Lafouresse, F.; Groom, J.R. A Task Force Against Local Inflammation and Cancer: Lymphocyte Trafficking to and Within the Skin. Front. Immunol. 2018, 9, 2454.
75. Hesketh, M.; Sahin, K.B.; West, Z.E.; Murray, R.Z. Macrophage Phenotypes Regulate Scar Formation and Chronic Wound Healing. Int. J. Mol. Sci. 2017, 18, 1545.
76. Krzyszczyk, P.; Schloss, R.; Palmer, A.; Berthiaume, F. The Role of Macrophages in Acute and Chronic Wound Healing and Interventions to Promote Pro-wound Healing Phenotypes. Front. Physiol. 2018, 9, 419.
77. Kim, S.Y.; Nair, M.G. Macrophages in wound healing: Activation and plasticity. Immunol. Cell Biol. 2019, 97, 258-267.
78. Shapouri-Moghaddam, A.; Mohammadian, S.; Vazini, H.; Taghadosi, M.; Esmaeili, S.A.; Mardani, F.; Seifi, B.; Mohammadi, A.; Afshari, J.T.; Sahebkar, A. Macrophage plasticity, polarization, and function in health and disease. J. Cell Physiol. 2018, 233, 6425-6440.
79. Vergadi, E.; Ieronymaki, E.; Lyroni, K.; Vaporidi, K.; Tsatsanis, C. Akt Signaling Pathway in Macrophage Activation and M1/M2 Polarization. J. Immunol. 2017, 198, 1006-1014.
80. Orecchioni, M.; Ghosheh, Y.; Pramod, A.B.; Ley, K. Macrophage Polarization: Different Gene Signatures in M1(LPS+) vs. Classically and M2(LPS-) vs. Alternatively Activated Macrophages. Front. Immunol. 2019, 10, 1084.
81. Das, A.; Sinha, M.; Datta, S.; Abas, M.; Chaffee, S.; Sen, C.K.; Roy, S. Monocyte and macrophage plasticity in tissue repair and regeneration. Am. J. Pathol. 2015, 185, 2596-2606.
82. Bouchery, T.; Harris, N. Neutrophil-macrophage cooperation and its impact on tissue repair. Immunol. Cell Biol. 2019, 97, 289-298.
83. De Oliveira, S.; Rosowski, E.E.; Huttenlocher, A. Neutrophil migration in infection and wound repair: Going forward in reverse. Nat. Rev. Immunol. 2016, 16, 378-391.
84. De Sousa, J.R.; Lucena Neto, F.D.; Sotto, M.N.; Quaresma, J.A.S. Immunohistochemical characterization of the M4 macrophage population in leprosy skin lesions. BMC Infect. Dis. 2018, 18, 576.
85. Kim, N.D.; Luster, A.D. The role of tissue resident cells in neutrophil recruitment. Trends Immunol. 2015, 36, 547-555.
86. Rohani, M.G.; Parks, W.C. Matrix remodeling by MMPs during wound repair. Matrix Biol. 2015, 44-46, 113-121.
87. Krishnaswamy, V.R.; Mintz, D.; Sagi, I. Matrix metalloproteinases: The sculptors of chronic cutaneous wounds. Biochim Biophys Acta Mol. Cell Res. 2017, 1864, 2220-2227.
88. Lazaro, J.L.; Izzo, V.; Meaume, S.; Davies, A.H.; Lobmann, R.; Uccioli, L. Elevated levels of matrix metalloproteinases and chronic wound healing: An updated review of clinical evidence. J. Wound Care 2016, 25, 277-287.
89. Mortaz, E.; Alipoor, S.D.; Adcock, I.M.; Mumby, S.; Koenderman, L. Update on Neutrophil Function in Severe Inflammation. Front. Immunol. 2018, 9, 2171.
90. Kovtun, A.; Messerer, D.A.C.; Scharffetter-Kochanek, K.; Huber-Lang, M.; Ignatius, A. Neutrophils in Tissue Trauma of the Skin, Bone, and Lung: Two Sides of the Same Coin. J. Immunol. Res. 2018, 2018, 8173983.
91. Brazil, J.C.; Quiros, M.; Nusrat, A.; Parkos, C.A. Innate immune cell-epithelial crosstalk during wound repair. J. Clin. Investig. 2019, 129, 2983-2993.
92. Geherin, S.A.; Fintushel, S.R.; Lee, M.H.; Wilson, R.P.; Patel, R.T.; Alt, C.; Young, A.J.; Hay, J.B.; Debes, G.F. The skin, a novel niche for recirculating B cells. J. Immunol. 2012, 188, 6027-6035.
93. Mauri, C.; Bosma, A. Immune regulatory function of B cells. Annu. Rev. Immunol. 2012, 30, 221-241.
94. Fillatreau, S. Regulatory roles of B cells in infectious diseases. Clin. Exp. Rheumatol. 2016, 34, 1-5.
95. Dai, Y.C.; Zhong, J.; Xu, J.F. Regulatory B cells in infectious disease (Review). Mol. Med. Rep. 2017, 16, 3-10.
96. Woodley, D.T. Distinct Fibroblasts in the Papillary and Reticular Dermis: Implications for Wound Healing. Dermatol. Clin. 2017, 35, 95-100.
97. Van Linthout, S.; Miteva, K.; Tschope, C. Crosstalk between fibroblasts and inflammatory cells. Cardiovasc. Res. 2014, 102, 258-269.
98. Kuhbacher, A.; Henkel, H.; Stevens, P.; Grumaz, C.; Finkelmeier, D.; Burger-Kentischer, A.; Sohn, K.; Rupp, S. Central Role for Dermal Fibroblasts in Skin Model Protection against Candida albicans. J. Infect. Dis 2017, 215, 1742-1752.
99. Fallahi, P.; Foddis, R.; Elia, G.; Ragusa, F.; Patrizio, A.; Benvenga, S.; Cristaudo, A.; Antonelli, A.; Ferrari, S.M. CXCL8 and CXCL11 chemokine secretion in dermal fibroblasts is differentially modulated by vanadium pentoxide. Mol. Med. Rep. 2018, 18, 1798-1803.
100. Gillitzer, R.; Goebeler, M. Chemokines in cutaneous wound healing. J. Leukoc. Biol. 2001, 69, 513-521.
101. Rees, P.A.; Greaves, N.S.; Baguneid, M.; Bayat, A. Chemokines in Wound Healing and as Potential Therapeutic Targets for Reducing Cutaneous Scarring. Adv. Wound Care 2015, 4, 687-703.
102. Takagi, H.; Arimura, K.; Uto, T.; Fukaya, T.; Nakamura, T.; Choijookhuu, N.; Hishikawa, Y.; Sato, K. Plasmacytoid dendritic cells orchestrate TLR7-mediated innate and adaptive immunity for the initiation of autoimmune inflammation. Sci. Rep. 2016, 6, 24477.
103. Furue, M.; Furue, K.; Tsuji, G.; Nakahara, T. Interleukin-17A and Keratinocytes in Psoriasis. Int. J. Mol. Sci. 2020, 21, 1275.
104. Feuerstein, R.; Kolter, J.; Henneke, P. Dynamic interactions between dermal macrophages and Staphylococcus aureus. J. Leukoc. Biol. 2017, 101, 99-106.
105. Sharma, A.; Rudra, D. Emerging Functions of Regulatory T Cells in Tissue Homeostasis. Front. Immunol. 2018, 9, 883.
106. Seneschal, J.; Clark, R.A.; Gehad, A.; Baecher-Allan, C.M.; Kupper, T.S. Human epidermal Langerhans cells maintain immune homeostasis in skin by activating skin resident regulatory T cells. Immunity 2012, 36, 873-884.
107. Rajilic-Stojanovic, M.; de Vos, W.M. The first 1000 cultured species of the human gastrointestinal microbiota. FEMS Microbiol. Rev. 2014, 38, 996-1047.
108. Ulrich, N.; Vonberg, R.P.; Gastmeier, P. Outbreaks caused by vancomycin-resistant Enterococcus faecium in hematology and oncology departments: A systematic review. Heliyon 2017, 3, e00473.
109. Whiteside, S.A.; Dave, S.; Seney, S.L.; Wang, P.; Reid, G.; Burton, J.P. Enterococcus faecalis persistence in pediatric patients treated with antibiotic prophylaxis for recurrent urinary tract infections. Future Microbiol. 2018, 13, 1095-1115.
110. Monticelli, J.; Knezevich, A.; Luzzati, R.; Di Bella, S. Clinical management of non-faecium non-faecalis vancomycin-resistant enterococci infection. Focus on Enterococcus gallinarum and Enterococcus casseliflavus/flavescens. J. Infect. Chemother. 2018, 24, 237-246.
111. Rajkumari, N.; Mathur, P.; Misra, M.C. Soft Tissue and Wound Infections Due to Enterococcus spp. Among Hospitalized Trauma Patients in a Developing Country. J. Glob. Infect. Dis. 2014, 6, 189-193.
112. Salem-Bekhit, M.M.; Moussa, I.M.; Muharram, M.M.; Alanazy, F.K.; Hefni, H.M. Prevalence and antimicrobial resistance pattern of multidrug-resistant enterococci isolated from clinical specimens. Indian J. Med. Microbiol. 2012, 30, 44-51.
113. Dworniczek, E.; Piwowarczyk, J.; Bania, J.; Kowalska-Krochmal, B.; Walecka, E.; Seniuk, A.; Dolna, I.; Gosciniak, G. Enterococcus in wound infections: Virulence and antimicrobial resistance. Acta Microbiol. Immunol. Hung. 2012, 59, 263-269.
114. Pendleton, J.N.; Gorman, S.P.; Gilmore, B.F. Clinical relevance of the ESKAPE pathogens. Expert Rev. Anti Infect. Ther. 2013, 11, 297-308.
115. Pochhammer, J.; Kramer, A.; Schaffer, M. Enterococci and surgical site infections: Causal agent or harmless commensals? Chirurg 2017, 88, 377-384.
116. Hinojosa, C.A.; Boyer-Duck, E.; Anaya-Ayala, J.E.; Nunez-Salgado, A.; Laparra-Escareno, H.; Torres-Machorro, A.; Lizola, R. Impact of the bacteriology of diabetic foot ulcers in limb loss. Wound Repair Regen. 2016, 24, 923-927.
117. Weintrob, A.C.; Murray, C.K.; Xu, J.; Krauss, M.; Bradley, W.; Warkentien, T.E.; Lloyd, B.A.; Tribble, D.R. Early Infections Complicating the Care of Combat Casualties from Iraq and Afghanistan. Surg. Infect. 2018, 19, 286-297.
118. Arias, M.; Hassan-Reshat, S.; Newsholme, W. Retrospective analysis of diabetic foot osteomyelitis management and outcome at a tertiary care hospital in the UK. PLoS ONE 2019, 14, e0216701.
119. Elhani, D.; Klibi, N.; Dziri, R.; Ben Hassan, M.; Asli Mohamed, S.; Ben Said, L.; Mahjoub, A.; Ben Slama, K.; Jemli, B.; Bellaj, R.; et al. vanA-containing E. faecium isolates of clonal complex CC17 in clinical and environmental samples in a Tunisian hospital. Diagn. Microbiol. Infect. Dis. 2014, 79, 60-63.
120. Huang, J.; Wang, M.; Gao, Y.; Chen, L.; Wang, L. Emergence of plasmid-mediated oxazolidinone resistance gene poxtA from CC17 Enterococcus faecium of pig origin. J. Antimicrob. Chemother. 2019, 74, 2524-2530.
121. Lee, T.; Pang, S.; Abraham, S.; Coombs, G.W. Antimicrobial-resistant CC17 Enterococcus faecium: The past, the present and the future. J. Glob. Antimicrob. Resist. 2019, 16, 36-47.
122. Sadowy, E. Linezolid resistance genes and genetic elements enhancing their dissemination in enterococci and streptococci. Plasmid 2018, 99, 89-98.
123. Hasman, H.; Clausen, P.; Kaya, H.; Hansen, F.; Knudsen, J.D.; Wang, M.; Holzknecht, B.J.; Samulioniene, J.; Roder, B.L.; Frimodt-Moller, N.; et al. LRE-Finder, a Web tool for detection of the 23S rRNA mutations and the optrA, cfr, cfr(B) and poxtA genes encoding linezolid resistance in enterococci from whole-genome sequences. J. Antimicrob. Chemother. 2019, 74, 1473-1476.
124. Bender, J.K.; Fleige, C.; Klare, I.; Werner, G. Development of a multiplex-PCR to simultaneously detect acquired linezolid resistance genes cfr, optrA and poxtA in enterococci of clinical origin. J. Microbiol. Methods 2019, 160, 101-103.
125. Gao, W.; Howden, B.P.; Stinear, T.P. Evolution of virulence in Enterococcus faecium, a hospital-adapted opportunistic pathogen. Curr. Opin. Microbiol. 2018, 41, 76-82.
126. Willems, R.J.; Top, J.; van Schaik, W.; Leavis, H.; Bonten, M.; Siren, J.; Hanage, W.P.; Corander, J. Restricted gene flow among hospital subpopulations of Enterococcus faecium. MBio 2012, 3, e00151-12.
127. Arias, C.A.; Murray, B.E. The rise of the Enterococcus: Beyond vancomycin resistance. Nat. Rev. Microbiol. 2012, 10, 266-278.
128. Garcia-Solache, M.; Rice, L.B. The Enterococcus: A Model of Adaptability to Its Environment. Clin. Microbiol. Rev. 2019, 32.
129. Wang, Y.; Lv, Y.; Cai, J.; Schwarz, S.; Cui, L.; Hu, Z.; Zhang, R.; Li, J.; Zhao, Q.; He, T.; et al. A novel gene, optrA, that confers transferable resistance to oxazolidinones and phenicols and its presence in Enterococcus faecalis and Enterococcus faecium of human and animal origin. J. Antimicrob. Chemother. 2015, 70, 2182-2190.
130. Golob, M.; Pate, M.; Kusar, D.; Dermota, U.; Avbersek, J.; Papic, B.; Zdovc, I. Antimicrobial Resistance and Virulence Genes in Enterococcus faecium and Enterococcus faecalis from Humans and Retail Red Meat. Biomed. Res. Int. 2019, 2019, 2815279.
131. Cassini, A.; Hogberg, L.D.; Plachouras, D.; Quattrocchi, A.; Hoxha, A.; Simonsen, G.S.; Colomb-Cotinat, M.; Kretzschmar, M.E.; Devleesschauwer, B.; Cecchini, M.; et al. Attributable deaths and disability-adjusted life-years caused by infections with antibiotic-resistant bacteria in the EU and the European Economic Area in 2015: A population-level modelling analysis. Lancet Infect. Dis. 2019, 19, 56-66.
132. Papadimitriou-Olivgeris, M.; Filippidou, S.; Drougka, E.; Fligou, F.; Kolonitsiou, F.; Dodou, V.; Marangos, M.; Anastassiou, E.D.; Vantarakis, A.; Spiliopoulou, I. Biofilm synthesis and presence of virulence factors among enterococci isolated from patients and water samples. J. Med. Microbiol. 2015, 64, 1270-1276.
133. Ahmed, M.O.; Baptiste, K.E. Vancomycin-Resistant Enterococci: A Review of Antimicrobial Resistance Mechanisms and Perspectives of Human and Animal Health. Microb. Drug Resist. 2018, 24, 590-606.
134. Lebreton, F.; Valentino, M.D.; Schaufler, K.; Earl, A.M.; Cattoir, V.; Gilmore, M.S. Transferable vancomycin resistance in clade B commensal-type Enterococcus faecium. J. Antimicrob. Chemother. 2018, 73, 1479-1486.
135. Teo, J.W.; Krishnan, P.; Jureen, R.; Lin, R.T. Detection of an unusual van genotype in a vancomycin-resistant Enterococcus faecium hospital isolate. J. Clin. Microbiol. 2011, 49, 4297-4298.
136. Evers, S.; Quintiliani, R., Jr.; Courvalin, P. Genetics of glycopeptide resistance in enterococci. Microb. Drug Resist. 1996, 2, 219-223.
137. Papagiannitsis, C.C.; Malli, E.; Florou, Z.; Medvecky, M.; Sarrou, S.; Hrabak, J.; Petinaki, E. First description in Europe of the emergence of Enterococcus faecium ST117 carrying both vanA and vanB genes, isolated in Greece. J. Glob. Antimicrob. Resist. 2017, 11, 68-70.
138. Sharifi, Y.; Hasani, A.; Ghotaslou, R.; Varshochi, M.; Hasani, A.; Aghazadeh, M.; Milani, M. Survey of Virulence Determinants among Vancomycin Resistant Enterococcus faecalis and Enterococcus faecium Isolated from Clinical Specimens of Hospitalized Patients of North west of Iran. Open Microbiol. J. 2012, 6, 34-39.
139. Wardal, E.; Kuch, A.; Gawryszewska, I.; Zabicka, D.; Hryniewicz, W.; Sadowy, E. Diversity of plasmids and Tn1546-type transposons among VanA Enterococcus faecium in Poland. Eur. J. Clin. Microbiol. Infect. Dis. 2017, 36, 313-328.
140. Labibzadeh, M.; Kaydani, G.A.; Savari, M.; Ekrami, A. Emergence of High-level Gentamicin Resistance among Enterococci Clinical Isolates from Burn Patients in South-west of Iran: Vancomycin Still Working. Pol. J. Microbiol. 2018, 67, 401-406.
141. Shettigar, K.; Bhat, D.V.; Satyamoorthy, K.; Murali, T.S. Severity of drug resistance and co-existence of Enterococcus faecalis in diabetic foot ulcer infections. Folia Microbiol. 2018, 63, 115-122.
142. Esmail, M.A.M.; Abdulghany, H.M.; Khairy, R.M. Prevalence of Multidrug-Resistant Enterococcus faecalis in Hospital-Acquired Surgical Wound Infections and Bacteremia: Concomitant Analysis of Antimicrobial Resistance Genes. Infect. Dis. 2019, 12, 1178633719882929.
143. Smith, J.R.; Barber, K.E.; Raut, A.; Aboutaleb, M.; Sakoulas, G.; Rybak, M.J. beta-Lactam combinations with daptomycin provide synergy against vancomycin-resistant Enterococcus faecalis and Enterococcus faecium. J. Antimicrob. Chemother. 2015, 70, 1738-1743.
144. Smith, J.R.; Barber, K.E.; Raut, A.; Rybak, M.J. beta-Lactams enhance daptomycin activity against vancomycin-resistant Enterococcus faecalis and Enterococcus faecium in in vitro pharmacokinetic/pharmacodynamic models. Antimicrob. Agents Chemother. 2015, 59, 2842-2848.
145. Kidd, J.M.; Abdelraouf, K.; Asempa, T.E.; Humphries, R.M.; Nicolau, D.P. Pharmacodynamics of Daptomycin against Enterococcus faecium and Enterococcus faecalis in the Murine Thigh Infection Model. Antimicrob. Agents Chemother. 2018, 62.
146. Yim, J.; Smith, J.R.; Rybak, M.J. Role of Combination Antimicrobial Therapy for Vancomycin-Resistant Enterococcus faecium Infections: Review of the Current Evidence. Pharmacotherapy 2017, 37, 579-592.
147. Park, B.; Min, Y.H. In vitro synergistic effect of retapamulin with erythromycin and quinupristin against Enterococcus faecalis. J. Antibiot. 2020, 73, 630-635.
148. Carter, G.P.; Harjani, J.R.; Li, L.; Pitcher, N.P.; Nong, Y.; Riley, T.V.; Williamson, D.A.; Stinear, T.P.; Baell, J.B.; Howden, B.P. 1,2,4-Oxadiazole antimicrobials act synergistically with daptomycin and display rapid kill kinetics against MDR Enterococcus faecium. J. Antimicrob. Chemother. 2018, 73, 1562-1569.
149. Wang, H.; Lee, M.; Peng, Z.; Blazquez, B.; Lastochkin, E.; Kumarasiri, M.; Bouley, R.; Chang, M.; Mobashery, S. Synthesis and evaluation of 1,2,4-triazolo[1,5-a]pyrimidines as antibacterial agents against Enterococcus faecium. J. Med. Chem. 2015, 58, 4194-4203.
150. Zou, J.; Shankar, N. Surface protein Esp enhances pro-inflammatory cytokine expression through NF-kappaB activation during enterococcal infection. Innate Immun. 2016, 22, 31-39.
151. Taglialegna, A.; Matilla-Cuenca, L.; Dorado-Morales, P.; Navarro, S.; Ventura, S.; Garnett, J.A.; Lasa, I.; Valle, J. The biofilm-associated surface protein Esp of Enterococcus faecalis forms amyloid-like fibers. NPJ Biofilms Microbiomes 2020, 6, 15.
152. Manias, D.A.; Dunny, G.M. Expression of Adhesive Pili and the Collagen-Binding Adhesin Ace Is Activated by ArgR Family Transcription Factors in Enterococcus faecalis. J. Bacteriol. 2018, 200.
153. Montealegre, M.C.; Singh, K.V.; Somarajan, S.R.; Yadav, P.; Chang, C.; Spencer, R.; Sillanpaa, J.; Ton-That, H.; Murray, B.E. Role of the Emp Pilus Subunits of Enterococcus faecium in Biofilm Formation, Adherence to Host Extracellular Matrix Components, and Experimental Infection. Infect. Immun. 2016, 84, 1491-1500.
154. Govyrin, V.A.; Didenko, A.V.; Iazykov, V.V. [Changes in the volume of blood vessel wall in the contractile process]. Dokl. Akad. Nauk. SSSR 1988, 300, 745-747.
155. Ike, Y. Pathogenicity of Enterococci. Nihon Saikingaku Zasshi 2017, 72, 189-211.
156. Comerlato, C.B.; Resende, M.C.; Caierao, J.; d'Azevedo, P.A. Presence of virulence factors in Enterococcus faecalis and Enterococcus faecium susceptible and resistant to vancomycin. Mem. Inst. Oswaldo Cruz 2013, 108, 590-595.
157. Heidari, H.; Emaneini, M.; Dabiri, H.; Jabalameli, F. Virulence factors, antimicrobial resistance pattern and molecular analysis of Enterococcal strains isolated from burn patients. Microb. Pathog. 2016, 90, 93-97.
158. Shokoohizadeh, L.; Ekrami, A.; Labibzadeh, M.; Ali, L.; Alavi, S.M. Antimicrobial resistance patterns and virulence factors of enterococci isolates in hospitalized burn patients. BMC Res. Notes 2018, 11, 1.
159. Darisipudi, M.N.; Nordengrun, M.; Broker, B.M.; Peton, V. Messing with the Sentinels-The Interaction of Staphylococcus aureus with Dendritic Cells. Microorganisms 2018, 6, 87.
160. Jenul, C.; Horswill, A.R. Regulation of Staphylococcus aureus Virulence. Microbiol. Spectr. 2019, 7.
161. Goldmann, O.; Medina, E. Staphylococcus aureus strategies to evade the host acquired immune response. Int. J. Med. Microbiol. 2018, 308, 625-630.
162. Hobbs, M.R.; Grant, C.C.; Thomas, M.G.; Berry, S.; Morton, S.M.B.; Marks, E.; Ritchie, S.R. Staphylococcus aureus colonisation and its relationship with skin and soft tissue infection in New Zealand children. Eur. J. Clin. Microbiol. Infect. Dis. 2018, 37, 2001-2010.
163. Petry, V.; Lipnharski, C.; Bessa, G.R.; Silveira, V.B.; Weber, M.B.; Bonamigo, R.R.; d'Azevedo, P.A. Prevalence of community-acquired methicillin-resistant Staphylococcus aureus and antibiotic resistance in patients with atopic dermatitis in Porto Alegre, Brazil. Int. J. Dermatol. 2014, 53, 731-735.
164. Bukowski, M.; Piwowarczyk, R.; Madry, A.; Zagorski-Przybylo, R.; Hydzik, M.; Wladyka, B. Prevalence of Antibiotic and Heavy Metal Resistance Determinants and Virulence-Related Genetic Elements in Plasmids of Staphylococcus aureus. Front. Microbiol. 2019, 10, 805.
165. McNeil, J.C.; Fritz, S.A. Prevention Strategies for Recurrent Community-Associated Staphylococcus aureus Skin and Soft Tissue Infections. Curr. Infect. Dis. Rep. 2019, 21, 12.
166. Jauneikaite, E.; Ferguson, T.; Mosavie, M.; Fallowfield, J.L.; Davey, T.; Thorpe, N.; Allsopp, A.; Shaw, A.M.; Fudge, D.; O'Shea, M.K.; et al. Staphylococcus aureus colonization and acquisition of skin and soft tissue infection among Royal Marines recruits: A prospective cohort study. Clin. Microbiol. Infect. 2020, 26, 381.e1-381.e6.
167. Planet, P.J.; Narechania, A.; Chen, L.; Mathema, B.; Boundy, S.; Archer, G.; Kreiswirth, B. Architecture of a Species: Phylogenomics of Staphylococcus aureus. Trends Microbiol. 2017, 25, 153-166.
168. Chaves-Moreno, D.; Wos-Oxley, M.L.; Jauregui, R.; Medina, E.; Oxley, A.P.; Pieper, D.H. Exploring the transcriptome of Staphylococcus aureus in its natural niche. Sci. Rep. 2016, 6, 33174.
169. Haaber, J.; Penades, J.R.; Ingmer, H. Transfer of Antibiotic Resistance in Staphylococcus aureus. Trends Microbiol. 2017, 25, 893-905.
170. Krismer, B.; Liebeke, M.; Janek, D.; Nega, M.; Rautenberg, M.; Hornig, G.; Unger, C.; Weidenmaier, C.; Lalk, M.; Peschel, A. Nutrient limitation governs Staphylococcus aureus metabolism and niche adaptation in the human nose. PLoS Pathog. 2014, 10, e1003862.
171. Krismer, B.; Weidenmaier, C.; Zipperer, A.; Peschel, A. The commensal lifestyle of Staphylococcus aureus and its interactions with the nasal microbiota. Nat. Rev. Microbiol. 2017, 15, 675-687.
172. Balasubramanian, D.; Harper, L.; Shopsin, B.; Torres, V.J. Staphylococcus aureus pathogenesis in diverse host environments. Pathog. Dis. 2017, 75.
173. Copin, R.; Shopsin, B.; Torres, V.J. After the deluge: Mining Staphylococcus aureus genomic data for clinical associations and host-pathogen interactions. Curr. Opin. Microbiol. 2018, 41, 43-50.
174. Alibayov, B.; Baba-Moussa, L.; Sina, H.; Zdenkova, K.; Demnerova, K. Staphylococcus aureus mobile genetic elements. Mol. Biol. Rep. 2014, 41, 5005-5018.
175. Plough, H.H. Penicillin resistance of Staphylococcus aureus and its clinical implications. Am. J. Clin. Pathol. 1945, 15, 446-451.
176. McGuinness, W.A.; Malachowa, N.; DeLeo, F.R. Vancomycin Resistance in Staphylococcus aureus. Yale J. Biol. Med. 2017, 90, 269-281.
177. Furtado, G.H.; Rocha, J.; Hayden, R.; Solem, C.; Macahilig, C.; Tang, W.Y.; Chambers, R.; Figueiredo, M.L.N.; Johnson, C.; Stephens, J.; et al. Early switch/early discharge opportunities for hospitalized patients with methicillin-resistant Staphylococcus aureus complicated skin and soft tissue infections in Brazil. Braz. J. Infect. Dis. 2019, 23, 86-94.
178. Hunter, C.; Rosenfield, L.; Silverstein, E.; Petrou-Zeniou, P. Methicillin-Resistant Staphylococcus aureus Infections: A Comprehensive Review and a Plastic Surgeon's Approach to the Occult Sites. Plast. Reconstr. Surg. 2016, 138, 515-523.
179. Shettigar, K.; Jain, S.; Bhat, D.V.; Acharya, R.; Ramachandra, L.; Satyamoorthy, K.; Murali, T.S. Virulence determinants in clinical Staphylococcus aureus from monomicrobial and polymicrobial infections of diabetic foot ulcers. J. Med. Microbiol. 2016, 65, 1392-1404.
180. Richardson, J.R.; Armbruster, N.S.; Gunter, M.; Biljecki, M.; Klenk, J.; Heumos, S.; Autenrieth, S.E. PSM Peptides From Community-Associated Methicillin-Resistant Staphylococcus aureus Impair the Adaptive Immune Response via Modulation of Dendritic Cell Subsets in vivo. Front. Immunol. 2019, 10, 995.
181. Dantes, R.; Mu, Y.; Belflower, R.; Aragon, D.; Dumyati, G.; Harrison, L.H.; Lessa, F.C.; Lynfield, R.; Nadle, J.; Petit, S.; et al. National burden of invasive methicillin-resistant Staphylococcus aureus infections, United States, 2011. JAMA Intern. Med. 2013, 173, 1970-1978.
182. Katayama, Y.; Ito, T.; Hiramatsu, K. A new class of genetic element, staphylococcus cassette chromosome mec, encodes methicillin resistance in Staphylococcus aureus. Antimicrob. Agents Chemother. 2000, 44, 1549-1555.
183. He, C.; Xu, S.; Zhao, H.; Hu, F.; Xu, X.; Jin, S.; Yang, H.; Gong, F.; Liu, Q. Leukotoxin and pyrogenic toxin Superantigen gene backgrounds in bloodstream and wound Staphylococcus aureus isolates from eastern region of China. BMC Infect. Dis. 2018, 18, 395.
184. Motallebi, M.; Jabalameli, F.; Asadollahi, K.; Taherikalani, M.; Emaneini, M. Spreading of genes encoding enterotoxins, haemolysins, adhesin and biofilm among methicillin resistant Staphylococcus aureus strains with staphylococcal cassette chromosome mec type IIIA isolated from burn patients. Microb. Pathog. 2016, 97, 34-37.
185. Goudarzi, M.; Bahramian, M.; Satarzadeh Tabrizi, M.; Udo, E.E.; Figueiredo, A.M.; Fazeli, M.; Goudarzi, H. Genetic diversity of methicillin resistant Staphylococcus aureus strains isolated from burn patients in Iran: ST239-SCCmec III/t037 emerges as the major clone. Microb. Pathog. 2017, 105, 1-7.
186. Scharn, C.R.; Tenover, F.C.; Goering, R.V. Transduction of staphylococcal cassette chromosome mec elements between strains of Staphylococcus aureus. Antimicrob. Agents Chemother. 2013, 57, 5233-5238.
187. Chlebowicz, M.A.; Maslanova, I.; Kuntova, L.; Grundmann, H.; Pantucek, R.; Doskar, J.; van Dijl, J.M.; Buist, G. The Staphylococcal Cassette Chromosome mec type V from Staphylococcus aureus ST398 is packaged into bacteriophage capsids. Int. J. Med. Microbiol. 2014, 304, 764-774.
188. Munier, A.L.; de Lastours, V.; Barbier, F.; Chau, F.; Fantin, B.; Ruimy, R. Comparative dynamics of the emergence of fluoroquinolone resistance in staphylococci from the nasal microbiota of patients treated with fluoroquinolones according to their environment. Int. J. Antimicrob. Agents 2015, 46, 653-659.
189. Olufunmiso, O.; Tolulope, I.; Roger, C. Multidrug and vancomycin resistance among clinical isolates of Staphylococcus aureus from different teaching hospitals in Nigeria. Afr. Health Sci. 2017, 17, 797-807.
190. Vanegas Munera, J.M.; Ocampo Rios, A.M.; Urrego, D.M.; Jimenez Quiceno, J.N. In vitro susceptibility of methicillin-resistant Staphylococcus aureus isolates from skin and soft tissue infections to vancomycin, daptomycin, linezolid and tedizolid. Braz. J. Infect. Dis. 2017, 21, 493-499.
191. Luque, Y.; Mesnard, L. [Vancomycin nephrotoxicity: Frequency and mechanistic aspects]. Nephrol. Ther. 2018, 14 (Suppl. 1), S133-S138.
192. Zeng, D.; Debabov, D.; Hartsell, T.L.; Cano, R.J.; Adams, S.; Schuyler, J.A.; McMillan, R.; Pace, J.L. Approved Glycopeptide Antibacterial Drugs: Mechanism of Action and Resistance. Cold Spring Harbor Perspect. Med. 2016, 6.
193. Krupa, P.; Bystron, J.; Bania, J.; Podkowik, M.; Empel, J.; Mroczkowska, A. Genotypes and oxacillin resistance of Staphylococcus aureus from chicken and chicken meat in Poland. Poult. Sci. 2014, 93, 3179-3186.
194. Krupa, P.; Bystron, J.; Podkowik, M.; Empel, J.; Mroczkowska, A.; Bania, J. Population Structure and Oxacillin Resistance of Staphylococcus aureus from Pigs and Pork Meat in South-West of Poland. BioMed Res. Int. 2015, 2015, 141475.
195. McCarthy, H.; Rudkin, J.K.; Black, N.S.; Gallagher, L.; O'Neill, E.; O'Gara, J.P. Methicillin resistance and the biofilm phenotype in Staphylococcus aureus. Front. Cell Infect. Microbiol. 2015, 5, 1.
196. Cong, Y.; Yang, S.; Rao, X. Vancomycin resistant Staphylococcus aureus infections: A review of case updating and clinical features. J. Adv. Res. 2020, 21, 169-176.
197. Uddin, M.J.; Ahn, J. Associations between resistance phenotype and gene expression in response to serial exposure to oxacillin and ciprofloxacin in Staphylococcus aureus. Lett. Appl. Microbiol. 2017, 65, 462-468.
198. Costa, S.S.; Viveiros, M.; Amaral, L.; Couto, I. Multidrug Efflux Pumps in Staphylococcus aureus: An Update. Open Microbiol. J. 2013, 7, 59-71.
199. Kaatz, G.W.; DeMarco, C.E.; Seo, S.M. MepR, a repressor of the Staphylococcus aureus MATE family multidrug efflux pump MepA, is a substrate-responsive regulatory protein. Antimicrob. Agents Chemother. 2006, 50, 1276-1281.
200. Floyd, J.L.; Smith, K.P.; Kumar, S.H.; Floyd, J.T.; Varela, M.F. LmrS is a multidrug efflux pump of the major facilitator superfamily from Staphylococcus aureus. Antimicrob. Agents Chemother. 2010, 54, 5406-5412.
201. Sweeney, D.; Shinabarger, D.L.; Arhin, F.F.; Belley, A.; Moeck, G.; Pillar, C.M. Comparative in vitro activity of oritavancin and other agents against methicillin-susceptible and methicillin-resistant Staphylococcus aureus. Diagn. Microbiol. Infect. Dis. 2017, 87, 121-128.
202. Dong, G.; Liu, H.; Yu, X.; Zhang, X.; Lu, H.; Zhou, T.; Cao, J. Antimicrobial and anti-biofilm activity of tannic acid against Staphylococcus aureus. Nat. Prod. Res. 2018, 32, 2225-2228.
203. Ashraf, S.; Chaudhry, U.; Raza, A.; Ghosh, D.; Zhao, X. In vitro activity of ivermectin against Staphylococcus aureus clinical isolates. Antimicrob. Resist. Infect. Control. 2018, 7, 27.
204. Low, D.E.; Nadler, H.L. A review of in-vitro antibacterial activity of quinupristin/dalfopristin against methicillin-susceptible and -resistant Staphylococcus aureus. J. Antimicrob. Chemother. 1997, 39 (Suppl. A), 53-58.
205. Godoy-Santos, F.; Pitts, B.; Stewart, P.S.; Mantovani, H.C. Nisin penetration and efficacy against Staphylococcus aureus biofilms under continuous-flow conditions. Microbiology 2019, 165, 761-771.
206. Delpech, G.; Ceci, M.; Lissarrague, S.; Garcia Allende, L.; Baldaccini, B.; Sparo, M. In vitro activity of the antimicrobial peptide AP7121 against the human methicillin-resistant biofilm producers Staphylococcus aureus and Staphylococcus epidermidis. Biofouling 2020, 36, 266-275.
207. Li, Z.; Mao, R.; Teng, D.; Hao, Y.; Chen, H.; Wang, X.; Wang, X.; Yang, N.; Wang, J. Antibacterial and immunomodulatory activities of insect defensins-DLP2 and DLP4 against multidrug-resistant Staphylococcus aureus. Sci. Rep. 2017, 7, 12124.
208. De Souza Feitosa Lima, I.M.; Zagmignan, A.; Santos, D.M.; Maia, H.S.; Dos Santos Silva, L.; da Silva Cutrim, B.; Vieira, S.L.; Bezerra Filho, C.M.; de Sousa, E.M.; Napoleao, T.H.; et al. Schinus terebinthifolia leaf lectin (SteLL) has anti-infective action and modulates the response of Staphylococcus aureus-infected macrophages. Sci. Rep. 2019, 9, 18159.
209. Bezerra Filho, C.M.; da Silva, L.C.N.; da Silva, M.V.; Lobner-Olesen, A.; Struve, C.; Krogfelt, K.A.; Correia, M.; Vilela Oliva, M.L. Antimicrobial and Antivirulence Action of Eugenia brejoensis Essential Oil in vitro and in vivo Invertebrate Models. Front. Microbiol. 2020, 11, 424.
210. Farahpour, M.R.; Vahid, M.; Oryan, A. Effectiveness of topical application of ostrich oil on the healing of Staphylococcus aureus- and Pseudomonas aeruginosa-infected wounds. Connect. Tissue Res. 2018, 59, 212-222.
211. Farahpour, M.R.; Pirkhezr, E.; Ashrafian, A.; Sonboli, A. Accelerated healing by topical administration of Salvia officinalis essential oil on Pseudomonas aeruginosa and Staphylococcus aureus infected wound model. Biomed. Pharmacother./Biomed. Pharmacother. 2020, 128, 110120.
212. Shahini Shams Abadi, M.; Nikokar, I.; Hoseini Alfatemi, S.M.; Malekzadegan, Y.; Azizi, A.; Sedigh Ebrahim-Saraie, H. Epidemiology of Panton-Valentine Leukocidin harbouring Staphylococcus aureus in cutaneous infections from Iran: A systematic review and meta-analysis. Infez Med. 2017, 25, 217-223.
213. Foster, T.J.; Geoghegan, J.A.; Ganesh, V.K.; Hook, M. Adhesion, invasion and evasion: The many functions of the surface proteins of Staphylococcus aureus. Nat. Rev. Microbiol. 2014, 12, 49-62.
214. Ghasemian, A.; Najar Peerayeh, S.; Bakhshi, B.; Mirzaee, M. The Microbial Surface Components Recognizing Adhesive Matrix Molecules (MSCRAMMs) Genes among Clinical Isolates of Staphylococcus aureus from Hospitalized Children. Iran. J. Pathol. 2015, 10, 258-264.
215. Lin, Q.; Sun, H.; Yao, K.; Cai, J.; Ren, Y.; Chi, Y. The Prevalence, Antibiotic Resistance and Biofilm Formation of Staphylococcus aureus in Bulk Ready-To-Eat Foods. Biomolecules 2019, 9, 524.
216. Horn, J.; Stelzner, K.; Rudel, T.; Fraunholz, M. Inside job: Staphylococcus aureus host-pathogen interactions. Int. J. Med. Microbiol. 2018, 308, 607-624.
217. Olaniyi, R.O.; Pancotto, L.; Grimaldi, L.; Bagnoli, F. Deciphering the Pathological Role of Staphylococcal alpha-Toxin and Panton-Valentine Leukocidin Using a Novel Ex Vivo Human Skin Model. Front. Immunol. 2018, 9, 951.
218. Hilliard, J.J.; Datta, V.; Tkaczyk, C.; Hamilton, M.; Sadowska, A.; Jones-Nelson, O.; O'Day, T.; Weiss, W.J.; Szarka, S.; Nguyen, V.; et al. Anti-alpha-toxin monoclonal antibody and antibiotic combination therapy improves disease outcome and accelerates healing in a Staphylococcus aureus dermonecrosis model. Antimicrob. Agents Chemother. 2015, 59, 299-309.
219. Yoong, P.; Torres, V.J. The effects of Staphylococcus aureus leukotoxins on the host: Cell lysis and beyond. Curr. Opin. Microbiol. 2013, 16, 63-69.
220. Alonzo, F., 3rd; Benson, M.A.; Chen, J.; Novick, R.P.; Shopsin, B.; Torres, V.J. Staphylococcus aureus leucocidin ED contributes to systemic infection by targeting neutrophils and promoting bacterial growth in vivo. Mol. Microbiol. 2012, 83, 423-435.
221. Grumann, D.; Nubel, U.; Broker, B.M. Staphylococcus aureus toxins--their functions and genetics. Infect. Genet. Evol. 2014, 21, 583-592.
222. Nishifuji, K.; Sugai, M.; Amagai, M. Staphylococcal exfoliative toxins: "molecular scissors" of bacteria that attack the cutaneous defense barrier in mammals. J. Dermatol. Sci. 2008, 49, 21-31.
223. Zhao, C.; Liu, Y.; Zhao, M.; Liu, Y.; Yu, Y.; Chen, H.; Sun, Q.; Chen, H.; Jiang, W.; Liu, Y.; et al. Characterization of community acquired Staphylococcus aureus associated with skin and soft tissue infection in Beijing: High prevalence of PVL+ ST398. PLoS ONE 2012, 7, e38577.
224. Santosaningsih, D.; Santoso, S.; Setijowati, N.; Rasyid, H.A.; Budayanti, N.S.; Suata, K.; Widhyatmoko, D.B.; Purwono, P.B.; Kuntaman, K.; Damayanti, D.; et al. Prevalence and characterisation of Staphylococcus aureus causing community-acquired skin and soft tissue infections on Java and Bali, Indonesia. Trop. Med. Int. Health 2018, 23, 34-44.
225. Harch, S.A.J.; MacMorran, E.; Tong, S.Y.C.; Holt, D.C.; Wilson, J.; Athan, E.; Hewagama, S. High burden of complicated skin and soft tissue infections in the Indigenous population of Central Australia due to dominant Panton Valentine leucocidin clones ST93-MRSA and CC121-MSSA. BMC Infect. Dis. 2017, 17, 405.
226. Ayepola, O.O.; Olasupo, N.A.; Egwari, L.O.; Schaumburg, F. Characterization of Panton-Valentine leukocidin-positive Staphylococcus aureus from skin and soft tissue infections and wounds in Nigeria: A cross-sectional study. F1000Res 2018, 7, 1155.
227. Goudarzi, M.; Tayebi, Z.; Dadashi, M.; Miri, M.; Amirpour, A.; Fazeli, M. Characteristics of community-acquired methicillin-resistant Staphylococcus aureus associated with wound infections in Tehran, Iran: High prevalence of PVL+ t008 and the emergence of new spa types t657, t5348, and t437 in Iran. Gene Rep. 2020, 19, 100603.
228. Syed, A.K.; Reed, T.J.; Clark, K.L.; Boles, B.R.; Kahlenberg, J.M. Staphlyococcus aureus phenol-soluble modulins stimulate the release of proinflammatory cytokines from keratinocytes and are required for induction of skin inflammation. Infect. Immun. 2015, 83, 3428-3437.
229. Nakagawa, S.; Matsumoto, M.; Katayama, Y.; Oguma, R.; Wakabayashi, S.; Nygaard, T.; Saijo, S.; Inohara, N.; Otto, M.; Matsue, H.; et al. Staphylococcus aureus Virulent PSMalpha Peptides Induce Keratinocyte Alarmin Release to Orchestrate IL-17-Dependent Skin Inflammation. Cell Host Microbe 2017, 22, 667-677 e665.
230. Liu, H.; Archer, N.K.; Dillen, C.A.; Wang, Y.; Ashbaugh, A.G.; Ortines, R.V.; Kao, T.; Lee, S.K.; Cai, S.S.; Miller, R.J.; et al. Staphylococcus aureus Epicutaneous Exposure Drives Skin Inflammation via IL-36-Mediated T Cell Responses. Cell Host Microbe 2017, 22, 653-666.e5.
231. Bjornsdottir, H.; Dahlstrand Rudin, A.; Klose, F.P.; Elmwall, J.; Welin, A.; Stylianou, M.; Christenson, K.; Urban, C.F.; Forsman, H.; Dahlgren, C.; et al. Phenol-Soluble Modulin alpha Peptide Toxins from Aggressive Staphylococcus aureus Induce Rapid Formation of Neutrophil Extracellular Traps through a Reactive Oxygen Species-Independent Pathway. Front. Immunol. 2017, 8, 257.
232. Talha, M.H.; Khazaal, S.S.; Al Hadraawy, M.K.; Mostafavi, S.K.S. Screening of antibiotic resistance genes and virulence determinants of Staphylococcus aureus from skin infections. Meta Gene 2020, 100682.
233. Koymans, K.J.; Feitsma, L.J.; Bisschop, A.; Huizinga, E.G.; van Strijp, J.A.G.; de Haas, C.J.C.; McCarthy, A.J. Molecular basis determining species specificity for TLR2 inhibition by staphylococcal superantigen-like protein 3 (SSL3). Vet. Res. 2018, 49, 115.
234. Pietrocola, G.; Nobile, G.; Rindi, S.; Speziale, P. Staphylococcus aureus Manipulates Innate Immunity through Own and Host-Expressed Proteases. Front. Cell Infect. Microbiol. 2017, 7, 166.
235. Sieprawska-Lupa, M.; Mydel, P.; Krawczyk, K.; Wojcik, K.; Puklo, M.; Lupa, B.; Suder, P.; Silberring, J.; Reed, M.; Pohl, J.; et al. Degradation of human antimicrobial peptide LL-37 by Staphylococcus aureus-derived proteinases. Antimicrob. Agents Chemother. 2004, 48, 4673-4679.
236. Peetermans, M.; Vanassche, T.; Liesenborghs, L.; Claes, J.; Vande Velde, G.; Kwiecinksi, J.; Jin, T.; De Geest, B.; Hoylaerts, M.F.; Lijnen, R.H.; et al. Plasminogen activation by staphylokinase enhances local spreading of S. aureus in skin infections. BMC Microbiol. 2014, 14, 310.
237. Nguyen, L.T.; Vogel, H.J. Staphylokinase has distinct modes of interaction with antimicrobial peptides, modulating its plasminogen-activation properties. Sci. Rep. 2016, 6, 31817.
238. Henig, O.; Cober, E.; Richter, S.S.; Perez, F.; Salata, R.A.; Kalayjian, R.C.; Watkins, R.R.; Marshall, S.; Rudin, S.D.; Domitrovic, T.N.; et al. A Prospective Observational Study of the Epidemiology, Management, and Outcomes of Skin and Soft Tissue Infections Due to Carbapenem-Resistant Enterobacteriaceae. Open Forum Infect. Dis. 2017, 4, ofx157.
239. Ramirez-Blanco, C.E.; Ramirez-Rivero, C.E.; Diaz-Martinez, L.A.; Sosa-Avila, L.M. Infection in burn patients in a referral center in Colombia. Burns 2017, 43, 642-653.
240. Piperaki, E.T.; Syrogiannopoulos, G.A.; Tzouvelekis, L.S.; Daikos, G.L. Klebsiella pneumoniae: Virulence, Biofilm and Antimicrobial Resistance. Pediatric Infect. Dis. J. 2017, 36, 1002-1005.
241. Keen, E.F., 3rd; Robinson, B.J.; Hospenthal, D.R.; Aldous, W.K.; Wolf, S.E.; Chung, K.K.; Murray, C.K. Prevalence of multidrug-resistant organisms recovered at a military burn center. Burns 2010, 36, 819-825.
242. Keen, E.F., 3rd; Robinson, B.J.; Hospenthal, D.R.; Aldous, W.K.; Wolf, S.E.; Chung, K.K.; Murray, C.K. Incidence and bacteriology of burn infections at a military burn center. Burns 2010, 36, 461-468.
243. Kus, H.; Arslan, U.; Turk Dagi, H.; Findik, D. Investigation of various virulence factors of Klebsiella pneumoniae strains isolated from nosocomial infections. Mikrobiyol. Bul. 2017, 51, 329-339.
244. Lee, C.R.; Lee, J.H.; Park, K.S.; Jeon, J.H.; Kim, Y.B.; Cha, C.J.; Jeong, B.C.; Lee, S.H. Antimicrobial Resistance of Hypervirulent Klebsiella pneumoniae: Epidemiology, Hypervirulence-Associated Determinants, and Resistance Mechanisms. Front. Cell Infect. Microbiol. 2017, 7, 483.
245. Chew, K.L.; Lin, R.T.P.; Teo, J.W.P. Klebsiella pneumoniae in Singapore: Hypervirulent Infections and the Carbapenemase Threat. Front. Cell Infect. Microbiol. 2017, 7, 515.
246. Wang, G.; Zhao, G.; Chao, X.; Xie, L.; Wang, H. The Characteristic of Virulence, Biofilm and Antibiotic Resistance of Klebsiella pneumoniae. Int. J. Environ. Res. Public Health 2020, 17, 6278.
247. Guo, Y.; Zhou, H.; Qin, L.; Pang, Z.; Qin, T.; Ren, H.; Pan, Z.; Zhou, J. Frequency, Antimicrobial Resistance and Genetic Diversity of Klebsiella pneumoniae in Food Samples. PLoS ONE 2016, 11, e0153561.
248. Huan, J. Controlling infection and spread of carbapenems-resistant Klebsiella pneumoniae among burn patients. Zhonghua Shao Shang Za Zhi 2015, 31, 5-8.
249. Wang, D.; Hou, W.; Chen, J.; Mou, Y.; Yang, L.; Yang, L.; Sun, X.; Chen, M. Characterization of the blaKPC-2 and blaKPC-3 genes and the novel blaKPC-15 gene in Klebsiella pneumoniae. J. Med. Microbiol. 2014, 63, 981-987.
250. Cui, X.; Zhang, H.; Du, H. Carbapenemases in Enterobacteriaceae: Detection and Antimicrobial Therapy. Front. Microbiol. 2019, 10, 1823.
251. Vena, A.; Castaldo, N.; Bassetti, M. The role of new beta-lactamase inhibitors in gram-negative infections. Curr. Opin Infect. Dis. 2019, 32, 638-646.
252. Chung, P.Y. The emerging problems of Klebsiella pneumoniae infections: Carbapenem resistance and biofilm formation. FEMS Microbiol. Lett. 2016, 363.
253. Pang, F.; Jia, X.Q.; Zhao, Q.G.; Zhang, Y. Factors associated to prevalence and treatment of carbapenem-resistant Enterobacteriaceae infections: A seven years retrospective study in three tertiary care hospitals. Ann. Clin. Microbiol. Antimicrob. 2018, 17, 13.
254. Ghanavati, R.; Kazemian, H.; Asadollahi, P.; Heidari, H.; Irajian, G.; Navab-Moghadam, F.; Razavi, S. Characterization of antimicrobial resistance patterns of Klebsiella pneumoniae isolates obtained from wound infections. Infect. Disord. Drug Targets 2020, 20, 1.
255. Ahn, C.; Yoon, S.S.; Yong, T.S.; Jeong, S.H.; Lee, K. The Resistance Mechanism and Clonal Distribution of Tigecycline-Nonsusceptible Klebsiella pneumoniae Isolates in Korea. Yonsei Med. J. 2016, 57, 641-646.
256. Bassetti, M.; Righi, E.; Carnelutti, A.; Graziano, E.; Russo, A. Multidrug-resistant Klebsiella pneumoniae: Challenges for treatment, prevention and infection control. Expert Rev. Anti-Infect. Ther. 2018, 16, 749-761.
257. Ayerbe-Algaba, R.; Gil-Marques, M.L.; Jimenez-Mejias, M.E.; Sanchez-Encinales, V.; Parra-Millan, R.; Pachon-Ibanez, M.E.; Pachon, J.; Smani, Y. Synergistic Activity of Niclosamide in Combination With Colistin Against Colistin-Susceptible and Colistin-Resistant Acinetobacter baumannii and Klebsiella pneumoniae. Front. Cell Infect. Microbiol. 2018, 8, 348.
258. Holloway, A.J.; Yu, J.; Arulanandam, B.P.; Hoskinson, S.M.; Eaves-Pyles, T. Cystatins 9 and C as a Novel Immunotherapy Treatment That Protects against Multidrug-Resistant New Delhi Metallo-Beta-Lactamase-1-Producing Klebsiella pneumoniae. Antimicrob. Agents Chemother. 2018, 62.
259. Rabin, N.; Zheng, Y.; Opoku-Temeng, C.; Du, Y.; Bonsu, E.; Sintim, H.O. Biofilm formation mechanisms and targets for developing antibiofilm agents. Future Med. Chem. 2015, 7, 493-512.
260. Stahlhut, S.G.; Chattopadhyay, S.; Kisiela, D.I.; Hvidtfeldt, K.; Clegg, S.; Struve, C.; Sokurenko, E.V.; Krogfelt, K.A. Structural and population characterization of MrkD, the adhesive subunit of type 3 fimbriae. J. Bacteriol. 2013, 195, 5602-5613.
261. Lin, T.H.; Chen, Y.; Kuo, J.T.; Lai, Y.C.; Wu, C.C.; Huang, C.F.; Lin, C.T. Phosphorylated OmpR Is Required for Type 3 Fimbriae Expression in Klebsiella pneumoniae Under Hypertonic Conditions. Front. Microbiol. 2018, 9, 2405.
262. Martin, R.M.; Bachman, M.A. Colonization, Infection, and the Accessory Genome of Klebsiella pneumoniae. Front. Cell Infect. Microbiol. 2018, 8, 4.
263. Loraine, J.; Heinz, E.; De Sousa Almeida, J.; Milevskyy, O.; Voravuthikunchai, S.P.; Srimanote, P.; Kiratisin, P.; Thomson, N.R.; Taylor, P.W. Complement Susceptibility in Relation to Genome Sequence of Recent Klebsiella pneumoniae Isolates from Thai Hospitals. mSphere 2018, 3, e00537-18.
264. Candan, E.D.; Aksoz, N. Klebsiella pneumoniae: Characteristics of carbapenem resistance and virulence factors. Acta Biochim. Pol. 2015, 62, 867-874.
265. Fang, C.T.; Shih, Y.J.; Cheong, C.M.; Yi, W.C. Rapid and Accurate Determination of Lipopolysaccharide O-Antigen Types in Klebsiella pneumoniae with a Novel PCR-Based O-Genotyping Method. J. Clin. Microbiol. 2016, 54, 666-675.
266. Clarke, B.R.; Ovchinnikova, O.G.; Kelly, S.D.; Williamson, M.L.; Butler, J.E.; Liu, B.; Wang, L.; Gou, X.; Follador, R.; Lowary, T.L.; et al. Molecular basis for the structural diversity in serogroup O2-antigen polysaccharides in Klebsiella pneumoniae. J. Biol. Chem. 2018, 293, 4666-4679.
267. Follador, R.; Heinz, E.; Wyres, K.L.; Ellington, M.J.; Kowarik, M.; Holt, K.E.; Thomson, N.R. The diversity of Klebsiella pneumoniae surface polysaccharides. Microb. Genom. 2016, 2, e000073.
268. Hsieh, P.F.; Wu, M.C.; Yang, F.L.; Chen, C.T.; Lou, T.C.; Chen, Y.Y.; Wu, S.H.; Sheu, J.C.; Wang, J.T. D-galactan II is an immunodominant antigen in O1 lipopolysaccharide and affects virulence in Klebsiella pneumoniae: Implication in vaccine design. Front. Microbiol. 2014, 5, 608.
269. Holden, V.I.; Wright, M.S.; Houle, S.; Collingwood, A.; Dozois, C.M.; Adams, M.D.; Bachman, M.A. Iron Acquisition and Siderophore Release by Carbapenem-Resistant Sequence Type 258 Klebsiella pneumoniae. mSphere 2018, 3, e00125-18.
270. Harding, C.M.; Hennon, S.W.; Feldman, M.F. Uncovering the mechanisms of Acinetobacter baumannii virulence. Nat. Rev. Microbiol. 2018, 16, 91-102.
271. Ranjbar, R.; Farahani, A. Study of genetic diversity, biofilm formation, and detection of Carbapenemase, MBL, ESBL, and tetracycline resistance genes in multidrug-resistant Acinetobacter baumannii isolated from burn wound infections in Iran. Antimicrob. Resist. Infect. Control. 2019, 8, 172.
272. Zurawski, D.V.; Banerjee, J.; Alamneh, Y.A.; Shearer, J.P.; Demons, S.T. Skin and Soft Tissue Models for Acinetobacter baumannii Infection. Methods Mol. Biol. 2019, 1946, 271-287.
273. Ali, A.; Botha, J.; Tiruvoipati, R. Fatal skin and soft tissue infection of multidrug resistant Acinetobacter baumannii: A case report. Int. J. Surg. Case Rep. 2014, 5, 532-536.
274. Sebeny, P.J.; Riddle, M.S.; Petersen, K. Acinetobacter baumannii skin and soft-tissue infection associated with war trauma. Clin. Infect. Dis. 2008, 47, 444-449.
275. Munier, A.L.; Biard, L.; Legrand, M.; Rousseau, C.; Lafaurie, M.; Donay, J.L.; Flicoteaux, R.; Mebazaa, A.; Mimoun, M.; Molina, J.M. Incidence, risk factors and outcome of multi-drug resistant Acinetobacter baumannii nosocomial infections during an outbreak in a burn unit. Int. J. Infect. Dis. 2019, 79, 179-184.
276. Ly, T.D.A.; Kerbaj, J.; Edouard, S.; Hoang, V.T.; Louni, M.; Dao, T.L.; Benkouiten, S.; Badiaga, S.; Tissot-Dupont, H.; Raoult, D.; et al. The Presence of Acinetobacter baumannii DNA on the Skin of Homeless People and Its Relationship with Body Lice Infestation. Preliminary Results. Front. Cell Infect. Microbiol. 2019, 9, 86.
277. Davis, K.A.; Moran, K.A.; McAllister, C.K.; Gray, P.J. Multidrug-resistant Acinetobacter extremity infections in soldiers. Emerg. Infect. Dis. 2005, 11, 1218-1224.
278. Johnson, E.N.; Burns, T.C.; Hayda, R.A.; Hospenthal, D.R.; Murray, C.K. Infectious complications of open type III tibial fractures among combat casualties. Clin. Infect. Dis. 2007, 45, 409-415.
279. Albrecht, M.C.; Griffith, M.E.; Murray, C.K.; Chung, K.K.; Horvath, E.E.; Ward, J.A.; Hospenthal, D.R.; Holcomb, J.B.; Wolf, S.E. Impact of Acinetobacter infection on the mortality of burn patients. J. Am. Coll. Surg. 2006, 203, 546-550.
280. Hammoudi, D.; Moubareck, C.A.; Sarkis, D.K. How to detect carbapenemase producers? A literature review of phenotypic and molecular methods. J. Microbiol. Methods 2014, 107, 106-118.
281. Alkasaby, N.M.; El Sayed Zaki, M. Molecular Study of Acinetobacter baumannii Isolates for Metallo-beta-Lactamases and Extended-Spectrum-beta-Lactamases Genes in Intensive Care Unit, Mansoura University Hospital, Egypt. Int. J. Microbiol. 2017, 2017, 3925868.
282. Pfeifer, Y.; Hunfeld, K.P.; Borgmann, S.; Maneg, D.; Blobner, W.; Werner, G.; Higgins, P.G. Carbapenem-resistant Acinetobacter baumannii ST78 with OXA-72 carbapenemase and ESBL gene blaCTX-M-115. J. Antimicrob. Chemother. 2016, 71, 1426-1428.
283. Uddin, F.; McHugh, T.D.; Roulston, K.; Platt, G.; Khan, T.A.; Sohail, M. Detection of carbapenemases, AmpC and ESBL genes in Acinetobacter isolates from ICUs by DNA microarray. J. Microbiol. Methods 2018, 155, 19-23.
284. Nemec, A.; Musilek, M.; Maixnerova, M.; De Baere, T.; van der Reijden, T.J.; Vaneechoutte, M.; Dijkshoorn, L. Acinetobacter beijerinckii sp. nov. and Acinetobacter gyllenbergii sp. nov., haemolytic organisms isolated from humans. Int. J. Syst. Evol. Microbiol. 2009, 59, 118-124.
285. Laudy, A.E. Non-antibiotics, Efflux Pumps and Drug Resistance of Gram-negative Rods. Pol. J. Microbiol. 2018, 67, 129-135.
286. Hamouda, A.; Amyes, S.G. Novel gyrA and parC point mutations in two strains of Acinetobacter baumannii resistant to ciprofloxacin. J. Antimicrob. Chemother. 2004, 54, 695-696.
287. Doi, Y.; Murray, G.L.; Peleg, A.Y. Acinetobacter baumannii: Evolution of antimicrobial resistance-treatment options. Semin. Respir. Crit. Care Med. 2015, 36, 85-98.
288. Doi, Y.; Adams, J.M.; Yamane, K.; Paterson, D.L. Identification of 16S rRNA methylase-producing Acinetobacter baumannii clinical strains in North America. Antimicrob. Agents Chemother. 2007, 51, 4209-4210.
289. Hasani, A.; Sheikhalizadeh, V.; Ahangarzadeh Rezaee, M.; Rahmati-Yamchi, M.; Hasani, A.; Ghotaslou, R.; Goli, H.R. Frequency of Aminoglycoside-Modifying Enzymes and ArmA Among Different Sequence Groups of Acinetobacter baumannii in Iran. Microb. Drug Resist. 2016, 22, 347-353.
290. Xu, C.; Bilya, S.R.; Xu, W. adeABC efflux gene in Acinetobacter baumannii. New Microbes New Infect. 2019, 30, 100549.
291. Trebosc, V.; Gartenmann, S.; Totzl, M.; Lucchini, V.; Schellhorn, B.; Pieren, M.; Lociuro, S.; Gitzinger, M.; Tigges, M.; Bumann, D.; et al. Dissecting Colistin Resistance Mechanisms in Extensively Drug-Resistant Acinetobacter baumannii Clinical Isolates. mBio 2019, 10.
292. Whitfield, C.; Trent, M.S. Biosynthesis and export of bacterial lipopolysaccharides. Annu. Rev. Biochem. 2014, 83, 99-128.
293. Vanegas, J.M.; Higuita, L.F.; Vargas, C.A.; Cienfuegos, A.V.; Rodriguez, E.A.; Roncancio, G.E.; Jimenez, J.N. Carbapenem-resistant Acinetobacter baumannii causing osteomyelitis and infections of skin and soft tissues in hospitals of Medellin, Colombia. Biomedica 2015, 35, 522-530.
294. Carvalho, V.C.; Oliveira, P.R.; Dal-Paz, K.; Paula, A.P.; Felix Cda, S.; Lima, A.L. Gram-negative osteomyelitis: Clinical and microbiological profile. Braz. J. Infect. Dis. 2012, 16, 63-67.
295. Baginska, N.; Pichlak, A.; Gorski, A.; Jonczyk-Matysiak, E. Specific and Selective Bacteriophages in the Fight against Multidrug-resistant Acinetobacter baumannii. Virol. Sin. 2019, 34, 347-357.
296. Jeon, J.; Park, J.H.; Yong, D. Efficacy of bacteriophage treatment against carbapenem-resistant Acinetobacter baumannii in Galleria mellonella larvae and a mouse model of acute pneumonia. BMC Microbiol. 2019, 19, 70.
297. Rouse, M.D.; Stanbro, J.; Roman, J.A.; Lipinski, M.A.; Jacobs, A.; Biswas, B.; Regeimbal, J.; Henry, M.; Stockelman, M.G.; Simons, M.P. Impact of Frequent Administration of Bacteriophage on Therapeutic Efficacy in an A. baumannii Mouse Wound Infection Model. Front. Microbiol. 2020, 11, 414.
298. Dai, T.; Murray, C.K.; Vrahas, M.S.; Baer, D.G.; Tegos, G.P.; Hamblin, M.R. Ultraviolet C light for Acinetobacter baumannii wound infections in mice: Potential use for battlefield wound decontamination? J. Trauma Acute Care Surg. 2012, 73, 661-667.
299. Zhang, Y.; Zhu, Y.; Gupta, A.; Huang, Y.; Murray, C.K.; Vrahas, M.S.; Sherwood, M.E.; Baer, D.G.; Hamblin, M.R.; Dai, T. Antimicrobial blue light therapy for multidrug-resistant Acinetobacter baumannii infection in a mouse burn model: Implications for prophylaxis and treatment of combat-related wound infections. J. Infect. Dis. 2014, 209, 1963-1971.
300. Ismail, M.M.; Samir, R.; Saber, F.R.; Ahmed, S.R.; Farag, M.A. Pimenta Oil as A Potential Treatment for Acinetobacter Baumannii Wound Infection: In Vitro and In Vivo Bioassays in Relation to Its Chemical Composition. Antibiotics 2020, 9, 679.
301. Thomas, V.M.; Brown, R.M.; Ashcraft, D.S.; Pankey, G.A. Synergistic effect between nisin and polymyxin B against pandrug-resistant and extensively drug-resistant Acinetobacter baumannii. Int. J. Antimicrob. Agents 2019, 53, 663-668.
302. Morroni, G.; Simonetti, O.; Brenciani, A.; Brescini, L.; Kamysz, W.; Kamysz, E.; Neubauer, D.; Caffarini, M.; Orciani, M.; Giovanetti, E.; et al. In vitro activity of Protegrin-1, alone and in combination with clinically useful antibiotics, against Acinetobacter baumannii strains isolated from surgical wounds. Med. Microbiol. Immunol. 2019, 208, 877-883.
303. Smith, M.G.; Gianoulis, T.A.; Pukatzki, S.; Mekalanos, J.J.; Ornston, L.N.; Gerstein, M.; Snyder, M. New insights into Acinetobacter baumannii pathogenesis revealed by high-density pyrosequencing and transposon mutagenesis. Genes Dev. 2007, 21, 601-614.
304. Lee, C.R.; Lee, J.H.; Park, M.; Park, K.S.; Bae, I.K.; Kim, Y.B.; Cha, C.J.; Jeong, B.C.; Lee, S.H. Biology of Acinetobacter baumannii: Pathogenesis, Antibiotic Resistance Mechanisms, and Prospective Treatment Options. Front. Cell Infect. Microbiol. 2017, 7, 55.
305. Gaddy, J.A.; Tomaras, A.P.; Actis, L.A. The Acinetobacter baumannii 19606 OmpA protein plays a role in biofilm formation on abiotic surfaces and in the interaction of this pathogen with eukaryotic cells. Infect. Immun. 2009, 77, 3150-3160.
306. Nie, D.; Hu, Y.; Chen, Z.; Li, M.; Hou, Z.; Luo, X.; Mao, X.; Xue, X. Outer membrane protein A (OmpA) as a potential therapeutic target for Acinetobacter baumannii infection. J. Biomed. Sci. 2020, 27, 26.
307. Sanchez-Encinales, V.; Alvarez-Marin, R.; Pachon-Ibanez, M.E.; Fernandez-Cuenca, F.; Pascual, A.; Garnacho-Montero, J.; Martinez-Martinez, L.; Vila, J.; Tomas, M.M.; Cisneros, J.M.; et al. Overproduction of Outer Membrane Protein A by Acinetobacter baumannii as a Risk Factor for Nosocomial Pneumonia, Bacteremia, and Mortality Rate Increase. J. Infect. Dis. 2017, 215, 966-974.
308. Palmer, J.; Flint, S.; Brooks, J. Bacterial cell attachment, the beginning of a biofilm. J. Ind. Microbiol. Biotechnol. 2007, 34, 577-588.
309. Renner, L.D.; Weibel, D.B. Physicochemical regulation of biofilm formation. MRS Bull. 2011, 36, 347-355.
310. Camarena, L.; Bruno, V.; Euskirchen, G.; Poggio, S.; Snyder, M. Molecular mechanisms of ethanol-induced pathogenesis revealed by RNA-sequencing. PLoS Pathog. 2010, 6, e1000834.
311. Fiester, S.E.; Arivett, B.A.; Schmidt, R.E.; Beckett, A.C.; Ticak, T.; Carrier, M.V.; Ghosh, R.; Ohneck, E.J.; Metz, M.L.; Sellin Jeffries, M.K.; et al. Iron-Regulated Phospholipase C Activity Contributes to the Cytolytic Activity and Virulence of Acinetobacter baumannii. PLoS ONE 2016, 11, e0167068.
312. Kareem, S.M.; Al-Kadmy, I.M.S.; Al-Kaabi, M.H.; Aziz, S.N.; Ahmad, M. Acinetobacter baumannii virulence is enhanced by the combined presence of virulence factors genes phospholipase C (plcN) and elastase (lasB). Microb. Pathog. 2017, 110, 568-572.
313. Tomaras, A.P.; Flagler, M.J.; Dorsey, C.W.; Gaddy, J.A.; Actis, L.A. Characterization of a two-component regulatory system from Acinetobacter baumannii that controls biofilm formation and cellular morphology. Microbiology 2008, 154, 3398-3409.
314. Espinal, P.; Marti, S.; Vila, J. Effect of biofilm formation on the survival of Acinetobacter baumannii on dry surfaces. J. Hosp. Infect. 2012, 80, 56-60.
315. Singh, J.K.; Adams, F.G.; Brown, M.H. Diversity and Function of Capsular Polysaccharide in Acinetobacter baumannii. Front. Microbiol. 2018, 9, 3301.
316. Geisinger, E.; Isberg, R.R. Antibiotic modulation of capsular exopolysaccharide and virulence in Acinetobacter baumannii. PLoS Pathog. 2015, 11, e1004691.
317. Fleming, I.D.; Krezalek, M.A.; Belogortseva, N.; Zaborin, A.; Defazio, J.; Chandrasekar, L.; Actis, L.A.; Zaborina, O.; Alverdy, J.C. Modeling Acinetobacter baumannii wound infections: The critical role of iron. J. Trauma Acute Care Surg. 2017, 82, 557-565.
318. Moore, J.L.; Becker, K.W.; Nicklay, J.J.; Boyd, K.L.; Skaar, E.P.; Caprioli, R.M. Imaging mass spectrometry for assessing temporal proteomics: Analysis of calprotectin in Acinetobacter baumannii pulmonary infection. Proteomics 2014, 14, 820-828.
319. Nairn, B.L.; Lonergan, Z.R.; Wang, J.; Braymer, J.J.; Zhang, Y.; Calcutt, M.W.; Lisher, J.P.; Gilston, B.A.; Chazin, W.J.; de Crecy-Lagard, V.; et al. The Response of Acinetobacter baumannii to Zinc Starvation. Cell Host Microbe 2016, 19, 826-836.
320. Balasubramanian, D.; Schneper, L.; Kumari, H.; Mathee, K. A dynamic and intricate regulatory network determines Pseudomonas aeruginosa virulence. Nucleic Acids Res. 2013, 41, 1-20.
321. Chevalier, S.; Bouffartigues, E.; Bodilis, J.; Maillot, O.; Lesouhaitier, O.; Feuilloley, M.G.J.; Orange, N.; Dufour, A.; Cornelis, P. Structure, function and regulation of Pseudomonas aeruginosa porins. FEMS Microbiol. Rev. 2017, 41, 698-722.
322. Bassetti, M.; Vena, A.; Croxatto, A.; Righi, E.; Guery, B. How to manage Pseudomonas aeruginosa infections. Drugs Context 2018, 7, 212527.
323. Haghi, F.; Zeighami, H.; Monazami, A.; Toutouchi, F.; Nazaralian, S.; Naderi, G. Diversity of virulence genes in multidrug resistant Pseudomonas aeruginosa isolated from burn wound infections. Microb. Pathog. 2018, 115, 251-256.
324. Morand, A.; Morand, J.J. [Pseudomonas aeruginosa in dermatology]. Ann. Dermatol. Venereol. 2017, 144, 666-675.
325. Pang, Z.; Raudonis, R.; Glick, B.R.; Lin, T.J.; Cheng, Z. Antibiotic resistance in Pseudomonas aeruginosa: Mechanisms and alternative therapeutic strategies. Biotechnol. Adv. 2019, 37, 177-192.
326. Moradali, M.F.; Ghods, S.; Rehm, B.H. Pseudomonas aeruginosa Lifestyle: A Paradigm for Adaptation, Survival, and Persistence. Front. Cell Infect. Microbiol. 2017, 7, 39.
327. Sun, J.; Deng, Z.; Yan, A. Bacterial multidrug efflux pumps: Mechanisms, physiology and pharmacological exploitations. Biochem. Biophys. Res. Commun. 2014, 453, 254-267.
328. Moskowitz, S.M.; Brannon, M.K.; Dasgupta, N.; Pier, M.; Sgambati, N.; Miller, A.K.; Selgrade, S.E.; Miller, S.I.; Denton, M.; Conway, S.P.; et al. PmrB mutations promote polymyxin resistance of Pseudomonas aeruginosa isolated from colistin-treated cystic fibrosis patients. Antimicrob. Agents Chemother. 2012, 56, 1019-1030.
329. Gutu, A.D.; Sgambati, N.; Strasbourger, P.; Brannon, M.K.; Jacobs, M.A.; Haugen, E.; Kaul, R.K.; Johansen, H.K.; Hoiby, N.; Moskowitz, S.M. Polymyxin resistance of Pseudomonas aeruginosa phoQ mutants is dependent on additional two-component regulatory systems. Antimicrob. Agents Chemother. 2013, 57, 2204-2215.
330. Liu, Y.Y.; Wang, Y.; Walsh, T.R.; Yi, L.X.; Zhang, R.; Spencer, J.; Doi, Y.; Tian, G.; Dong, B.; Huang, X.; et al. Emergence of plasmid-mediated colistin resistance mechanism MCR-1 in animals and human beings in China: A microbiological and molecular biological study. Lancet Infect. Dis. 2016, 16, 161-168.
331. Mataseje, L.F.; Peirano, G.; Church, D.L.; Conly, J.; Mulvey, M.; Pitout, J.D. Colistin-Nonsusceptible Pseudomonas aeruginosa Sequence Type 654 with blaNDM-1 Arrives in North America. Antimicrob. Agents Chemother. 2016, 60, 1794-1800.
332. Fernandez, L.; Breidenstein, E.B.; Hancock, R.E. Creeping baselines and adaptive resistance to antibiotics. Drug Resist. Updates 2011, 14, 1-21.
333. Khaledi, A.; Schniederjans, M.; Pohl, S.; Rainer, R.; Bodenhofer, U.; Xia, B.; Klawonn, F.; Bruchmann, S.; Preusse, M.; Eckweiler, D.; et al. Transcriptome Profiling of Antimicrobial Resistance in Pseudomonas aeruginosa. Antimicrob. Agents Chemother. 2016, 60, 4722-4733.
334. Azzopardi, E.A.; Azzopardi, E.; Camilleri, L.; Villapalos, J.; Boyce, D.E.; Dziewulski, P.; Dickson, W.A.; Whitaker, I.S. Gram negative wound infection in hospitalised adult burn patients--systematic review and metanalysis. PLoS ONE 2014, 9, e95042.
335. Elmassry, M.M.; Mudaliar, N.S.; Colmer-Hamood, J.A.; San Francisco, M.J.; Griswold, J.A.; Dissanaike, S.; Hamood, A.N. New markers for sepsis caused by Pseudomonas aeruginosa during burn infection. Metabolomics 2020, 16, 40.
336. Ul Hassan, F.; Qudus, M.S.; Sehgal, S.A.; Ahmed, J.; Khan, M.; Ul Haq, K.; Mumtaz, S.; Arshad, M.; Siraj, S. Prevalence of Extended-Spectrum beta-Lactamases in Multi-drug Resistant Pseudomonas aeruginosa from Diabetic Foot Patients. Endocr. Metab. Immune Disord. Drug Targets 2019, 19, 443-448.
337. Al-Khudhairy, M.K.; Al-Shammari, M.M.M. Prevalence of metallo-beta-lactamase-producing Pseudomonas aeruginosa isolated from diabetic foot infections in Iraq. New Microbes New Infect. 2020, 35, 100661.
338. Otta, S.; Debata, N.K.; Swain, B. Bacteriological profile of diabetic foot ulcers. CHRISMED J. Heal. Res. 2019, 6, 7.
339. Aditi; Shariff, M.; Chhabra, S.K.; Rahman, M.U. Similar virulence properties of infection and colonization associated Pseudomonas aeruginosa. J. Med Microbiol. 2017, 66, 1489-1498.
340. Ahmed, M.A.S.; Hadi, H.A.; Hassan, A.A.I.; Abu Jarir, S.; Al-Maslamani, M.A.; Eltai, N.O.; Dousa, K.M.; Hujer, A.M.; Sultan, A.A.; Soderquist, B.; et al. Evaluation of in vitro activity of ceftazidime/avibactam and ceftolozane/tazobactam against MDR Pseudomonas aeruginosa isolates from Qatar. J. Antimicrob. Chemother. 2019, 74, 3497-3504.
341. Hirsch, E.B.; Brigman, H.V.; Zucchi, P.C.; Chen, A.; Anderson, J.C.; Eliopoulos, G.M.; Cheung, N.; Gilbertsen, A.; Hunter, R.C.; Emery, C.L.; et al. Ceftolozane-tazobactam and ceftazidime-avibactam activity against beta-lactam-resistant Pseudomonas aeruginosa and extended-spectrum beta-lactamase-producing Enterobacterales clinical isolates from U.S. medical centres. J. Glob. Antimicrob. Resist. 2020, 22, 689-694.
342. Garcia-Fernandez, S.; Garcia-Castillo, M.; Melo-Cristino, J.; Pinto, M.F.; Goncalves, E.; Alves, V.; Vieira, A.R.; Ramalheira, E.; Sancho, L.; Diogo, J.; et al. In vitro activity of ceftolozane-tazobactam against Enterobacterales and Pseudomonas aeruginosa causing urinary, intra-abdominal and lower respiratory tract infections in intensive care units in Portugal: The STEP multicenter study. Int. J. Antimicrob. Agents 2020, 55, 105887.
343. Sader, H.S.; Carvalhaes, C.G.; Streit, J.M.; Doyle, T.B.; Castanheira, M. Antimicrobial Activity of Ceftazidime-Avibactam, Ceftolozane-Tazobactam and Comparators Tested Against Pseudomonas aeruginosa and Klebsiella pneumoniae Isolates from United States Medical Centers in 2016-2018. Microb. Drug Resist. 2020.
344. Stone, G.G.; Newell, P.; Gasink, L.B.; Broadhurst, H.; Wardman, A.; Yates, K.; Chen, Z.; Song, J.; Chow, J.W. Clinical activity of ceftazidime/avibactam against MDR Enterobacteriaceae and Pseudomonas aeruginosa: Pooled data from the ceftazidime/avibactam Phase III clinical trial programme. J. Antimicrob. Chemother. 2018, 73, 2519-2523.
345. Mikhail, S.; Singh, N.B.; Kebriaei, R.; Rice, S.A.; Stamper, K.C.; Castanheira, M.; Rybak, M.J. Evaluation of the Synergy of Ceftazidime-Avibactam in Combination with Meropenem, Amikacin, Aztreonam, Colistin, or Fosfomycin against Well-Characterized Multidrug-Resistant Klebsiella pneumoniae and Pseudomonas aeruginosa. Antimicrob. Agents Chemother. 2019, 63.
346. Delgado-Valverde, M.; Conejo, M.D.C.; Serrano, L.; Fernandez-Cuenca, F.; Pascual, A. Activity of cefiderocol against high-risk clones of multidrug-resistant Enterobacterales, Acinetobacter baumannii, Pseudomonas aeruginosa and Stenotrophomonas maltophilia. J. Antimicrob. Chemother. 2020, 75, 1840-1849.
347. Iregui, A.; Khan, Z.; Landman, D.; Quale, J. Activity of Cefiderocol Against Enterobacterales, Pseudomonas aeruginosa, and Acinetobacter baumannii Endemic to Medical Centers in New York City. Microb. Drug Resist. 2020, 26, 722-726.
348. Lob, S.H.; Karlowsky, J.A.; Young, K.; Motyl, M.R.; Hawser, S.; Kothari, N.D.; Gueny, M.E.; Sahm, D.F. Activity of imipenem/relebactam against MDR Pseudomonas aeruginosa in Europe: SMART 2015-17. J. Antimicrob. Chemother. 2019, 74, 2284-2288.
349. Mwangi, J.; Yin, Y.; Wang, G.; Yang, M.; Li, Y.; Zhang, Z.; Lai, R. The antimicrobial peptide ZY4 combats multidrug-resistant Pseudomonas aeruginosa and Acinetobacter baumannii infection. Proc. Natl. Acad. Sci. USA 2019, 116, 26516-26522.
350. Meskini, M.; Esmaeili, D. The study of formulated Zoush ointment against wound infection and gene expression of virulence factors Pseudomonas aeruginosa. BMC Complement. Altern. Med. 2018, 18, 185.
351. Lenzmeier, T.D.; Mudaliar, N.S.; Stanbro, J.A.; Watters, C.; Ahmad, A.; Simons, M.P.; Ventolini, G.; Zak, J.C.; Colmer-Hamood, J.A.; Hamood, A.N. Application of Lactobacillus gasseri 63 AM supernatant to Pseudomonas aeruginosa-infected wounds prevents sepsis in murine models of thermal injury and dorsal excision. J. Med. Microbiol. 2019, 68, 1560-1572.
352. Argenta, A.; Satish, L.; Gallo, P.; Liu, F.; Kathju, S. Local Application of Probiotic Bacteria Prophylaxes against Sepsis and Death Resulting from Burn Wound Infection. PLoS ONE 2016, 11, e0165294.
353. Ferro, T.A.F.; Souza, E.B.; Suarez, M.A.M.; Rodrigues, J.F.S.; Pereira, D.M.S.; Mendes, S.J.F.; Gonzaga, L.F.; Machado, M.; Bomfim, M.R.Q.; Calixto, J.B.; et al. Topical Application of Cinnamaldehyde Promotes Faster Healing of Skin Wounds Infected with Pseudomonas aeruginosa. Molecules 2019, 24, 1627.
354. Hegerle, N.; Choi, M.; Sinclair, J.; Amin, M.N.; Ollivault-Shiflett, M.; Curtis, B.; Laufer, R.S.; Shridhar, S.; Brammer, J.; Toapanta, F.R.; et al. Development of a broad spectrum glycoconjugate vaccine to prevent wound and disseminated infections with Klebsiella pneumoniae and Pseudomonas aeruginosa. PLoS ONE 2018, 13, e0203143.
355. Hashemi, F.B.; Behrouz, B.; Irajian, G.; Laghaei, P.; Korpi, F.; Fatemi, M.J. A trivalent vaccine consisting of "flagellin A+B and pilin" protects against Pseudomonas aeruginosa infection in a murine burn model. Microb. Pathog. 2020, 138, 103697.
356. Ben Haj Khalifa, A.; Moissenet, D.; Vu Thien, H.; Khedher, M. Virulence factors in Pseudomonas aeruginosa: Mechanisms and modes of regulation. Ann. Biol. Clin. 2011, 69, 393-403.
357. Finlayson, E.A.; Brown, P.D. Comparison of antibiotic resistance and virulence factors in pigmented and non-pigmented Pseudomonas aeruginosa. West. Indian Med. J. 2011, 60, 24-32.
358. Chaney, S.B.; Ganesh, K.; Mathew-Steiner, S.; Stromberg, P.; Roy, S.; Sen, C.K.; Wozniak, D.J. Histopathological comparisons of Staphylococcus aureus and Pseudomonas aeruginosa experimental infected porcine burn wounds. Wound Repair Regen. 2017, 25, 541-549.
359. Hauser, A.R. The type III secretion system of Pseudomonas aeruginosa: Infection by injection. Nat. Rev. Genet. 2009, 7, 654-665.
360. Mishra, M.; Panda, S.; Barik, S.; Sarkar, A.; Singh, D.V.; Mohapatra, H. Antibiotic Resistance Profile, Outer Membrane Proteins, Virulence Factors and Genome Sequence Analysis Reveal Clinical Isolates of Enterobacter Are Potential Pathogens Compared to Environmental Isolates. Front. Cell Infect. Microbiol. 2020, 10, 54.
361. Annavajhala, M.K.; Gomez-Simmonds, A.; Uhlemann, A.C. Multidrug-Resistant Enterobacter cloacae Complex Emerging as a Global, Diversifying Threat. Front. Microbiol. 2019, 10, 44.
362. Zhao, Y.; Zhang, J.; Fu, Y.; Li, C.; Hu, K.; Su, S.; Yu, L.; Guo, Y.; Fu, Y.; Zhang, X. Molecular characterization of metallo-beta-lactamase- producing carbapenem-resistant Enterobacter cloacae complex isolated in Heilongjiang Province of China. BMC Infect. Dis. 2020, 20, 94.
363. Davin-Regli, A.; Lavigne, J.P.; Pages, J.M. Enterobacter spp.: Update on Taxonomy, Clinical Aspects, and Emerging Antimicrobial Resistance. Clin. Microbiol. Rev. 2019, 32.
364. Mezzatesta, M.L.; Gona, F.; Stefani, S. Enterobacter cloacae complex: Clinical impact and emerging antibiotic resistance. Future Microbiol. 2012, 7, 887-902.
365. Lee, J.H.; Bae, I.K.; Lee, C.H.; Jeong, S. Molecular Characteristics of First IMP-4-Producing Enterobacter cloacae Sequence Type 74 and 194 in Korea. Front. Microbiol. 2017, 8, 2343.
366. Gomez-Simmonds, A.; Annavajhala, M.K.; Wang, Z.; Macesic, N.; Hu, Y.; Giddins, M.J.; O'Malley, A.; Toussaint, N.C.; Whittier, S.; Torres, V.J.; et al. Genomic and Geographic Context for the Evolution of High-Risk Carbapenem-Resistant Enterobacter cloacae Complex Clones ST171 and ST78. mBio 2018, 9, e00542-18.
367. Alvarez-Marin, R.; Navarro-Amuedo, D.; Gasch-Blasi, O.; Rodriguez-Martinez, J.M.; Calvo-Montes, J.; Lara-Contreras, R.; Lepe-Jimenez, J.A.; Tubau-Quintano, F.; Cano-Garcia, M.E.; Rodriguez-Lopez, F.; et al. A prospective, multicenter case control study of risk factors for acquisition and mortality in Enterobacter species bacteremia. J. Infect. 2020, 80, 174-181.
368. Jolivet, S.; Lescure, F.X.; Armand-Lefevre, L.; Raffoul, R.; Dilly, M.P.; Ghodbane, W.; Nataf, P.; Lucet, J.C. Surgical site infection with extended-spectrum beta-lactamase-producing Enterobacteriaceae after cardiac surgery: Incidence and risk factors. Clin. Microbiol. Infect. 2018, 24, 283-288.
369. Azevedo, P.A.A.; Furlan, J.P.R.; Oliveira-Silva, M.; Nakamura-Silva, R.; Gomes, C.N.; Costa, K.R.C.; Stehling, E.G.; Pitondo-Silva, A. Detection of virulence and beta-lactamase encoding genes in Enterobacter aerogenes and Enterobacter cloacae clinical isolates from Brazil. Braz. J. Microbiol. 2018, 49 (Suppl. 1), 224-228.
370. Park, H.S.; Pham, C.; Paul, E.; Padiglione, A.; Lo, C.; Cleland, H. Early pathogenic colonisers of acute burn wounds: A retrospective review. Burns 2017, 43, 1757-1765.
371. Yuan, S.; Wu, G.; Zheng, B. Complete genome sequence of an IMP-8, CTX-M-14, CTX-M-3 and QnrS1 co-producing Enterobacter asburiae isolate from a patient with wound infection. J. Glob. Antimicrob. Resist. 2019, 18, 52-54.
372. Hadano, Y.; Tamagawa, K.; Ohkusu, K. Trauma Wound Related Infection Caused by Enterobacter cancerogenus and Aeromonas hydrophilia. Intern. Med. 2018, 57, 131-133.
373. Yang, H.; Wang, W.S.; Tan, Y.; Zhang, D.J.; Wu, J.J.; Lei, X. Investigation and analysis of the characteristics and drug sensitivity of bacteria in skin ulcer infections. Chin. J. Traumatol. 2017, 20, 194-197.
374. Haciseyitoglu, D.; Dokutan, A.; Abulaila, A.; Erdem, F.; Cag, Y.; Ozer, S.; Aktas, Z. The First Enterobacter cloacae Co-Producing NDM and OXA-48 Carbapenemases and Interhospital Spread of OXA-48 and NDM-Producing Klebsiella pneumoniae in Turkey. Clin. Lab. 2017, 63, 1213-1222.
375. Chavda, K.D.; Chen, L.; Fouts, D.E.; Sutton, G.; Brinkac, L.; Jenkins, S.G.; Bonomo, R.A.; Adams, M.D.; Kreiswirth, B.N. Comprehensive Genome Analysis of Carbapenemase-Producing Enterobacter spp.: New Insights into Phylogeny, Population Structure, and Resistance Mechanisms. mBio 2016, 7, e02093-16.
376. Yang, H.; Chen, G.; Hu, L.; Liu, Y.; Cheng, J.; Ye, Y.; Li, J. Enhanced efficacy of imipenem-colistin combination therapy against multiple-drug-resistant Enterobacter cloacae: In vitro activity and a Galleria mellonella model. J. Microbiol. Immunol. Infect. 2018, 51, 70-75.
377. Manohar, P.; Nachimuthu, R.; Lopes, B.S. The therapeutic potential of bacteriophages targeting gram-negative bacteria using Galleria mellonella infection model. BMC Microbiol. 2018, 18, 97.
378. Manohar, P.; Tamhankar, A.J.; Lundborg, C.S.; Nachimuthu, R. Therapeutic Characterization and Efficacy of Bacteriophage Cocktails Infecting Escherichia coli, Klebsiella pneumoniae, and Enterobacter Species. Front. Microbiol. 2019, 10, 574.
Gustavo Henrique Rodrigues Vale de Macedo
1,2,
Gabrielle Damasceno Evangelista Costa
1,2,
Elane Rodrigues Oliveira
2,
Glauciane Viera Damasceno
2,
Juliana Silva Pereira Mendonça
1,2,
Lucas dos Santos Silva
2,
Vitor Lopes Chagas
2,
José Manuel Noguera Bazán
3,
Amanda Silva dos Santos Aliança
4,
Rita de Cássia Mendonça de Miranda
1,5,
Adrielle Zagmignan
2,
Andrea de Souza Monteiro
1,6 and
Luís Cláudio Nascimento da Silva
1,2,3,*
1Programa de Pós-graduação em Biologia Microbiana, Universidade CEUMA, 65075-120 São Luís, Brazil
2Laboratório de Patogenicidade Microbiana, Universidade CEUMA, 65075-120 São Luís, Brazil
3Programa de Pós-graduação em Odontologia, Universidade CEUMA, 65075-120 São Luís, Brazil
4Programa de Pós-graduação em Gestão de Programas e Serviços de Saúde, Universidade CEUMA, 65075-120 São Luís, Brazil
5Programa de Pós-graduação em Meio Ambiente, Universidade CEUMA, 65075-120 São Luís, Brazil
6Laboratório de Microbiologia Aplicada, Universidade CEUMA, 65075-120 São Luís, Brazil
*Author to whom correspondence should be addressed.
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