Commonly Encountered Skin Biome-Derived Pathogens after Orthopedic Surgery

Abstract

Background:

Normal skin microbiota influence susceptibility to surgical infections. The distribution of skin bacteria differs by anatomic site, and given the right conditions, almost any of these bacteria can become an opportunistic pathogen.

Methods:

This paper provides a thorough review of the most commonly encountered bacteria in various regions of the body and their isolation from operative incisions at those locations. These data may be useful in optimizing targeted antibiotic therapy for surgical site infections and provide a better understanding of the skin biome distribution at specific surgical sites.

Conclusion:

Typical skin-borne flora, surgical site infections, orthopedic infections by body part, and drug-resistant pathogens are reviewed.

Understanding which microbial species are present on the skin and mucosal tissues has become a useful method for predicting which pathogens are likely to be isolated from surgical site infections (SSIs). Some species of skin-borne bacteria and fungi are found predominately at specific sites on the body; therefore, understanding their distribution will help identify whether their presence at a particular surgical site is to be expected. Knowledge of the common skin biome and the respective locations during surgical intervention will allow more informed selection of antibiotics to prevent or manage SSIs.

Normal Skin Microbiome

Roughly one trillion bacteria live on human skin. Until recently, our knowledge of the skin microbiota has depended largely on media culture assays, although it is estimated that fewer than 1% of bacterial species can be cultured [1]. Phylogenetic identification via 16s ribosomal RNA gene sequences, which carry a common threshold of approximately 97% sequence identity, allows species-specific phylotype determination [2].

Recent studies have shown significant intra-personal and inter-personal variability in the composition of bacterial communities. Fierer et al. observed that microbiota on the two hands of the same individual shared only 17% of their phylotypes, and those of different individuals shared only 13% [3]. Furthermore, it was noted that women had distinctly higher bacterial diversity than men, and community composition was affected significantly by handedness, time since last hand washing, and an individual's sex.

The most commonly characterized skin-borne bacterium is Staphylococcus aureus, which is carried by approximately one-third of the population in the anterior nares, as well as on the skin and in the respiratory tract [4 –8]. Staphylococcus aureus is a commensal organism of the normal skin biome; however, if the skin barrier is disrupted, it can cause infection. Indeed, the organism is the most prevalent pathogen isolated from skin and soft tissue infections (SSTIs) [9,10]. With an estimated incidence of 32 infections per 100,000 persons, S. aureus has surpassed Streptococcus pneumoniae and Haemophilus influenzae as the most common invasive bacterial pathogen in the United States [11,12].

In general, bacterial diversity and composition are dependent on the normal physiology of the skin site, with specific bacteria associated with moist, dry, and sebaceous environments [13]. Diversity seems to be lowest around regions rich with sebaceous glands, suggesting that a specific subset of organisms can better tolerate these regions. Areas such as the back, forehead, retroauricular crease, and alar crease exhibit low phylotype diversity [13 –15]. Propionibacterium species, such as P. (Cutibacterium) acnes, are the dominant organisms at these sites, which supports classical microbiological studies describing these species as lipophilic residents of the pilosebaceous unit (Table 1) [13].

Table 1.

Proportion of Major Bacterial Groups when Sites Are Clustered into Microenvironments

	Type of site
	Sebaceous (%)	Moist (%)	Dry (%)
Actinobacteria
Propionibacterium	46	7	13
Corynebacterium	10	28	15
Other	4	1	0
Firmicutes
Staphylococcus	16	22	5
Lactobacillales	3	2	4
Clostridiales	1	2	3
Proteobacteria
Alpha	1	1	2
Beta	9	21	32
Gamma	1	3	7
Bacteroidetes
Flavobacteriales	3	9	14
Bacteroidales	1	0	0
Other	5	4	5

Data from references 1, 13, and 15.

Staphylococcus and Corynebacterium spp. are the most dominant organisms colonizing moist areas, which is consistent with culture data, suggesting that these organisms prefer regions of high humidity [1,13,15]. Moist sites include the umbilicus, axilla, inguinal crease, gluteal crease, sole of the foot, popliteal fossa, and antecubital fossa. The sites with the highest bacterial diversity are the dry areas, such as the forearm, buttock, and various regions of the hand, which display mixed representation from the phyla Actinobacteria, Proteobacteria, Firmicutes, and Bacteriodetes [1,13,15].

Skin, Soft Tissue, and Surgical Site Infections

Most SSIs are associated with skin-derived gram-positive cocci, including S. aureus, coagulase-negative staphylococci (CoNS) (usually Staphylococcus epidermidis), streptococci, and Enterococcus spp. [16,17]. In addition to SSIs caused by staphylococci, gastrointestinal and some head and neck operations have a higher likelihood of being associated with infection caused by enteric aerobic (e.g., Escherichia coli) and anaerobic (e.g., Bacteroides fragilis) gram-negative bacilli [18,19].

Overall, S. aureus is the pathogen most commonly encountered in infections following surgical procedures. Table 2 outlines some of the pathogens commonly associated with various surgical procedures. From the accrued data, it appears that specific types of procedures align with specific groups of organisms [20], thus providing a possible map for use in prophylactic care.

Table 2.

Pathogens Commonly Associated with Different Surgical Procedures

Type of surgery	Common pathogens
Placement of graft, prosthesis, or implant	Staphylococcus aureus; CoNS
Cardiac	S. aureus; CoNS
Neurosurgery	S. aureus; CoNS
Breast	S. aureus; CoNS
Ophthalmic	S. aureus; CoNS; streptococci; gram-negative bacilli
Orthopaedic	S. aureus; CoNS; gram-negative bacilli
Non-cardiothoracic	S. aureus; CoNS; Streptococcus pneumonia; gram-negative bacilli
Vascular	S. aureus; CoNS
Appendectomy	Gram-negative bacilli; anaerobes
Biliary tract	Gram-negative bacilli; anaerobes
Colorectal	Gram-negative bacilli; anaerobes
Gastroduodenal	Gram-negative bacilli; streptococci; oropharyngeal anaerobes (e.g., peptostreptococci)
Head and neck	S. aureus; streptococci; oropharyngeal anaerobes (e.g., peptostreptococci)
Obstetric and gynecologic	Gram-negative bacilli; enterococci; Group B streptococci; anaerobes
Urologic	Gram-negative bacilli

CoNS = coagulase-negative staphylococci [17].

The SENTRY Antimicrobial Surveillance Program database provides a long-term analysis of the microbial causes of skin and soft tissue infections in hospitalized patients [21]. The pathogens most consistently encountered over time are S. aureus, Pseudomonas aeruginosa, E. coli, and Enterococcus spp.

The microbiology of SSTIs can differ remarkably with the means of entry (Table 3) [22]. The cause of SSTIs may be normal host flora transferred from an instrument or from the environment [23,24]. Etiologies can differ significantly between hospital-acquired and community-acquired SSTIs. As mentioned, hospital-acquired SSTIs are caused predominantly by S. aureus, whereas community-acquired SSTIs typically are caused by β-hemolytic streptococci, usually Lancefield groups A, C, and G, with group B being seen in patients with diabetes mellitus and in the elderly [25 –29]. Clinically, the microbial etiology of SSTIs often can be predicted with some accuracy. Localized pus-producing lesions such as boils, abscesses, and carbuncles usually are staphylococcal in etiology. Alternatively, rapidly spreading infections such as erysipelas, lymphangitis, and cellulitis typically are caused by β-hemolytic streptococci [25], which also are prevalent in cases of necrotizing fasciitis.

Table 3.

Etiologic Risk Factors for Skin and Soft Tissue Infections and Their Associated Bacterial Causes

Risk factor	Associated etiologic pathogen
Diabetes mellitus	Staphylococcus aureus, group B streptococci, anaerobes, gram-negative bacilli
Cirrhosis	Campylobacter fetus, Klebsiella pneumoniae, Escherichia coli, Capnocytophaga canimorsus, other gram-negative bacilli, Vibrio vulnificus
Neutropenia	Pseudomonas aeruginosa
Bite wounds
Human	Oral flora (Eikenella corrodens)
Cat	Pastureurella multocida
Dog	C canimorsus, P multocida
Rat	Streptobacillus moniliformis
Animal contact	Campylobacter spp.
Reptile contact	Salmonella spp.
Hot tub exposure/loofah sponge	P. aeruginosa
Freshwater exposure	Aeromonas hydrophila
Seawater (fish tank) exposure	V. vulnificus, Mycobacterium marinum
Intravenous drug abuse	Methicillin-resistant S. aureus, P. aeruginosa
Subcutaneous drug abuse	Anaerobes, especially E corrodens

Data from reference 19.

Orthopedic Infections

Orthopedic-specific procedures make up a significant number of all operations. For example, when Molina et al. surveyed the American College of Surgeons' National Surgical Quality Improvement Program (NSQIP) to quantify the number of orthopedic procedures performed between 2005 and 2011 in more than 400 hospitals around the world [30], they found that orthopedic procedures accounted for roughly 7% of all operations. Of these, total knee arthroplasty (TKA), total hip arthroplasty (THA), and knee arthroscopy with meniscectomy comprised 55% of all orthopedic procedures, which matched the numbers reported by the American Academy of Orthopaedic Surgeons in their 2014 practitioner census [31]. Other frequently performed procedures are laminectomy and nerve root decompression, total shoulder arthroplasty, and revision TKAs and THAs. Among the most frequently performed procedures, those investigators found a 5% complication rate [30].

As the United States' population continues to age, the prevalence of age-related disorders, such as osteoarthritis, will continue to rise. According to the Centers for Disease Control and Prevention (CDC), the total number of THAs and TKAs performed has been increasing. In 2010, approximately 332,000 primary THAs and 719,000 primary TKAs were performed on patients 45 years or older [32]. Kurtz et al. project that by 2030, the demand for THAs will increase by 174% and the demand for TKAs by 673%, accounting for roughly 3.48 million surgical procedures annually [33].

As the total number of orthopedic procedures increases, a concurrent rise in the number of SSIs is expected. Furthermore, the incidence of musculoskeletal infection, including peri-prosthetic joint infection, soft tissue infection, septic arthritis, and osteomyelitis, is increasing in proportion to the aging population and higher rates of diabetes mellitus and obesity [34]. Therefore, a thorough understanding of which types of bacterial infections are most prevalent at each surgical site can benefit subsequent intervention greatly. Figure 1 describes some of the most common surgical interventions at each anatomic site, as well as the pathogens most commonly found in the associated SSIs.

FIG. 1.

Common procedures by anatomic site and most common pathogens encountered in surgical site infections [data from references 2, 5, 8, 10, 12, and 16].

As Figure 1 details, frequent surgical sites, including the spine, shoulder, elbow, hand, hip, knee, and foot and ankle, are associated with various commonly cultured pathogenic species when post-surgical infection occurs. These data are represented in Table 4.

Table 4.

Pathogen Species Most Commonly Isolated from Infections after Non-Emergency Spinal Surgery by Site

	Organism	Percent of total infections
Spine [30]	Staphylococcus aureus	45.2
	Methicillin sensitive	29.7
	Methicillin resistant	15.5
	S. epidermidis	31.4
	Enterococcus spp.	17.6
	Escherichia coli	10
	Propionibacterium acnes	7.9
Elbow [31]	S. aureus	69
	Serratia marcescens	10
	Enterococcus spp.	7
	Bacillus spp.	5
	Other	9
Hand [32]	S. aureus	76
	Methicillin resistant	53
	Methicillin sensitive	23
	Streptococcal spp.	25
	Coagulase-negative Staphylococcus (CoNS)	6
	Klebsiella	4
	Diphtheroids	2
Foot and ankle [33]	S. aureus	17
	Methicillin sensitive	11
	Methicillin resistant	6
	CoNS	6
	Enterobacter cloacae	2
	P. acnes	2
	Acinetobacter baumannii	1
Hip [38]	CoNS	67
	S. aureus	13
	Streptococcal spp.	9
	E. coli	6
	All other	5
Knee [39]	CoNS	49
	S. aureus	13
	E. coli	7
	Enterococcus faecalis	6
	Other	25
Shoulder [42 –44]	P. acnes	27.5
	CoNS	13.7
	S. aureus	13.7
	Methicillin sensitive	7.8
	Methicillin resistant	5.9
	Ent. cloacae	5.9
	Finegoldia magna	5.9

Staphylococcus aureus is the most common pathogen in infections of the spine, elbow, hand, and foot and ankle, accounting for 45.2%, 69%, 76%, and 17% of infections, respectively (Table 4) [35 –38]. Although they share the most common etiologic agent, the incidence of SSIs at these sites differs greatly. In spine surgery, for example, the overall incidence of SSIs ranges from 0.5% to 18.8% [39]. In contrast, because of its thin soft-tissue envelope, SSIs resulting from surgical intervention on the elbow are frequent, with an incidence of 4.5% [40]. Rates of post-operative infection in hand surgery are poorly documented but include a wide range of both emergency and non-emergency procedures. The incidence of SSIs in hand surgery has been reported to be as high as 9.7% [41]. Lastly, because of the wide array of procedures involving the foot and ankle, the incidence of SSIs associated with these interventions ranges from 1% to 8% [42].

Currently, CoNS are the most common pathogen in SSIs of the hip and knee, accounting for 67% and 49% of post-operative infection at these sites, respectively. At both sites, S. aureus is the second most commonly isolated pathogen but is becoming increasingly implicated in infections (Table 4) [43,44]. With regard to hip surgery, the rate of infection differs significantly depending on the procedure, with SSI rates reported to be as low as 1% for non-emergency cases [45]. Likewise, the incidence of SSIs during knee surgery is low, ranging from 0.86% to 2.5% [46].

The incidence of SSIs in shoulder surgery ranges from 0.76% to 5.8% [47]. In contrast to the sites discussed earlier, SSIs after shoulder surgery are not caused predominantly by S. aureus or CoNS but rather often by P. acnes, the most commonly isolated pathogen in shoulder-related SSIs, accounting for roughly 27.5% of such infections [48,49]. Propionibacterium acnes requires 14 days to culture, and contamination often is mistaken for pathogen growth. Because it is notoriously difficult to culture and quantify, P. acnes typically is underdiagnosed as a cause of infection. Additionally, the organism has shown resistance to commonly used skin preparation agents, such as chlorhexidine gluconate [50]. Thus, it is possible that the prevalence of P. acnes in shoulder SSIs is underestimated.

Skin Preparation Prior to Incision in Orthopedic Surgery

A plethora of skin preparation solutions are currently used prior to incision, with the goal of removing skin-dwelling bacteria to prevent inoculation of the incision during surgery. Because the skin microbiome differs across locations, it is reasonable that different skin preparation methods are favored depending on the operative site. Commonly used preparations include ChloraPrep, which is 2% chlorhexidine gluconate and 70% isopropyl alcohol (BD, Franklin Lakes, NJ); DuraPrep, 0.7% iodophor and 74% isopropyl alcohol (3M, Chelmsford, MA); povidone–iodine scrub and paint (0.75% and 1.0% iodine, respectively); Techni-Care, 3.0% chloroxylenol (Care-Tech Laboratories, St. Louis, MO); Betadine 10% povidone–iodine; and 2% chlorhexidine gluconate (CHG) cloth.

In shoulder surgery, ChloraPrep is more effective than both DuraPrep and povidone–iodine scrub and paint at removing bacteria from the region, with post-treatment positive-culture rates of 7%, 19%, and 31%, respectively. Although ChloraPrep and DuraPrep are more effective than povidone–iodine at eliminating CoNS from the shoulder, all are equally able to remove P. acnes. Of note, there was no difference among the preparation agents in the post-operative infection rate [51].

Similarly, ChloraPrep is more effective than DuraPrep and Techni-Care at eliminating bacteria from the skin prior to foot and ankle surgery. However, ChloraPrep still was associated with a high post-application positive culture rate, 21%, and there were no significant differences in the rate of post-operative infections among the three agents [52].

When comparing Betadine, DuraPrep, and Chloraprep as preparation for hand surgery, the overall culture rates are 43.2%, 30.4%, and 40%, respectively. Both Betadine and DuraPrep are more effective than ChloraPrep at removing Bacillus, but this organism rarely causes SSIs [37,53]. All three solutions yield equally low rates of positive CoNS and P. acnes cultures. Similar results were obtained in shoulder and foot and ankle surgery; the three preparations did not differ in the post-operative infection rate [53].

In the lumbar spine, ChloraPrep and DuraPrep yield similar post-preparation positive culture rates: 0 and 6%, respectively. Pathogens present on the skin after DuraPrep application were Propionibacterium and Peptostreptococcus [54]. However, on the upper back, where P. acnes commonly resides, ChloraPrep is ineffective at penetrating deep into the dermal layer, and persistence of P. acnes is seen in as many as 70% of patients [55].

With hip and knee surgery, at-home use of a 2% CHG cloth prior to surgery shows additional benefits. Applying this cloth to the surgical site the night before and the morning of surgery reduced the infection rate by half (3.19% to 1.59%) [56]. The rate of SSI after total hip arthroplasty was reduced further when the cloth was applied to the whole body as opposed to only the hip. Additionally, two at-home total body 2% CHG applications followed by DuraPrep application in the operating room consistently resulted in fewer SSIs than DuraPrep alone [57,58]. However, it is unclear which pathogens contributed to SSI and which were eliminated by skin preparation.

Drug-Resistant Bacteria Strains

Clinically important drug-resistant bacteria that commonly cause nosocomial infections include methicillin-resistant S. aureus (MRSA), methicillin-resistant CoNS, vancomycin-resistant enterococci, and multi-drug-resistant gram-negative bacilli, including strains of Pseudomonas aeruginosa and E. coli [59]. Each of these will be considered in turn, with a special focus on MRSA S. aureus.

Methicillin-resistant Staphylococcus aureus (MRSA)

In orthopedics, staphylococci are the primary cause of implant-related infections, being responsible for nearly 80% of all prosthetic infections [60]. Although initially considered primarily a nosocomial pathogen, MRSA has become increasingly prevalent over the last several years. The CDC reports that 0.8% to 1.5% of the general public is now colonized by MRSA, which accounts for nearly 60% of all S. aureus isolates recovered from hospital intensive care units [61]. At present, it is projected that MRSA infections account for more than 100,000 hospitalizations each year in the United States [62].

Considerable variation in the resistance rate of S. aureus to methicillin has been noted between countries and continents, with the highest rate in North America (35.9%), followed by Latin America (29.4%) and Europe (22.8%) [21]. The MRSA strains are further classified into either community-acquired (CA) or healthcare-acquired (HA), with each type differing in pathogenicity, virulence, antibiotic resistance profile, and the patient population affected [63]. However, the boundaries between HA-MRSA and CA-MRSA are becoming blurred because of the movement of patients between hospitals and the community [64,65]. For example, there are many documented HA outbreaks of CA-MRSA following the admission of colonized or infected patients [65]. In the United States, where CA-MRSA is becoming common, it is increasingly difficult to distinguish between CA- and HA-MRSA on clinical and epidemiologic grounds [64,66]. Because HA- and CA-MRSA strains often are genotypically and phenotypically different, the microbiological characteristics of the isolates may help to distinguish between the two types of infection [27,67]. Methicillin-resistant S. aureus colonizes the anterior nares, throat, rectum, and skin (axilla, inguinal area, and perineum) [5 –8, 68 –71]. Roughly 10% of healthy individuals in the United States are colonized with MRSA, and the prevalence has increased significantly over the past decade, accompanied by an increase in MRSA infection rates [68,72]. Whereas notable risk factors for acquiring HA-MRSA include antibiotic use, MRSA carriage, proximity to infected individuals, prolonged hospitalization, intensive care admission, and hemodialysis, CA-MRSA typically is associated with skin and soft-tissue infections in young and healthy individuals with no recent healthcare exposure [63,73 –79].

Although CA-MRSA always has been a prominent cause of staphylococcal skin and soft-tissue infections, this drug-resistant pathogen has become increasingly prevalent recently. In a review of MRSA SSTIs in South Texas, there were fewer than 10 cases reported annually throughout the 1990s, but by 2003, roughly 500 cases were being reported each year [80]. By 2005, several centers across the United States reported that CA-MRSA accounted for nearly 75% of all staphylococcal infections [81]. With such unexpectedly high rates of CA-MRSA, changes in empirical antibiotic therapy were needed whenever MRSA was suspected, particularly for SSTI, among which more than 50% were caused by CA-MRSA [80 –85]. A retrospective review of more than 40 tertiary-care children's hospitals in the United States identified almost 60,000 children with S. aureus infections from 2002 to 2007. Of those, 51% were infected with MRSA. Over the course of the years studied, MRSA rates increased significantly, whereas rates of methicillin-sensitive S. aureus infection remained stable. Of the nearly 40,000 patients whose type of infection was identified, 61% had SSTIs. As a result, the authors deduced that the increasing rate of MRSA infections is driven largely by an increase in SSTIs [88].

Children and African Americans are disproportionally affected by increases in CA-MRSA [87 –89]. Additionally, military personnel, prisoners, and athletes exhibit larger numbers of SSTIs caused by CA-MRSA [76,89 –95]. In each of these high-risk groups, the risk factors for infection are close contact, compromised skin integrity, and an increased prevalence of colonization. Furthermore, outbreaks often are linked to periods of more common colonization or exposure to specific strains of S. aureus, such as USA300 CA-MRSA [96].

Multi-resistant and opportunistic pathogens

Overall, antibiotic-resistant pathogens other than MRSA, namely, methicillin-resistant CoNS, vancomycin-resistant enterococci, and extended spectrum beta-lactamase-producing gram-negative bacteria, have been associated increasingly with healthcare-acquired infections [97, 98]. Moreover, there are various other bacterial pathogens known to be opportunistic infectious agents, especially in orthopedics.

Pseudomonas aeruginosa typically is present on the skin (Proteobacteria). The organism is most commonly associated with infections in immunocompromised individuals and burn victims and often is multi-drug resistant [99]. Although P. aeruginosa typically is not enteric, it has been identified recently as a gastrointestinal source of blood stream infections after hematopoietic stem-cell transplantation [100].

Serratia is another gram-negative opportunistic pathogen of the Enterobacteriaceae family that typically is spread by hand-to-hand contact. Serratia marcescens is the most pathogenic species within the genus and a frequent cause of healthcare-acquired infections, particularly after spinal surgery. The bacterium is prevalent in the environment and causes a wide range of healthcare-acquired infections, such as respiratory tract infections, osteomyelitis, urinary tract infections, and endocarditis [101 –105].

Enterococcal species, such as vancomycin-resistant Enterococcus faecalis (VRE) and Enterococcus faecium, are gut bacteria that frequently colonize orthopedic and soft tissues and typically spread through direct contact. Enterococci have become the second most common organisms recovered from nosocomial urinary tract and wound infections and one of the most common causes of nosocomial bacteremia in the United States [106].

Escherichia coli also accounts for many soft tissue infections. This organism has been identified as a causative agent of many conditions, including, but not limited to, cellulitis localized to the lower or upper limbs, necrotizing fasciitis, surgical site infections, and infection after burn injuries [107 –112].

Fungal pathogens

Several common fungal pathogens are opportunistic infectious agents.

Candida albicans is a commensal fungus that resides mainly on human mucosal surfaces. It is a known opportunistic pathogen in immunocompromised individuals and commonly is associated with biofilm formation, making it difficult to eradicate [113]. It is the third leading cause of infections associated with intravascular catheters, having the second highest rate of colonization transitioning to infection and the highest overall crude mortality rate [114,115]. Notably, C. albicans can colonize prosthetic devices either endogenously or exogenously. Other Candida species notable for healthcare-acquired infections are C. glabrata, C. parapsilosis, C. krusei, and C. tropicalis [116 –120].

Cryptococcus neoformans is an encapsulated fungus that causes life-threatening meningoencephalitis in immunocompromised individuals. It colonizes and subsequently forms biofilms on various prosthetic devices, including ventricular shunts, peritoneal dialysis fistulas, and cardiac valves [121 –124].

Although normally it is a benign fungus, some species of Trichosporon can disseminate in immunocompromised hosts, leading to systemic disease. Trichosporon has been associated with infections related to devices such as catheters, breast implants, and cardiac grafts [125 –127].

Lastly, various fungi have been isolated as opportunistic agents in specific situations. Blastoschizomyces capitatus has been associated with catheter-related fungemia, and M. pachydermatis has been isolated from infected patients receiving parenteral nutrition. The baker's yeast Saccharomyces cerevisiae has been found in association with the dentures of patients with stomatitis, and persistent meningitis has been linked to a C. immitis biofilm at the tip of ventriculoperitoneal shunt tubing [117,128 –130]. Aspergillus fumigatus has caused infections of medical implant devices, including cardiac pacemakers, joint prostheses, and breast implants [131,132].

Conclusion

During surgical interventions, understanding which species of flora-bacteria are already present on the skin and mucosal tissues at various anatomic sites can help prepare providers for post-operative surgical infections. Furthermore, this knowledge allows surgeons to implement more specific and effective preventative measures in the pre-operative stages. Orthopedic-specific operations make up a significant number of all surgical procedures, and these numbers are expected to rise considerably over the next several years. Better recognizing which pathogens are typically found at specific anatomic sites and during specific procedures may improve surgical and medical interventions for peri-operative infections.

Footnotes

Acknowledgements

We thank Diane N. Weiss and the Sipprelle Family Foundation for their continued support.

Author Disclosure Statement

Dioscaris R. Garcia holds equity in BI Medical, LLC. Adam E. M. Eltorai, MSc receives royalties from LWW and Springer. John D. Jarrell holds equity in and is a Board member of Biointraface, Inc. and holds equity in and is a consultant for BI Medical, LLC. Christopher T. Born holds equity in and is a board member of Biointraface, Inc. and holds equity in and is a consultant for BI Medical, LLC and Illuminoss, LLC. David Deckely, Carole SL Spake, Jack M. Haglin, Toby Emanuel, and Cory Mayfield have no disclosures.

References

Grice

, Kong

, Renaud

, et al. A diversity profile of the human skin microbiota. Genome Res, 2008; 18:1043–1050.

Clarridge

. Impact of 16s rRNA gene sequence analysis for identification of bacteria on clinical microbiology and infectious diseases. Clin Microbiol Rev, 2004; 17:840–862.

Fierer

, Hamady

, Lauber

, Knight

. The influence of sex, handedness, and washing on the diversity of hand surface bacteria. Proc Natl Acad Sci USA, 2008; 105:17994–17999.

Wertheim

, Melles

, Vos

, et al. The role of nasal carriage in Staphylococcus aureus infections. Lancet Infect Dis, 2005; 5:751–762.

Faden

, Lesse

, Trask

, et al. Importance of colonization site in the current epidemic of staphylococcal skin abscesses. Pediatrics, 2010; 125:e618–e624.

Fritz

, Hogan

, Hayek

, et al. Staphylococcus aureus colonization in children with community-associated Staphylococcus aureus skin infections and their household contacts. Arch Pediatr Adolesc Med, 2012; 166:551–557.

Kluytmans

, Van Belkum

, Verbrugh

. Nasal carriage of Staphylococcus aureus: Epidemiology, underlying mechanisms, and associated risks. Clin Microbiol Rev, 1997; 10:505–520.

Mertz

, Frei

, Jaussi

, et al. Throat swabs are necessary to reliably detect carriers of Staphylococcus aureus. Clin Infect Dis, 2007; 45:475–477.

Weems

Jr . The many faces of Staphylococcus aureus infection: Recognizing and managing its life-threatening manifestations. Postgrad Med, 2001; 110:24–36.

10.

Lindsay

, Holden

. Staphylococcus aureus: Superbug, super genome?. Trends Microbiol, 2004; 12:378–385.

11.

Klevens

, Morrison

, Nadle

, et al. Invasive methicillin-resistant Staphylococcus aureus infections in the United States. JAMA, 2007; 298:1763–1771.

12.

Becker

, Heilmann

, Peters

. Coagulase-negative staphylococci. Clin Microbiol Rev, 2014; 27:870–926.

13.

Grice

, Segre

. The skin microbiome. Nat Rev Microbiol, 2011; 9:244.

14.

Grice

, Kong

, Conlan

, et al. Topographical and temporal diversity of the human skin microbiome. Science, 2009; 324:1190–1192.

15.

Costello

, Lauber

, Hamady

, et al. Bacterial community variation in human body habitats across space and time. Science, 2009; 326:1694–1697.

16.

Weigelt

, Lipsky

, Tabak

, et al. Surgical site infections: Causative pathogens and associated outcomes. Am J Infect Control, 2010; 38:112–1208.

17.

Lundberg

, Smith

, Heaney

, et al. Pre-operative antisepsis protocol compliance and the effect on bacterial load reduction. Surg Infect, 2016; 17:32–37.

18.

Barie

, Eachempati

. Surgical site infections. Surg Clin North Am, 2005; 85:1115–1135.

19.

Bennett-Guerrero

, Pappas

, Koltun

, et al. Gentamicin–collagen sponge for infection prophylaxis in colorectal surgery. N Engl J Med, 2010; 363:1038–1049.

20.

Mangram

, Horan

, Pearson

, et al. Guideline for prevention of surgical site infection, 1999. Am J Infect Control, 1999; 27:97–134.

21.

Moet

, Jones

, Biedenbach

, et al. Contemporary causes of skin and soft tissue infections in North America, Latin America, and Europe: Report from the SENTRY Antimicrobial Surveillance Program (1998–2004). Diagn Microbiol Infect Dis, 2007; 57:7–13.

22.

, Rotstein

. Bacterial skin and soft tissue infections in adults: A review of their epidemiology, pathogenesis, diagnosis, treatment and site of care. Can J Infect Dis Med Microbiol, 2008; 19:173–184.

23.

Dancer

, Stewart

, Coulombe

, et al. Surgical site infections linked to contaminated surgical instruments. J Hosp Infect, 2012; 81:231–238.

24.

Saito

, Kobayashi

, Uetera

, et al. Microbial contamination of surgical instruments used for laparotomy. Am J Infect Control, 2014; 42:43–47.

25.

Dryden

. Skin and soft tissue infection: Microbiology and epidemiology. Int J Antimicrob Agents, 2009; 34:S2–S7.

26.

Rennie

, Jones

, Mutnick

. Occurrence and antimicrobial susceptibility patterns of pathogens isolated from skin and soft tissue infections: Report from the SENTRY Antimicrobial Surveillance Program (United States and Canada, 2000). Diagn Microbiol Infect Dis, 2003; 45:287–293.

27.

King

, Humphrey

, Wang

, et al. Emergence of community-acquired methicillin-resistant Staphylococcus aureus USA 300 clone as the predominant cause of skin and soft-tissue infections. Ann Intern Med, 2006; 144:309–317.

28.

Frazee

, Lynn

, Charlebois

, et al. High prevalence of methicillin-resistant Staphylococcus aureus in emergency department skin and soft tissue infections. Ann Emerg Med, 2005; 45:311–320.

29.

Eady

, Cove

. Staphylococcal resistance revisited: Community-acquired methicillin-resistant Staphylococcus aureus: An emerging problem for the management of skin and soft tissue infections. Curr Opin Infect Dis, 2003; 16:103–124.

30.

Molina

, Thakore

, Blumer

, et al. Use of the National Surgical Quality Improvement Program in orthopaedic surgery. Clin Orthopaed Rel Res, 2015; 473:1574–1581.

31.

American Academy of Orthopaedic Surgeons. Orthopaedic Practice in the U.S. 2014. AAOS. Rosemont, IL. 2015

32.

U.S. Centers for Disease Control and Prevention/National Center for Health Statistics. National Hospital Discharge Survey. Available at: https://www.cdc.gov/nchs/data/nhds/4procedures/2010pro4_numberprocedureage.pdf Accessed June 21, 2016.

33.

Kurtz

, Ong

, Lau

, et al. Projections of primary and revision hip and knee arthroplasty in the United States from 2005 to 2030. J Bone Joint Surg, 2007; 89:780–785.

34.

Mears

, Edwards

. Bone and joint infections in older adults. Clin Geriatr Med, 2016; 32:555–570.

35.

Abdul-Jabbar

, Berven

, Hu

, et al. Surgical site infections in spine surgery: Identification of microbiologic and surgical characteristics in 239 cases. Spine, 2013; 38:E1425–E1431.

36.

Claessen

, Braun

, van Leeuwen

, et al. What factors are associated with a surgical site infection after operative treatment of an elbow fracture?. Clin Orthop Rel Res, 2016; 474:562–570.

37.

Fowler

, Ilyas

. Epidemiology of adult acute hand infections at an urban medical center. J Hand Surg, 2013; 38:1189–1193.

38.

Zalavras

, Christensen

, Rigopoulos

, et al. Infection following operative treatment of ankle fractures. Clin Orthop Rel Res, 2009; 467:1715–2720.

39.

Chahoud

, Kanafani

, Kanj

. Surgical site infections following spine surgery: Eliminating the controversies in the diagnosis. Front Med, 2014; 1:7.

40.

Sanchez-Sotelo

. Total elbow arthroplasty. Open Orthop J, 2011; 5:115.

41.

Platt

, Page

. Post-operative infection following hand surgery: Guidelines for antibiotic use. J Hand Surg, 1995; 20:685–690.

42.

Marsh

, Saltzman

. Ankle fractures. In: Bucholz

, Heckman

, Court-Brown

, eds. Rockwood and Green's Fractures in Adults. Sixth Edition. JB Lippincott; Philadelphia. 2005:2148–2247.

43.

Rafiq

, Gambhir

, Wroblewski

, Kay

. The microbiology of infected hip arthroplasty. Int Orthop, 2006; 30:532–535

44.

Nickinson

, Board

, Gambhir

, et al. The microbiology of the infected knee arthroplasty. Int Orthop, 2010; 34:505–510.

45.

Moran

, Byren

, Atkins

. The diagnosis and management of prosthetic joint infections. J Antimicrob Chemother, 2010; 65(suppl_3):iii45–iii54.

46.

Blom

, Brown

, Taylor

, et al. Infection after total knee arthroplasty. Bone Joint J, 2004; 86:688–691.

47.

Noud

, Esch

. Complications of arthroscopic shoulder surgery. Sports Med Arthrosc Rev, 2013; 21:89–96.

48.

Hsu

, Bumgarner

, Matsen III

. Propionibacterium in shoulder arthroplasty: What we think we know today. J Bone Joint Surg, 2016; 98:597–606.

49.

Richards

, Inacio

, Beckett

, et al. Patient and procedure-specific risk factors for deep infection after primary shoulder arthroplasty. Clin Orthop Rel Res, 2014; 472:2809–2815.

50.

Hackett

Jr , Crosby

. Infection prevention in shoulder surgery. Bull NYU Hosp Joint Dis, 2015; 73(1_suppl):S140–S141.

51.

Saltzman

, Nuber

, Gryzlo

, et al. Efficacy of surgical preparation solutions in shoulder surgery. J Bone Joint Surg, 2009; 91:1949–1953.

52.

Ostrander

, Botte

, Brage

. Efficacy of surgical preparation solutions in foot and ankle surgery. J Bone Joint Surg, 2005; 87:980–985.

53.

, Fowler

, Goitz

. Prospective randomized trial comparing the efficacy of surgical preparation solutions in hand surgery. Hand, 2017; 12:258–264.

54.

Savage

, Weatherford

, Sugrue

, et al. Efficacy of surgical preparation solutions in lumbar spine surgery. J Bone Joint Surg, 2012; 94:490–494.

55.

Lee

, Pottinger

, Butler-Wu

, et al. Propionibacterium persists in the skin despite standard surgical preparation. J Bone Joint Surg, 2014; 96:1447–1450.

56.

Eiselt

. Presurgical skin preparation with a novel 2% chlorhexidine gluconate cloth reduces rates of surgical site infection in orthopaedic surgical patients. Orthop Nursing, 2009; 28:141–145.

57.

Johnson

, Daley

, Zywiel

, et al. Preoperative chlorhexidine preparation and the incidence of surgical site infections after hip arthroplasty. J Arthrop, 2010:25:98–102.

58.

Kapadia

, Johnson

, Daley

et al. Pre-admission cutaneous chlorhexidine preparation reduces surgical site infections in total hip arthroplasty. J Arthrop, 2013; 28:490–493.

59.

Howard

, Scott

, Packard

, Jones

. The global impact of drug resistance. Clin Infect Dis, 2003; 36(Supplement 1):S4–S10.

60.

Arciola

, Campoccia

, Ravaioli

, Montanaro

. Polysaccharide intercellular adhesin in biofilm: Structural and regulatory aspects. Front Cellular Infect Microbiol, 2015; 5:7.

61.

National Nosocomial Infections Surveillance (NNIS) System Report. Data summary from January 1992 through June 2004, issued October 2004. Am J Infect Control, 2004; 32:470–485.

62.

Kuehnert

, Hill

, Kupronis

, et al. Methicillin-resistant–Staphylococcus aureus hospitalizations, United States. Emerg Infect Dis, 2005; 11:468.

63.

Fridkin

, Hageman

, Morrison

, et al. Methicillin-resistant Staphylococcus aureus disease in three communities. N Engl J Med, 2005; 352:1436–1444.

64.

Miller

, Remington

, Bayer

, et al. Clinical and epidemiologic characteristics cannot distinguish community-associated methicillin-resistant Staphylococcus aureus infection from methicillin-susceptible S. aureus infection: A prospective investigation. Clin Infect Dis, 2007; 44:471–482.

65.

Dryden

. Complicated skin and soft tissue infections caused by methicillin-resistant Staphylococcus aureus: Epidemiology, risk factors, and presentation. Surg Infect, 2008; 9(suppl 1):s3–s10.

66.

Napolitano

. Early appropriate parenteral antimicrobial treatment of complicated skin and soft tissue infections caused by methicillin-resistant Staphylococcus aureus. Surg Infect, 2008; 9(suppl 1):s17–s27.

67.

Otter

, French

. Nosocomial transmission of community-associated methicillin-resistant Staphylococcus aureus: An emerging threat. Lancet Infect Dis, 2006; 6:753–755.

68.

Eveillard

, de Lassence

, Lancien

, et al. Evaluation of a strategy of screening multiple anatomical sites for methicillin-resistant Staphylococcus aureus at admission to a teaching hospital. Infect Control Hosp Epidemiol, 2006; 27:181–184.

69.

Nilsson

, Ripa

. Staphylococcus aureus throat colonization is more frequent than colonization in the anterior nares. J Clin Microbiol, 2006; 44:3334–3339.

70.

Peters

, Brooks

, Limbago

, Lowery

, et al. Methicillin-resistant Staphylococcus aureus colonization in HIV-infected outpatients is common and detection is enhanced by groin culture. Epidemiol Infect, 2011; 139:998–1008.

71.

Kaplan

, Forbes

, Hammerman

, et al. Randomized trial of “bleach baths” plus routine hygienic measures vs routine hygienic measures alone for prevention of recurrent infections. Clin Infect Dis, 2013; 58:679–682.

72.

Gorwitz

, Kruszon-Moran

, McAllister

, et al. Changes in the prevalence of nasal colonization with Staphylococcus aureus in the United States, 2001–2004. J Infect Dis, 2008; 197:1226–1234.

73.

Von Eiff

, Becker

, Machka

, et al. Nasal carriage as a source of Staphylococcus aureus bacteremia. N Engl J Med, 2001; 344:11–16.

74.

Toshkova

, Annemüller

, Akineden

, Lämmler

. The significance of nasal carriage of Staphylococcus aureus as risk factor for human skin infections. FEMS Microbiol Lett, 2001; 202:17–24.

75.

Davis

, Stewart

, Crouch

, et al. Methicillin-resistant Staphylococcus aureus (MRSA) nares colonization at hospital admission and its effect on subsequent MRSA infection. Clin Infect Dis, 2004; 39:776–782.

76.

Ellis

, Hospenthal

, Dooley

, et al. Natural history of community-acquired methicillin-resistant Staphylococcus aureus colonization and infection in soldiers. Clin Infect Dis, 2004; 39:971–979.

77.

Fritz

, Epplin

, Garbutt

, Storch

. Skin infection in children colonized with community-associated methicillin-resistant Staphylococcus aureus. J Infect, 2009; 59:394–401.

78.

Thompson

, Cabezudo

, Wenzel

. Epidemiology of nosocomial infections caused by methicillin-resistant Staphylococcus aureus. Ann Intern Med, 1982; 97:309–317.

79.

Boyce

. Methicillin-resistant Staphylococcus aureus: Detection, epidemiology, and control measures. Infect Dis Clin North Am, 1989; 3:901–913.

80.

Purcell

, Fergie

. Epidemic of community-acquired methicillin-resistant Staphylococcus aureus infections: A 14-year study at Driscoll Children's Hospital. Arch Pediatr Adolesc Med, 2005; 159:980–985.

81.

Kaplan

, Hulten

, Gonzalez

, et al. Three-year surveillance of community-acquired Staphylococcus aureus infections in children. Clin Infect Dis, 2005; 40:1785–1791.

82.

Gonzalez

, Martinez-Aguilar

, Hulten

, et al. Severe staphylococcal sepsis in adolescents in the era of community-acquired methicillin-resistant Staphylococcus aureus. Pediatrics, 2005; 115:642–648.

83.

Levison

, Fung

. Community-associated methicillin-resistant Staphylococcus aureus: Reconsideration of therapeutic options. Curr Infect Dis Rep, 2006; 8:23.

84.

Stankovic

, Mahajan

. Healthy children with invasive community-acquired methicillin-resistant Staphylococcus aureus infections. Pediatr Emerg Care, 2006; 22:361–363.

85.

Creech

, Johnson

, Bartilson

, et al. Increasing use of extracorporeal life support in methicillin-resistant Staphylococcus aureus sepsis in children. Pediatr Crit Care Med, 2007; 8:231–235.

86.

Gerber

, Coffin

, Smathers

, Zaoutis

. Trends in the incidence of methicillin-resistant Staphylococcus aureus infection in children's hospitals in the United States. Clin Infect Dis, 2009; 49:65–71.

87.

Ray

, Suaya

, Baxter

. Incidence, microbiology, and patient characteristics of skin and soft-tissue infections in a US population: A retrospective population-based study. BMC Infect Dis, 2013; 13:252.

88.

Ray

, Suaya

, Baxter

. Microbiology of skin and soft tissue infections in the age of community-acquired methicillin-resistant Staphylococcus aureus. Diagn Microbiol Infect Dis, 2013; 76:24–30.

89.

Kazakova

, Hageman

, Matava

, et al. A clone of methicillin-resistant Staphylococcus aureus among professional football players. N Engl J Med, 2005; 352:468–475.

90.

Jiménez-Truque

, Saye

, Soper

, et al. Longitudinal assessment of colonization with Staphylococcus aureus in healthy collegiate athletes. J Pediatr Infect Dis Soc, 2014; 5:105–113.

91.

Wright

M-O

, Furuno

, Venezia

, et al. Methicillin-resistant Staphylococcus aureus infection and colonization among hospitalized prisoners. Infect Control Hosp Epidemiol, 2007; 28:877–879.

92.

Creech

, Saye

, McKenna

, et al. One-year surveillance of methicillin-resistant Staphylococcus aureus nasal colonization and skin and soft tissue infections in collegiate athletes. Arch Pediatr Adolesc Med, 2010; 164:615–620.

93.

David

, Mennella

, Mansour

, et al. Predominance of methicillin-resistant Staphylococcus aureus among pathogens causing skin and soft tissue infections in a large urban jail: Risk factors and recurrence rates. J Clin Microbiol, 2008; 46:3222–3227.

94.

U.S. Centers for Disease Control and Prevention. Methicillin-resistant Staphylococcus aureus infections in correctional facilities—Georgia, California, and Texas, 2001–2003. MMWR Morbid Mortal Wkly Rep, 2003; 52:992.

95.

U.S. Centers for Disease Control and Prevention. Methicillin-resistant Staphylococcus aureus infections among competitive sports participants—Colorado, Indiana, Pennsylvania, and Los Angeles County, 2000–2003. MMWR Morbid Mortal Wkly Rep, 2003; 52:793.

96.

Carrel

, Perencevich

, David

. USA300 methicillin-resistant Staphylococcus aureus, United States, 2000–2013. Emerg Infect Dis, 2015; 21:1973.

97.

Krukerink

, Kievit

, Marang-van de Mheen

. Evaluation of routinely reported surgical site infections against microbiological culture results: A tool to identify patient groups where diagnosis and treatment may be improved. BMC Infect Dis, 2009; 9:176.

98.

Martinez-Pastor

, Vilchez

, Pitart

, et al. Antibiotic resistance in orthopaedic surgery: Acute knee prosthetic joint infections due to extended-spectrum beta-lactamase (ESBL)-producing Enterobacteriaceae. Eur J Clin Microbiol Infect Dis, 2010; 29:1039–1041.

99.

de Bentzmann

, Plésiat

. The Pseudomonas aeruginosa opportunistic pathogen and human infections. Environ Microbiol, 2011; 13:1655–1665.

100.

Tamburini

, Andermann

, Tkachenko

, et al. Precision identification of diverse bloodstream pathogens from the gut microbiome. bioRxiv, 2018:310441.

101.

Castelli

, Fedrigo

, Clementín

, et al. Enterobacterial common antigen integrity is a checkpoint for flagellar biogenesis in Serratia marcescens. J Bacteriol, 2008; 190:213–220.

102.

Hejazi

, Falkiner

. Serratia marcescens. J Med Microbiol, 1997; 46:903–912.

103.

Kida

, Inoue

, Shimizu

, Kuwano

. Serratia marcescens serralysin induces inflammatory responses through protease-activated receptor 2. Infect Immun, 2007; 75:164–174.

104.

Koh

, Lam

, Alhede

, et al. Phenotypic diversification and adaptation of Serratia marcescens MG1 biofilm-derived morphotypes. J Bacteriol, 2007; 189:119–130.

105.

Matsuo

, Chen

, Minato

, et al. SmdAB, a heterodimeric ABC-type multidrug efflux pump, in Serratia marcescens. J Bacteriol, 2008; 190:648–654.

106.

Schaberg

, Culver

, Gaynes

. Major trends in the microbial etiology of nosocomial infection. Am J Med, 1991; 91:S72–S75.

107.

Afifi

, El-Hindawi

. Acute necrotizing fasciitis in Egyptian patients: A case series. Int J Surg, 2008; 6:7–14.

108.

Corredoira

, Ariza

, Pallares

, et al. Gram-negative bacillary cellulitis in patients with hepatic cirrhosis. Eur J Clin Microbiol Infect Dis, 1994; 13:19–24.

109.

, De Lun

, Chen

. Necrotising fasciitis with Escherichia coli. Lancet Infect Dis, 2006; 6:456.

110.

Krebs

VLJ

, Koga

, Diniz

EMdA

, et al. Necrotizing fasciitis in a newborn infant: A case report. Revis Hosp Clín, 2001; 56:59–62.

111.

Brzozowski

, Ross

. Upper limb Escherichia coli cellulitis in the immunocompromised. J Hand Surg, 1997; 22:679–680.

112.

Tourmousoglou

, Yiannakopoulou

, Kalapothaki

, et al. Surgical-site infection surveillance in general surgery: A critical issue. J Chemother, 2008; 20:312–318.

113.

Ramage

, Martínez

, López‐Ribot

. Candida biofilms on implanted biomaterials: A clinically significant problem. FEMS Yeast Res, 2006; 6:979–986.

114.

Crump

, Collignon

. Intravascular catheter-associated infections. Eur J Clin Microbiol Infect Dis, 2000; 19:1–8.

115.

Wisplinghoff

, Bischoff

, Tallent

, et al. Nosocomial bloodstream infections in US hospitals: Analysis of 24,179 cases from a prospective nationwide surveillance study. Clin Infect Dis, 2004; 39:309–317.

116.

Hawser

, Douglas

. Biofilm formation by Candida species on the surface of catheter materials in vitro. Infect Immun, 1994; 62:915–921.

117.

Coco

, Bagg

, Cross

, et al. Mixed Candida albicans and Candida glabrata populations associated with the pathogenesis of denture stomatitis. Mol Oral Microbiol, 2008; 23:377–383.

118.

Choi

, Shin

, Jung

, et al. Species-specific differences in the susceptibilities of biofilms formed by Candida bloodstream isolates to echinocandin antifungals. Antimicrob Agents Chemother, 2007; 51:1520–1523.

119.

Shin

, Kee

, Shin

, et al. Biofilm production by isolates of Candida species recovered from nonneutropenic patients: Comparison of bloodstream isolates with isolates from other sources. J Clin Microbiol, 2002; 40:1244–1248.

120.

Tumbarello

, Posteraro

, Trecarichi

, et al. Biofilm production by Candida species and inadequate antifungal therapy as predictors of mortality for patients with candidemia. J Clin Microbiol, 2007; 45:1843–1850.

121.

Walsh

, Schlegel

, Moody

, et al. Ventriculoatrial shunt infection due to Cryptococcus neoformans: An ultrastructural and quantitative microbiological study. Neurosurgery, 1986; 18:376–382.

122.

Bach

, Tally

, Godofsky

. Use of cerebrospinal fluid shunts in patients having acquired immunodeficiency syndrome with cryptococcal meningitis and uncontrollable intracranial hypertension. Neurosurgery, 1997; 41:1280–1283.

123.

Banerjee

, Gupta

, Venugopal

. A case of prosthetic valve endocarditis caused by Cryptococcus neoformans var. neoformans. J Med Vet Mycol, 1997; 35:139–141.

124.

Braun

, Janssen

, Marcus

, Kauffman

. Cryptococcal infection of a prosthetic dialysis fistula. Am J Kidney Dis, 1994; 24:864–867.

125.

Krzossok

, Birck

, Henke

, et al. Trichosporon asahii infection of a dialysis PTFE arteriovenous graft. Clin Nephrol, 2004; 62:66–68.

126.

Pini

, Faggi

, Donato

, Fanci

. Isolation of Trichosporon in a hematology ward. Mycoses, 2005; 48:45–49.

127.

Reddy

, Torres

, Kontoyiannis

. Breast implant infection caused by Trichosporon beigelii. Scand J Infect Dis, 2002; 34:143–144.

128.

Cannizzo

, Eraso

, Ezkurra

, et al. Biofilm development by clinical isolates of Malassezia pachydermatis. Med Mycol, 2007; 45:357–361.

129.

D'Antonio

, Parruti

, Pontieri

, et al. Slime production by clinical isolates of Blastoschizomyces capitatus from patients with hematological malignancies and catheter-related fungemia. Eur J Clin Microbiol Infect Dis, 2004; 23:787–789.

130.

Davis

, Cook

, Costerton

. Biofilm on ventriculo-peritoneal shunt tubing as a cause of treatment failure in coccidioidal meningitis. Emerg Infect Dis, 2002; 8:376.

131.

Langer

, Kassim

, Macari

, Saleh

. Aspergillus infection after total knee arthroplasty. Am J Orthop, 2003; 32:402–404.

132.

Rosenblatt

, Pollock

. Aspergillus flavus cultured from a saline-filled implant. Plast Reconstruct Surg, 1997; 99:1470–1472.