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Rapidly growing mycobacterial infections: Mycobacteria abscessus, chelonae, and fortuitum

Rapidly growing mycobacterial infections: Mycobacteria abscessus, chelonae, and fortuitum
Literature review current through: Sep 2023.
This topic last updated: Jul 17, 2023.

INTRODUCTION — Rapidly growing mycobacteria (RGM) are environmental organisms that are found worldwide and usually grow in subculture within one week (ie, more rapidly than other mycobacteria) (table 1). The three most common clinically relevant species are:

Mycobacterium abscessus

Mycobacterium fortuitum

Mycobacterium chelonae

Among this group, M. abscessus is the species most commonly isolated from clinical respiratory specimens; M. fortuitum and M. chelonae are more commonly isolated from non-respiratory specimens.

Appropriate management of RGM depends on awareness of the variable susceptibility patterns of the different species and subspecies and the potential diagnostic limitations in distinguishing them.

The disease associations, diagnosis, and treatment of RGM infections will be discussed here.

The microbiology of nontuberculous mycobacterial (NTM) infections, the diagnosis of NTM pulmonary disease, and bone and joint NTM infections are discussed in detail elsewhere:

(See "Overview of nontuberculous mycobacterial infections" and "Microbiology of nontuberculous mycobacteria".)

(See "Diagnosis of nontuberculous mycobacterial infections of the lungs".)

(See "Epidemiology, clinical manifestations, and diagnosis of osteomyelitis due to nontuberculous mycobacteria" and "Treatment of osteomyelitis due to nontuberculous mycobacteria in adults".)

EPIDEMIOLOGY — The epidemiology of nontuberculous mycobacterial (NTM) infections, including those caused by RGM, is discussed separately. (See "Epidemiology of nontuberculous mycobacterial infections".)

NOMENCLATURE FOR M. ABSCESSUS — The taxonomy and nomenclature for M. abscessus have been in flux; there are now three proposed M. abscessus subspecies:

M. abscessus subspecies abscessus (the organism traditionally labeled "M. abscessus")

M. abscessus subspecies bolletii

M. abscessus subspecies massiliense

When possible, it is important to distinguish clinical M. abscessus isolates to the subspecies level because of treatment implications: the majority of M. abscessus subspecies abscessus and M. abscessus subspecies bolletii isolates, with occasional exceptions, have an active inducible macrolide resistance (erm) gene, whereas M. abscessus subspecies massiliense does not (with rare exceptions). This difference impacts treatment decisions regarding the use of macrolide-containing regimens. (See 'Detecting macrolide resistance' below.)

However, these subspecies are difficult to distinguish in the clinical laboratory, and so subspecies designation is frequently unavailable (see 'Mycobacterial species identification' below). Further confusing the subspecies designation is that some investigators had historically labeled M. abscessus subspecies massiliense as M. bolletii (or included it as part of M. abscessus subspecies bolletii).

The nomenclature detailed above is consistent with a proposal from an international group (including the author of this topic) to designate these three subspecies based on genomic data and phenotypic data regarding the erm gene [1]. We use this nomenclature throughout this topic. If data on M. abscessus are reported without a subspecies designation, it is because no subspecies information is available.

CLINICAL MANIFESTATIONS

Spectrum of disease by species — The clinical disease spectra caused by M. abscessus, M. fortuitum, and M. chelonae vary:

M. abscessus is the most pathogenic of the RGM group and the RGM most likely to cause pulmonary infection, particularly in patients with underlying lung disease such as cystic fibrosis-related and non-cystic fibrosis-related bronchiectasis.

M. fortuitum causes human infection primarily by direct inoculation (eg, through trauma or surgery), resulting in primary skin and soft tissue infections [2], surgical wound infections [3], and catheter-related sepsis [4]. Rarely, it causes keratitis [5], prosthetic valve endocarditis [6], cervical lymphadenitis [6], and pulmonary disease [4]. However, the majority of M. fortuitum respiratory isolates are from individuals with underlying pulmonary diseases, such as bronchiectasis, and represent colonization or transient infection that does not require treatment [7]; pulmonary disease is also associated with underlying gastrointestinal disorders, such as achalasia [8].

M. chelonae primarily causes human infection in immunosuppressed patients, including hematogenously disseminated disease. M. chelonae may also cause surgical wound infections and keratitis [5].

Pulmonary infection — Pulmonary disease due to RGM is predominantly due to M. abscessus [4,9]. In one report that reviewed the epidemiology and clinical manifestations of 154 patients with RGM pulmonary disease (82 percent of whom had M. abscessus) at two centers in Texas [9]:

The median age at onset was 58 years (range 1 to 85 years).

The majority were female (65 percent).

Specific underlying diseases were documented in the minority: previously treated mycobacterial disease (18 percent), cystic fibrosis (8 percent), current or former smoking (34 percent).

The diagnosis was usually not established until more than two years after the onset of symptoms.

Cough was the most frequent symptom at presentation (71 percent); fever, weight loss, hemoptysis, and dyspnea were also common.

More than half had involvement of three or more lobes on chest radiograph. An interstitial pattern, a mixed interstitial and alveolar infiltrative pattern, and a reticulonodular pattern were each seen with similar frequency (36 to 40 percent). Cavitation was infrequent (16 percent). These findings are similar to those seen with nodular/bronchiectatic (noncavitary) Mycobacterium avium complex (MAC) lung disease. (See "Diagnosis of nontuberculous mycobacterial infections of the lungs", section on 'Chest imaging'.)

Death was attributed to progressive RGM lung disease with respiratory failure in 21 cases (14 percent). The mean time from diagnosis to death was shorter among those with severe underlying lung disease (1 versus 7.8 years); however, there were minimal data about disease course prior to death.

Other case series have highlighted the association between RGM pulmonary infection and pre-existing lung disease, particularly bronchiectasis [10-12] as well as underlying conditions such as esophageal disease [13], malignancy [14,15], and rheumatologic conditions [10]. As an example, in a review of RGM infections in 341 patients with cancer, the most common type of infection was pulmonary (47 percent of cases) [15]. Underlying lung disease was documented in 70 percent.

Similar to MAC lung disease, clinical features associated with progressive RGM lung disease include low body mass index (BMI), bilateral lung involvement, and fibrocavitary disease [16]. Certain genetic characteristics of a particular RGM isolate may also affect the natural history of pulmonary infection [17].

Lymphadenitis — M. abscessus and M. fortuitum are occasional causes of superficial lymphadenitis, especially cervical lymphadenitis, in children. Most cases of nontuberculous lymphadenitis are due to M. avium complex organisms. (See "Nontuberculous mycobacterial lymphadenitis in children", section on 'Microbiology'.)

Disseminated disease — Disseminated RGM infection occurs most commonly in immunocompromised patients, including patients on glucocorticoids, and can present as multiple subcutaneous nodules (pseudo-erythema nodosum) or abscesses that drain spontaneously [2]. The nodules associated with disseminated RGM are often purple and not associated with skin penetration, and it is important to maintain a low threshold of suspicion for disseminated disease when these are present.

In a review of patients with cancer and RGM infections, 26 (8 percent) had disseminated disease [15]. The majority of those with disseminated disease had a hematologic malignancy (83 percent) or had undergone recent chemotherapy (87 percent). The most common species involved were M. abscessus (46 percent), M. chelonae (27 percent), and M. fortuitum (23 percent). Among the 22 patients in whom the outcome was reported, 15 (68 percent) died; RGM infection was a cause or contributing factor in half of the deaths.

Skin and soft tissue infection — RGM are an uncommon cause of soft tissue infections, but the incidence is increasing (possibly because of enhanced detection) [18]. Signs and symptoms include nodules (frequently with purple discoloration), recurrent abscesses, or chronic discharging sinuses. Cutaneous nodules are also associated with disseminated RGM disease [2]; in particular, the possibility of disseminated disease should be considered in immunocompromised patients (including those on glucocorticoids) with cutaneous RGM (see 'Disseminated disease' above). A high index of suspicion is necessary for diagnosis and timely collection of specimens for acid-fast bacilli (AFB) analysis.

In a retrospective case series of 63 patients with skin and soft tissue infections due to RGM [19]:

M. fortuitum infections were more likely to present as a single lesion (89 percent).

M. chelonae and M. abscessus were more likely to present as multiple lesions (62 percent).

Patients with multiple lesions were more likely to be immunosuppressed (67 percent; and many of these patients may have had disseminated disease)

Severe RGM skin and soft tissue infections have been associated with exposure to nail salon whirlpool footbaths [20-23]. Suboptimal cleaning of footbaths appears to increase the risk of infection [23]. An outbreak of furunculosis due to M. fortuitum was reported in patients using footbaths in a pedicure salon; the organism with the same pulsed field gel electrophoretic pattern observed in the patients was isolated from the footbaths [21]. Over 100 pedicure customers had prolonged boils on the lower legs that left scars when healed.

RGM, in particular M. chelonae and M. abscessus, have also been associated with skin infections after tattooing [24-30]. In case series, infection presented as an erythematous, papular rash along the inked areas of the skin that developed one to three weeks following receipt of the tattoo [27-29,31].

Musculoskeletal infection — Infection of the musculoskeletal system with RGM usually involves tenosynovitis and occurs from either percutaneous inoculation (eg, trauma or surgery) or hematogenous seeding. The clinical course is indolent, slowly progressive, and destructive, in part because of a delay in diagnosis [32]. The clinical manifestations of osteomyelitis due to RGM are discussed elsewhere. (See "Epidemiology, clinical manifestations, and diagnosis of osteomyelitis due to nontuberculous mycobacteria", section on 'Clinical findings'.)

Surgical site infection — Postoperative infections with RGM have occurred following various procedures including cosmetic surgery procedures (such as augmentation mammoplasty), laser in situ keratomileusis (LASIK), and heart surgery [33-41]. Nonsterile water (or ice made from nonsterile water) is a frequent source of RGM in health care-associated infections. Avoidance of nonsterile tap water for surgical procedures or other interventions is the single most important approach to preventing such infections.

Infection is characterized by multiple recurrent abscesses around the surgical wound. Microbiologic diagnosis is often delayed because routine cultures are not held long enough for mycobacterial growth.

Illustrative cases include:

M. abscessus was responsible for an outbreak of wound infections among 20 individuals who underwent abdominoplasty in the Dominican Republic [42]. This outbreak highlighted the risks of infection associated with surgical procedures performed in settings with inadequate infection control practices.

An epidemic of surgical site infections occurred following video-assisted surgeries in Rio de Janeiro, Brazil [43]. Of 1051 possible cases at 63 hospitals, 302 were attributed to M. massiliense [44]. A single clone of M. massiliense was detected in 74 of the 148 isolates that were sequenced. All of five isolates that were tested were tolerant to the glutaraldehyde solution used to sterilize surgical instruments, and the epidemic ended following discontinuation of glutaraldehyde use.

Sixteen patients under the care of a single clinician developed RGM skin infection at the site of mesotherapy injections [34]; most were caused by M. chelonae. Mesotherapy involves the subcutaneous injection of small quantities of substances, such as vitamins, silica, or lecithin, for the purpose of fat or wrinkle reduction [45].

Catheter-related infections — M. fortuitum and M. chelonae are an uncommon cause of catheter-related bacteremia in patients with cancer [46]. RGM have also been reported as a cause of infection complicating long-term peritoneal dialysis catheters [47,48]. In review of patients with cancer and RGM infections, catheter-associated bloodstream infections occurred most commonly in patients with hematologic malignancies (58 percent of cases) [15]. Mycobacterium mucogenicum was the most common species (30 percent), followed by M. fortuitum (22 percent), and M. chelonae (15 percent). The high rate of M. mucogenicum infection compared with other studies is most likely due to the fact that three of the studies included in this review involved outbreaks of infection caused by M. mucogenicum.

Prosthetic device infection — Infections related to prosthetic devices have been rarely described with tympanostomy tubes, pacemakers, and prosthetic joints [49-51]. As an example, a case series identified 21 cases of chronic M. abscessus otitis media after tympanostomy tube placement; the authors noted that mycobacteria should be considered as a cause of refractory post-tympanostomy tube otorrhea [49]. (See "Tympanostomy tube otorrhea in children: Causes, prevention, and management".)

Another case series with a review of the literature identified 18 cases of prosthetic joint infection, which included the knee, hip, and elbow [51]. The causative organisms were M. fortuitum (n = 10), M. chelonae (n = 6), Mycobacterium smegmatis (n = 1), and M. abscessus (n = 1).

DIAGNOSIS — Identifying an acid-fast bacillus (AFB) in a clinical specimen as an RGM and distinguishing it from Mycobacterium tuberculosis are important for several reasons.

M. tuberculosis requires public health tracking, whereas RGM (and other nontuberculous mycobacteria [NTM]) do not.

RGM are not susceptible in vitro to antituberculous drugs and warrant specific antimycobacterial therapy.

Recognizing the typical clinical settings for RGM infection and rapid molecular techniques for identifying M. tuberculosis facilitate the distinction.

Identifying the specific RGM species is also important, as treatment approaches vary by species. For organisms such as M. abscessus subspecies abscessus or bolletii or M. fortuitum, determination of erm gene activity with inducible macrolide resistance is a rapid alternative to early species identification.

When cultures are submitted, the lab should be alerted to the possibility of RGM, and special mycobacterial media should be used. (See 'Susceptibility testing' below.)

Approach

Pulmonary disease — The diagnosis of pulmonary disease caused by RGM is made in patients who have all of the following (table 2):

A compatible clinical syndrome (eg, cough, other pulmonary symptoms, fatigue, weight loss) with consistent radiographic findings

No other clear explanation for the clinical and radiographic findings

Microbiologic evidence that meets criteria for RGM infection

Minimal laboratory evaluation should include three or more sputum samples (either spontaneously expectorated or induced) for AFB and testing to exclude other confounding disorders, such as tuberculosis and lung malignancy. In most patients, a diagnosis can be made without a lung biopsy. Diagnostic criteria are primarily based on experience with M. avium complex infection and on expert consensus; these criteria have also been applied to other NTM species, including M. abscessus subspecies [52,53]. A notable exception is M. fortuitum, for which more rigorous diagnostic criteria should be applied. The diagnosis of NTM lung disease is discussed in detail elsewhere. (See "Diagnosis of nontuberculous mycobacterial infections of the lungs".)

The diagnosis of RGM pulmonary infection cannot be made by isolation of the organism on culture of respiratory specimens alone. As an example, M. abscessus respiratory isolates are sometimes recovered from patients undergoing treatment for M. avium complex lung disease, but these are not necessarily clinically significant [54]. Such isolates are more likely to be clinically significant and associated with progressive infection when they are recovered on multiple occasions and accompanied by new or enlarging cavitary lesions on chest radiograph. While the identification of certain genetic markers or virulence factors is a potential future tool for distinguishing clinically significant M. abscessus isolates, diagnosis continues to require careful clinical, microbiologic, and radiographic assessment and follow-up.

Nonpulmonary disease — Diagnosis of nonpulmonary RGM disease is made by culture of the specific pathogen from drainage material or biopsy of the affected site. The isolation of RGM from sterile, closed sites, such as bone marrow or blood or from a skin biopsy (in the setting of multiple lesions), is diagnostic of the disease.

In patients who have skin lesions due to RGM without obvious explanation (eg, no clear portal of entry or injury), the possibility of disseminated RGM (with blood and other closed site cultures) and underlying immunodeficiency should be evaluated. It is a common misperception that purple nodules that grow RGM (typically M. chelonae) on aspirate culture are due to localized disease, when these are more likely to be a manifestation of disseminated infection, particularly in immunocompromised patients [2].

Mycobacterial species identification — Once an isolate has been identified as an NTM, the species (and for M. abscessus, the subspecies) should be identified. Treatment recommendations rely on accurate laboratory identification of the specific RGM isolated. Accurate interpretation of studies of RGM also depends on the methods used for identifying RGM isolates; many older studies did not use adequate laboratory techniques to make accurate species and/or subspecies identification. (See 'Nomenclature for M. abscessus' above.)

However, identification of RGM in most laboratories is either incomplete or imprecise. M. abscessus isolates are sometimes identified simply as "M. chelonae/abscessus complex." If "M. abscessus" is specifically identified, sometimes the isolate is not accurately sub-speciated. Clinicians should be familiar with the capabilities of their clinical laboratories to speciate and report RGM (in particular, M. abscessus).

Only molecular methods, such as polymerase chain reaction (PCR)-restriction endonuclease assay (PRA), can reliably differentiate between M. abscessus and M. chelonae. Molecular identification can also be accomplished using gene sequencing, although this is only feasible for laboratories with access to sequencing facilities. Several target genes have been described (eg, 16S rRNA, hsp65, rpoB, and the 16S-23S internal transcribed spacer [ITS]) [53]. 16S rRNA gene sequencing alone offers limited discriminatory power, particularly for the M. abscessus-M. chelonae group. The hsp65 and rpoB genes and ITS are more discriminative. Complementing 16S rRNA sequencing with additional targets where required yields the best discriminatory power, allowing identifications up to subspecies level for M. abscessus [53]. (See "Microbiology of nontuberculous mycobacteria".)

Discriminatory power of the matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) mass spectrometry method for NTM has increased with recent improvements in protein extraction protocols and databases, but not all species and subspecies, including RGM, can be differentiated with this approach [53].

Widely available rapid methods for NTM identification in many laboratories include high-pressure liquid chromatography (HPLC) and commercial DNA probes, but neither is optimal for RGM. No commercial DNA probes are available for RGM. HPLC, which examines the mycolic acid fingerprint patterns that differ among most species of mycobacteria, cannot reliably differentiate between M. abscessus and M. chelonae.

Antibiotic susceptibility testing can also provide valuable clues to the species identification of RGM as each species has a relatively specific in vitro antibiotic susceptibility pattern. (See 'Agents to test' below.)

Susceptibility testing

Agents to test — Antimicrobial susceptibility varies among species of RGM. Susceptibility testing should be performed on all clinically significant isolates, including isolates that have been recovered after treatment failure or relapse.

The Clinical and Laboratory Standards Institute (CLSI) recommends susceptibility testing of RGM for clarithromycin (ie, macrolides), amikacin, imipenem, cefoxitin, linezolid, ciprofloxacin, moxifloxacin, doxycycline (or minocycline), trimethoprim-sulfamethoxazole, and, for M. chelonae only, tobramycin [55]. We also request susceptibility testing for tigecycline and clofazimine.

Additionally, all M. abscessus and M. fortuitum isolates must be tested for erm gene activity; this can be done by determining minimum inhibitory concentrations (MICs) for clarithromycin before and after extended (eg, 14 days) incubation with clarithromycin. (See 'Detecting macrolide resistance' below.)

Susceptibility patterns vary among the different species and subspecies [2,56-61]; interpretation of in vitro susceptibility testing warrants several caveats that are discussed elsewhere (see 'Caveats to MIC interpretation' below):

M. abscessus:

Usually “susceptible” – amikacin (90 percent), clofazimine (90 percent), cefoxitin (70 percent), tigecycline (most have MIC <1 mcg/mL)

Sometimes “susceptible” – imipenem (50 percent), linezolid (23 percent)

Additional agents that appear to have in vitro activity – omadacycline, eravacycline, tedizolid, bedaquiline

Clarithromycin (macrolide) susceptibility depends on the presence of an active erm gene (see 'Detecting macrolide resistance' below):

-Most M. abscessus subspecies abscessus and M. abscessus subspecies bolletii have an active erm gene and are thus resistant to macrolides. (Approximately 20 percent of M. abscessus subspecies abscessus have an inactive erm gene and are thus susceptible to macrolides [62]).

-Almost all M. abscessus subspecies massiliense lack an active erm gene and are susceptible to macrolides.

M. fortuitum:

Usually susceptible – amikacin (100 percent), imipenem (100 percent) fluoroquinolones (100 percent), sulfonamides (100 percent), linezolid (86 percent), cefoxitin (80 percent)

Sometimes susceptible – doxycycline and minocycline (50 percent)

M. fortuitum isolates have an active erm gene and are thus resistant to macrolides (see 'Detecting macrolide resistance' below)

M. chelonae:

Usually susceptible – clarithromycin (100 percent), tobramycin (100 percent), amikacin (80 percent), moxifloxacin (75 percent), tigecycline (most have MIC <1 mcg/mL)

Sometimes susceptible – imipenem (60 percent), linezolid (54 percent), clofazimine (25 percent), doxycycline (25 percent), ciprofloxacin and levofloxacin (20 percent)

M. abscessus, M. fortuitum, and M. chelonae are resistant to the antituberculosis agents rifampin, ethambutol, and isoniazid, so susceptibility testing of RGM should not be performed with these.

In vitro susceptibility testing is not clinically available at most labs for bedaquiline, omadacycline, or tedizolid. It is assumed that an isolate with a low MIC to tigecycline would also have a low MIC to omadacycline, and MICs for tedizolid are typically lower than those for linezolid. As below, however, there are important caveats to MIC interpretation. (See 'Caveats to MIC interpretation' below.)

The susceptibility testing method for RGM recommended by the CLSI is the broth microdilution method.

Caveats to MIC interpretation — CLSI has set MIC breakpoints for susceptibility and resistance for certain antimicrobial agents for RGM in general. However, there are important caveats in interpreting these MIC values, in particular for M. abscessus:

For M. abscessus, the only two agents for which in vitro susceptibility has been associated with clinical outcomes are clarithromycin (which represents the macrolide class) and amikacin [53,63]. There is no evidence that other antimicrobials with MIC levels in the “susceptible” range are associated with in vivo activity or successful treatment outcomes. Nevertheless, regimens are generally selected according to in vitro susceptibility results, consistent with recommendations from the Infectious Diseases Society of America (IDSA)/American Thoracic Society (ATS) guidelines, despite these limitations and uncertainties. (See 'Treatment' below.)

Most untreated macrolide-susceptible M. abscessus isolates without inducible resistance to macrolides have an MIC ≤2 mcg/mL [63]. However, MICs to clarithromycin (macrolides) that are measured with standard incubation times may be within the susceptible range even if the isolate has inducible resistance to macrolides, which thus needs to be evaluated separately. This is discussed in detail elsewhere. (See 'Detecting macrolide resistance' below.)

The cutoff for intravenous amikacin resistance is an MIC ≥64 mcg/mL; resistance is mediated by a mutation in the 16S rRNA gene [55].

The cutoff for inhaled amikacin resistance is an MIC ≥128 mcg/mL.

Some M. abscessus isolates grow slowly in broth and must be incubated beyond five days for adequate growth and susceptibility testing. However, when this is necessary, only clarithromycin and amikacin MIC results can be reliably reported because other antimicrobials are unstable after this period [55]. As an example, MIC data for imipenem may be unreliable or challenging to reproduce; 85 percent of imipenem in vitro activity is lost after 24 hours incubation [64].

In contrast to M. abscessus, M. fortuitum isolates respond more predictably to antibiotics based on MICs as recommended by CLSI.

Detecting macrolide resistance — RGM can be resistant to macrolides through two mechanisms:

Mutational resistance – A mutation in the 23S ribosomal RNA gene can result in macrolide resistance [55]. This is detected in susceptibility testing assays with standard incubation times (eg, three to five days), which will result in MICs in the resistant range. It can also be detected through a line probe assay.

Inducible resistance – The erm gene is an inducible macrolide resistance gene. Isolates that contain an active erm gene become resistant during exposure to a macrolide (because this induces expression of the erm gene) but may have MICs in the susceptible range when tested with susceptibility assays using standard incubation times. Inducible macrolide resistance through an active erm gene can be identified through susceptibility assays using prolonged (eg, 14 day) incubation times.

Most M. abscessus subspecies abscessus, M. abscessus subspecies bolletii, and M. fortuitum isolates have an active erm gene [65]. Most M. abscessus subspecies massiliense and M. chelonae do not. However, species identification alone cannot predict erm gene activity. Some isolates that would typically have an active erm gene have mutations to the erm gene that render it inactive and restore the in vivo macrolide susceptibility.

Molecular testing can also detect the erm gene as well as an inactivating mutation in the erm gene, thus providing indirect evidence of erm gene activity. However, only phenotypic drug susceptibility testing with macrolide incubation offers a direct indication of erm gene functionality. Thus, we suggest determining erm gene activity through phenotypic drug susceptibility testing with macrolide incubation for these species. (See 'Susceptibility testing' above.)

Determining macrolide susceptibility (which includes ruling out inducible resistance) is critical to management decisions. For M. abscessus, use of macrolide-containing regimens for macrolide-susceptible infection is associated with treatment success, whereas macrolide resistance is associated with poor treatment response [66-68]. The recognition of an inducible macrolide resistance gene offers insight into the historically poor response of M. fortuitum and M. abscessus subspecies abscessus to macrolide-based regimens. (See 'Antimicrobial therapy for M. fortuitum and M. chelonae' below and 'Antimicrobial therapy of M. abscessus species' below.)

TREATMENT — No controlled clinical trials of treatment for disease caused by RGM have been performed. Thus, our treatment suggestions are based on case series, in vitro susceptibility testing, and the clinical experience of experts [53,56-59,63,69,70]. Patients with RGM infections should be treated in consultation with an expert in mycobacterial diseases (eg, infectious disease or pulmonary disease specialists).

Antimicrobial therapy for M. fortuitum and M. chelonae

Limited skin and soft tissue infection — For limited skin and soft tissue infection due to M. fortuitum or M. chelonae, we suggest oral therapy with at least two agents that have in vitro activity against the isolate for a minimum of four months [69,70]. We choose two oral agents based on susceptibility testing from among the following:

Trimethoprim-sulfamethoxazole (1 double-strength tablet twice daily)

Doxycycline (100 to 200 mg daily)

Levofloxacin (500 to 750 mg daily)

Clarithromycin (500 mg twice daily) or azithromycin (250 to 500 mg daily) for isolates that do not have an active inducible erm gene (see 'Detecting macrolide resistance' above)

Monotherapy with these agents should not be used because of concern for acquired resistance and treatment failure.

Other potential agents that may have in vitro activity against M. fortuitum or M. chelonae isolates include moxifloxacin, linezolid, minocycline, and clofazimine [56,57,60,61].

Severe, active, or progressive disease — Severe disease due to M. fortuitum or M. chelonae includes infection that has progressed beyond localized skin involvement, such as extensive skin and soft tissue disease, pulmonary disease, and other forms of disseminated infection.

For severe infection due to M. fortuitum or M. chelonae, we suggest initial therapy with three agents (including one to two intravenous agents) to which the isolate is susceptible [69]. We typically treat with this initial regimen until clinical improvement is evident (two to six weeks), at which point we transition to therapy with two or three oral agents with in vitro activity. For patients with pulmonary disease, treatment is continued for at least 12 months after sputum cultures become negative. For severe skin and soft tissue infection, bone involvement, and disseminated disease, treatment duration is at least 6 to 12 months. The treatment of osteomyelitis due to RGM is discussed in detail elsewhere. (See "Treatment of osteomyelitis due to nontuberculous mycobacteria in adults", section on 'Rapidly growing mycobacteria'.)

A typical initial intravenous regimen includes:

An aminoglycoside:

For M. fortuitum Amikacin is the preferred aminoglycoside. The author typically uses 10 to 15 mg/kg daily as the starting dose for adult patients with normal renal function; some experts use three times weekly dosing. The dose should be adjusted to provide peak serum levels above the amikacin minimum inhibitory concentration (MIC) of the organism (preferably two or three times the MIC if it is ≤16 mcg/mL).

For M. chelonaeTobramycin is the preferred aminoglycoside, although some M. chelonae isolates are amikacin susceptible in vitro as well. The author typically uses 5 mg/kg intravenously daily; some experts use three times weekly dosing.

PLUS one of the following:

Cefoxitin (8 to 12 g intravenously in divided doses; we typically use 3 to 4 g twice daily).

Imipenem (0.5 to 1 g intravenously, given two times per day).

PLUS:

Levofloxacin (500 to 750 mg intravenously or orally once daily).

Following the initial intravenous course, we switch to a combination of at least two oral agents selected among the following based on in vitro susceptibility testing:

Trimethoprim-sulfamethoxazole (1 double-strength tablet twice daily)

Doxycycline (100 to 200 mg daily)

Levofloxacin (500 to 750 mg daily)

Clarithromycin (500 mg twice daily) or azithromycin (250 to 500 mg daily) for isolates that do not have an active inducible erm gene (see 'Detecting macrolide resistance' above)

Other potential agents that may have in vitro activity against M. fortuitum or M. chelonae isolates include tigecycline, moxifloxacin, linezolid, minocycline, and clofazimine. (See 'Agents to test' above.)

Antimicrobial therapy of M. abscessus species — The approach to treatment of M. abscessus species depends on whether the isolate is macrolide susceptible or has inducible or mutational resistance to macrolides (clarithromycin and azithromycin). Assessment for macrolide resistance is discussed elsewhere. (See 'Detecting macrolide resistance' above.)

Patients should be managed in consultation with a mycobacterial diseases specialist, or infectious disease and pulmonary specialists with experience treating M. abscessus disease. Data to guide antimicrobial therapy of M. abscessus are limited, and the optimal approach is uncertain, particularly for isolates with macrolide resistance. The antibiotic selection and dosing recommendations presented reflect the author’s approach and are consistent with expert guidelines on the treatment of nontuberculous mycobacteria (NTM) [53]. These guidelines use in vitro susceptibility results to guide the selection of agents, although, as noted elsewhere, it is not clear if in vitro susceptibility reliably predicts treatment response for antibiotics other than clarithromycin (macrolides) or amikacin. (See 'Caveats to MIC interpretation' above.)

The treatment of osteomyelitis due to M. abscessus is discussed in detail elsewhere. (See "Treatment of osteomyelitis due to nontuberculous mycobacteria in adults", section on 'Rapidly growing mycobacteria'.)

Deciding to treat

Pulmonary disease – For patients who meet diagnostic criteria for M. abscessus pulmonary disease (table 2), the decision to treat is similar to that for M. avium complex lung disease and depends on clinical symptoms and comorbidities as well as radiographic findings (eg, presence of cavitary disease) and microbiologic findings (eg, sputum smear positivity), which are associated with disease progression. This is discussed in detail elsewhere. (See "Treatment of Mycobacterium avium complex pulmonary infection in adults", section on 'Deciding to treat'.)

Nonpulmonary disease – Treatment is warranted for all patients with a clear diagnosis of infection at nonpulmonary sites. (See 'Nonpulmonary disease' above.)

Macrolide-susceptible isolates (without inducible resistance) — For patients with disease due to M. abscessus subspecies massiliense or other M. abscessus subspecies that do not have an active erm gene (ie, they are macrolide susceptible), macrolides are the cornerstone of therapy [53,63].

Initial therapy – For initial therapy, we suggest a macrolide (eg, azithromycin) in combination with at least two other agents (including at least one intravenous agent) that have in vitro activity. Doses are listed in the table (table 3):

Azithromycin (or clarithromycin)

PLUS one of the following intravenous agents

-Amikacin (preferred)

-Imipenem

-Cefoxitin

PLUS at least one of the following:

-Omadacycline (preferred, can be given orally or intravenously) or tigecycline

-An oxazolidinone, either tedizolid (preferred oxazolidinone) or linezolid

-Clofazimine

For macrolide-susceptible M. abscessus lung infection, an example of an initial regimen is azithromycin, intravenous amikacin, and oral omadacycline. Based on clinical experience, the author favors other drugs over clofazimine.

Maintenance therapy – Following the initial intravenous regimen of macrolide-susceptible M. abscessus infection, we generally transition to oral/inhaled maintenance therapy. In such cases, we continue the oral macrolide and combine it with at least two other agents, typically inhaled amikacin (if pulmonary disease) and omadacycline. Tedizolid, linezolid, and clofazamine are additional options. Doses are listed in the table (table 3).

The optimal time to switch to maintenance therapy is not well defined. The practice of switching is based not on prospective data but on concerns about toxicity and inconvenience of prolonged intravenous therapy. Thus, common practice is to switch after about three months of initial intravenous antibiotics. However, the author of this topic believes that there are not sufficient data to be dogmatic about this approach, and the time to switch should depend on the maintenance regimen that can be used. As an example, for a patient with macrolide-susceptible M. abscessus lung infection, a maintenance regimen of azithromycin, inhaled amikacin, and omadacycline, with or without clofazimine may be reasonably expected to be effective and minimize the risk of acquired mutational resistance to the azithromycin and/or amikacin; thus, switching from an intravenous agent to this regimen after three months would be appropriate. However, if omadacycline is not available (because of cost or other reasons), the author is concerned, based on clinical experience, that transition to a different oral/inhaled maintenance regimen might not sufficiently prevent acquired mutational resistance to azithromycin and/or amikacin, and so would favor continuing the intravenous regimen for a longer duration (ie, as long as tolerated up to the total duration of therapy). In particular, the author discourages use of azithromycin plus clofazimine alone, or amikacin plus clofazimine alone, because of the concern about acquired mutational resistance to azithromycin or amikacin and the resulting poor prognosis.

Total duration of therapy – For patients with pulmonary disease, treatment is continued for at least 12 months after sputum cultures become negative [71]. For severe skin and soft tissue infection, bone involvement, and disseminated disease, treatment duration is at least 6 to 12 months.

The treatment response and prognosis for patients with macrolide-susceptible isolates are good and better than for patients with macrolide-resistant isolates [16,72]. In a systematic review of 13 observational studies evaluating macrolide-containing regimens, rates of sustained sputum culture clearance without relapse were substantially higher for M. abscessus subspecies massiliense, which does not typically have an active erm gene, compared with M. abscessus subspecies abscessus, which typically does have an active erm gene and thus inducible resistance (84 versus 23 percent) [73]. One of those studies included 43 patients with M. abscessus subspecies massiliense pulmonary disease who received a macrolide plus amikacin plus cefoxitin or imipenem for two weeks followed by a macrolide until sputum cultures were negative for 12 months [72]. All had symptomatic improvement, 91 percent had radiographic improvement, and 91 percent converted to negative sputum cultures after 12 months of therapy. Seven percent had microbiologic relapse. Isolates in two patients acquired macrolide resistance.

The rationale to use multiple drugs in combination is to prevent acquired resistance to any one agent (in particular, the macrolide) during treatment. Given that macrolide-resistant infection is associated with substantially worse treatment outcomes than macrolide-susceptible infection, we prioritize preservation of macrolide susceptibility during treatment. Thus, despite its successful use in some observational studies (including the study described above), we strongly recommend against macrolide monotherapy for M. abscessus infection of any kind, as this can lead to acquired macrolide resistance and significantly lower the chance for treatment success [74]. Similarly, we ensure that a sufficient number of companion drugs that have in vitro activity are used in combination with the macrolide, as inadequate companion therapy can also increase the risk of acquired macrolide resistance. Clinicians should be cognizant of this risk when selecting maintenance regimen antibiotics.

Macrolide-resistant isolates (with inducible or mutational resistance)

Initial intravenous therapy — For initial treatment of infections due to M. abscessus subspecies abscessus and M. abscessus subspecies bolletii isolates that have an active erm gene (or for other macrolide-resistant isolates), we suggest a multidrug regimen that includes intravenous amikacin plus at least three additional agents, one of which is intravenous, chosen based on results of in vitro susceptibility testing and expected tolerance (doses are listed in the table (table 3)):

Amikacin

PLUS one or two of the following intravenous agents:

Imipenem (preferred)

Cefoxitin

PLUS at least one or two of the following agents:

Omadacycline (preferred, can be given orally or intravenously) or tigecycline

An oxazolidinone, either tedizolid (preferred oxazolidinone) or linezolid

Clofazimine

For macrolide-resistant M. abscessus lung infection, an example of an initial regimen is intravenous amikacin, intravenous imipenem, oral omadacycline, and clofazimine.

Although we do not suggest a macrolide for treatment of erm gene-active M. abscessus isolates, a macrolide may still be a useful component of the treatment regimen for patients with bronchiectasis as an immune-modulating agent [53,75,76]. (See "Bronchiectasis in adults: Maintaining lung health", section on 'Macrolides'.)

Because of the difficulty treating (and the low likelihood of cure with medical therapy alone), the possibility of surgical resection should also be evaluated. (See 'Adjunctive surgical therapy' below.)

Evidence informing treatment regimens for macrolide-resistant M. abscessus infections are limited mainly to observational studies of patients with pulmonary infection, and those suggest overall poor microbiologic outcomes [66-68]. In a systematic review of 13 observational studies, rates of sustained sputum culture clearance without relapse for M. abscessus subspecies abscessus, which usually has an active erm gene resulting in inducible macrolide resistance, was 23 percent [73]. The antimicrobial regimens in these studies included at least three agents, but the specific agents varied; all included a macrolide, most included amikacin, although at different doses and durations, and other companion agents included imipenem, cefoxitin, tetracyclines, and fluoroquinolones. In another analysis evaluating the impact of individual agents for M. abscessus subspecies abscessus pulmonary infection based on data from 303 patients in eight studies, intravenous amikacin was associated with treatment success (sustained sputum clearance), although this occurred only 39 percent of those who received amikacin [77].

The rationale to use multiple drugs in combination is to prevent acquired resistance to any one agent (in particular, amikacin) during treatment.

Whether to include a macrolide as part of the treatment regimen of an isolate with an active erm gene is controversial. In one study that included 120 patients with M. abscessus pulmonary infection treated with regimens that did not include a macrolide, the sputum conversion rate was approximately 8 percent [9], which is lower than that reported with macrolide-containing regimens. Additionally, in the analysis of the impact of individual agents described above, a macrolide was also associated with sustained sputum clearance [77]. However, erm gene activity was not documented in most studies, so it could be that some of the possible benefit from a macrolide in these analyses was related to inclusion of isolates that were macrolide susceptible. Because of this, we do not count a macrolide as an active agent for macrolide-resistant infections; nevertheless, as above, macrolides may be useful for lung disease due to macrolide-resistant M. abscessus because of the immune-modulating effects. (See "Bronchiectasis in adults: Maintaining lung health", section on 'Macrolides'.)

Antimicrobial treatment can be associated with clinical improvement, even in the absence of microbiologic cure. In one retrospective study that followed 69 patients treated for M. abscessus pulmonary disease for a mean of 34 months, cough, sputum production, and fatigue at least remained stable in 80 percent and improved or resolved in 69 and 59 percent, respectively, even though the sustained sputum clearance rate was only 48 percent [78]. In another retrospective study, the symptom response rate was 83 percent in the setting of a sustained sputum clearance rate of 58 percent [79].

Deciding whether and when to transition to oral/inhaled maintenance therapy — The approach to transitioning to a nonintravenous regimen for macrolide-resistant M. abscessus isolates is uncertain given the lack of oral agents with demonstrated efficacy (see 'Oral/inhaled maintenance therapy' below). Options include:

Continuing intravenous therapy for a complete treatment course. (See 'Total duration of therapy' below.)

Continuing intravenous therapy until symptoms have improved or resolved. At that point, parenteral therapy can be stopped without transitioning to an oral regimen with plans to reinstitute the same parenteral regimen if symptoms recur or worsen.

Continuing initial intravenous therapy for a predefined duration and then transitioning to an oral/inhaled maintenance regimen for the remainder of the treatment course. The optimal duration of initial intravenous therapy in this context is uncertain. An arbitrary minimal target duration of 8 to 12 weeks is often used. However, because of limitations with the oral regimens used for maintenance therapy, we try to maximize the duration of initial parenteral therapy in patients who are improving on it for as long as it is tolerated before transitioning the regimen. Potential oral/inhaled maintenance regimens and their limitations are discussed elsewhere. (See 'Oral/inhaled maintenance therapy' below.)

The choice among the options depends on the extent of disease, the severity of symptoms, and how well the patient tolerates the intravenous regimen. These same considerations apply to pulmonary and extrapulmonary infections. However, since total treatment durations are generally shorter for extrapulmonary infections, it may be easier to continue intravenous medications for a longer proportion of the total treatment duration (rather than switching to an oral maintenance regimen at an arbitrary time). (See 'Total duration of therapy' below.)

Available data do not inform whether one approach is superior to the others. Most studies describing treatment of M. abscessus pulmonary infection have used two to four months of initial intravenous therapy followed by an oral maintenance regimen for a total treatment duration greater than one year; however, in some cases, intravenous therapy was continued for up to 20 months [71]. Even with prolonged treatment, microbiologic response rates are low, as discussed elsewhere (see 'Initial intravenous therapy' above). There are also no data informing whether continuation with oral maintenance therapy improves response rates compared with initial intravenous therapy alone.

Oral/inhaled maintenance therapy

Potential agents to use in combination For patients with M. abscessus isolates with mutational or inducible resistance to macrolides who transition to an oral/inhaled maintenance phase following the initial parenteral treatment phase, we suggest a regimen with at least three oral or inhaled agents with in vitro activity against the M. abscessus isolate [53].

Oral agents that may have in vitro activity against M. abscessus species include clofazimine, omadacycline, either tedizolid or linezolid, and possibly bedaquiline [60,80-82]. We also use inhaled amikacin for susceptible isolates in patients with pulmonary disease. Doses are listed in the table (table 3).

Although we do not suggest a macrolide for treatment of erm gene-active M. abscessus isolates, a macrolide may still be a useful component of the treatment regimen for patients with bronchiectasis as an immune-modulating agent [53]. (See "Bronchiectasis in adults: Maintaining lung health", section on 'Macrolides'.)

Clofazimine is frequently used as part of treatment regimens for M. abscessus because the MIC for this agent is typically low, it is administered orally, and it appears to be safe at a 100 mg daily dose. However, evidence for its efficacy for treating M. abscessus is limited; although it has been studied in multidrug regimens, its contribution to the efficacy of those regimens is unknown [81-83]. As below, the author discourages use of clofazimine as the sole companion drug to amikacin because of concerns about acquired mutational resistance to amikacin.

Data on other non-macrolide oral drugs are similarly limited, and it is not yet clear whether any of these have significant clinical impact. Omadacycline has garnered particular interest because it is an oral agent with in vitro activity similar to tigecycline, and limited early clinical experience is promising [84-86]. The M. abscessus MICs to tedizolid are generally several-fold lower than to linezolid, but use of either may be limited by hematologic adverse effects or peripheral neuropathy [60,80,87,88]. Clinical data on bedaquiline are extremely limited [89,90].

Data on inhaled amikacin for pulmonary disease are emerging but remain limited [91-93]. In a small observational study that included 36 patients with M. abscessus subspecies abscessus infection, a maintenance regimen including inhaled amikacin was associated with microbiologic cure in 38 percent and clinical improvement in 78 percent [91].

There is no evidence that fluoroquinolones (ie, moxifloxacin) add to the efficacy of any regimen for any M. abscessus subspecies.

Limitations to oral/inhaled maintenance therapy Constructing an effective maintenance regimen for M. abscessus isolates with mutational or inducible resistance to macrolides is frequently challenging because of the limited number of oral agents that have in vitro activity against such isolates. Even for those agents that have low MICs for M. abscessus in vitro, data demonstrating clinical efficacy are lacking. Additionally, some promising oral agents (omadacycline, tedizolid, bedaquiline) have prohibitively high cost for patients. Finally, long-term exposure to these drugs increases the risk of toxicities, and there is no clear evidence indicating that prolonged therapy can reliably achieve a clinical response.

Another important potential limitation of oral/inhaled maintenance therapy is the possibility of acquired mutational resistance to amikacin if companion drugs are insufficient to protect against such resistance. For instance, a regimen of inhaled amikacin, omadacycline, clofazimine, and tedizolid (or linezolid) would likely be sufficient to minimize the risk of acquired mutational resistance to amikacin. However, if omadacycline and the oxazolidinones are not available due to cost, it is unclear based on the author’s clinical experience whether clofazimine alone is adequate as a single companion drug to protect against amikacin resistance. Given the potential negative impact of emergent amikacin resistance on treatment response, the author discourages the use of inhaled amikacin plus clofazimine alone.

Total duration of therapy — For patients with pulmonary disease, the goal is to treat until the patient has negative sputum cultures for 12 months, although this may not be an attainable outcome for macrolide-resistant infection [71]. For extrapulmonary disease, treatment duration is generally 6 to 12 months.

Investigational approaches — Novel approaches for M. abscessus therapies include dual beta-lactam therapy and bacteriophages. Experience with these is highly limited, and clinicians should consult with an expert in the management of NTM if considering these treatment modalities.

Combinations of two beta-lactam agents (eg, imipenem with ceftaroline or ceftazidime) with or without a beta-lactamase inhibitor (avibactam, relebactam) have been proposed as therapy for M. abscessus infection [94,95]. Thus far, there have only been anecdotal reports of effective use of dual beta-lactam therapy. The author of this topic has limited experience with this approach, and results have been inconsistent. The several-times daily dosing required with these regimens also limit their utility.

Engineered bacteriophages are another investigational approach [96-98]. Thus far, there have only been a few anecdotal reports, also with inconsistent outcomes. Additionally, the bacteriophage must be tailored to the individual M. abscessus isolate.

Adjunctive surgical therapy — Surgery can be an important adjunct to medical therapy and should be considered whenever possible for these difficult-to-treat patients. Surgery is generally indicated for extensive cutaneous disease, abscess formation, or insufficient response to antimicrobial therapy. Removal of foreign bodies (eg, breast implants, percutaneous catheters) is essential for cure [52].

Surgical resection for limited pulmonary disease may also be curative; some observational data suggest a higher rate of microbiologic cure of lung disease with surgical intervention, which has been performed for localized bronchiectasis, cavitary disease, hemoptysis, and poor response to antimicrobial therapy [9,78]. However, the decision to perform surgery should be made by clinicians with experience in the treatment of NTM disease. Thoracic surgery for M. abscessus infection may be associated with significant complications (eg, postoperative hemorrhage, bronchopleural fistula, frozen shoulder, wound infection, brachial plexus injury, and respiratory failure and/or death), even in expert hands, and should only be performed in centers with extensive experience [9,53].

Monitoring for drug toxicity — Monitoring for drug toxicity of patients during treatment of NTM disease is important, given the number and type of drugs used and the older age of these patients. Monitoring includes routine interval testing of the complete blood count, blood urea nitrogen and creatinine, and liver enzymes.

For patients taking aminoglycosides, monitoring should also include routine questioning about balance, ability to walk (especially in the dark), tinnitus, dizziness, and difficulty hearing. A baseline hearing test should be done and then repeated periodically for the duration of aminoglycoside use.

Monitoring is similar to that during treatment for M. avium complex infection, which is discussed elsewhere. (See "Treatment of Mycobacterium avium complex pulmonary infection in adults", section on 'Monitoring for side effects'.)

Prognosis — In retrospective studies, the mortality directly attributable to M. abscessus infections treated with combination antibiotic regimens with or without surgery has been approximately 15 percent, although that figure likely overestimates M. abscessus-related mortality for patients with macrolide-susceptible disease [9,53,63,78]. Time from diagnosis of M. abscessus disease to infection-related death is variable and depends on the severity of underlying lung disease [9]. (See 'Pulmonary infection' above.)

SOCIETY GUIDELINE LINKS — Links to society and government-sponsored guidelines from selected countries and regions around the world are provided separately. (See "Society guideline links: Nontuberculous mycobacteria".)

SUMMARY AND RECOMMENDATIONS

Disease associations – The diseases caused by the three most clinically relevant rapidly growing mycobacteria (RGM) species vary:

Mycobacterium abscessus is the most pathogenic RGM and usually causes pulmonary infection in patients with or without underlying lung disease. It is also associated with disseminated disease in immunocompromised patients, skin and soft tissue infections, and musculoskeletal infections.

Mycobacterium fortuitum primarily causes infection by direct inoculation, resulting in skin and soft tissue, surgical site, and catheter-related infections. Most respiratory isolates are from individuals with underlying pulmonary diseases and represent colonization or transient infection.

Mycobacterium chelonae most commonly causes infection in immunocompromised patients. (See 'Clinical manifestations' above.)

Diagnosis – The diagnosis of RGM infection is made by culture of the specific pathogen from a relevant clinical specimen (eg, blood, drainage material, tissue biopsy, or specimen from typically sterile sites). For pulmonary disease, the diagnosis requires meeting clinical, radiographic, and microbiologic criteria for nontuberculous mycobacterial (NTM) lung disease (table 2) because of the possibility that respiratory specimens could reflect colonization rather than clinically significant infection.

Importance of species identification – Distinction of RGM from Mycobacterium tuberculosis is essential because of the public health and treatment implications. Treatment recommendations also rely on accurate identification of the RGM species. In particular, M. abscessus should be speciated and reported to the subspecies level, if possible. (See 'Mycobacterial species identification' above and 'Nomenclature for M. abscessus' above.)

Susceptibility testing – This should be performed on all clinically significant isolates, including isolates that have been recovered after treatment failure or relapse. Drugs to test include clarithromycin, amikacin, tigecycline, imipenem, cefoxitin, clofazimine, doxycycline, linezolid, fluoroquinolones, and a sulfonamide. Additionally, all M. abscessus and M. fortuitum isolates should be tested for inducible macrolide resistance. (See 'Susceptibility testing' above.)

Expert involvement in management – Data informing optimal management of RGM are limited; management is complicated and involves prolonged therapy with antimicrobials associated with substantial toxicity. Antimicrobial regimens are selected based on susceptibility testing results, which are challenging to interpret. Patients should be managed in consultation with a specialist with expertise in treating NTM diseases. (See 'Treatment' above.)

The treatment of osteomyelitis due to RGM is discussed elsewhere. (See "Treatment of osteomyelitis due to nontuberculous mycobacteria in adults", section on 'Rapidly growing mycobacteria'.)

Antimicrobial regimens for M. fortuitum or M. chelonae

For patients with limited skin and soft tissue infection due to M. fortuitum or M. chelonae, we suggest therapy with at least two agents to which the isolate is susceptible (Grade 2C). Options generally include trimethoprim-sulfamethoxazole, doxycycline, levofloxacin, and, for isolates without inducible macrolide resistance, azithromycin or clarithromycin. Duration is generally a minimum of four months. (See 'Limited skin and soft tissue infection' above.)

For more severe infections due to M. fortuitum or M. chelonae, we suggest initial therapy with an intravenous aminoglycoside plus at least two other active agents (Grade 2C). Options include imipenem, cefoxitin, and levofloxacin. Once clinical improvement is evident (generally after two to six weeks), the regimen can be transitioned to an oral regimen with two or three active agents. For severe skin and soft tissue infection, bone involvement, and disseminated disease, treatment duration is at least 6 to 12 months. (See 'Severe, active, or progressive disease' above.)

Antimicrobial regimens for M. abscessus infection

For M. abscessus isolates that have no inducible macrolide resistance (eg, most M. abscessus subspecies massiliense or M. abscessus isolates with an inactive erm gene), we suggest initial combination therapy with a macrolide plus amikacin plus at least one other active agent (table 3) (Grade 2C). After 8 to 12 weeks of initial therapy, the regimen can generally be transitioned to maintenance therapy with oral (and inhaled) agents. For maintenance therapy, we suggest a macrolide plus at least two other active agents (Grade 2C). For pulmonary disease, treatment continues until sputum cultures are negative for 12 months; for nonpulmonary infection, duration is generally at least 6 to 12 months. (See 'Macrolide-susceptible isolates (without inducible resistance)' above.)

For M. abscessus isolates with inducible or mutational resistance to macrolides (eg, M. abscessus subspecies abscessus and M. abscessus subspecies bolletii that have an active erm gene), we suggest initial combination therapy with amikacin plus at least three other active agents (Grade 2C). Additional options include imipenem, omadacycline, tedizolid/linezolid, and clofazimine (table 3). Options for ongoing therapy include continuing the parenteral regimen for the duration of treatment (eg, until sputum cultures have cleared for 12 months for pulmonary disease), continuing the parenteral regimen until symptoms resolve then discontinuing it with plans to restart if disease worsens, and continuing the parenteral regimen for a discrete period (eg, 8 to 12 weeks) or for as long as the patient can tolerate it before transitioning to a regimen of at least three oral and/or inhaled agents to which the isolate is susceptible in vitro. (See 'Macrolide-resistant isolates (with inducible or mutational resistance)' above.)

Adjunctive surgery – Surgery is generally indicated for extensive cutaneous disease, abscess formation, foreign body removal, or cases in which antimicrobial therapy is difficult or unsuccessful. Surgical resection of limited pulmonary disease has been associated with improved outcomes but should be performed by clinicians with extensive experience in NTM disease. (See 'Adjunctive surgical therapy' above.)

  1. Tortoli E, Kohl TA, Brown-Elliott BA, et al. Emended description of Mycobacterium abscessus, Mycobacterium abscessus subsp. abscessus and Mycobacteriumabscessus subsp. bolletii and designation of Mycobacteriumabscessus subsp. massiliense comb. nov. Int J Syst Evol Microbiol 2016; 66:4471.
  2. Wallace RJ Jr, Brown BA, Onyi GO. Skin, soft tissue, and bone infections due to Mycobacterium chelonae chelonae: importance of prior corticosteroid therapy, frequency of disseminated infections, and resistance to oral antimicrobials other than clarithromycin. J Infect Dis 1992; 166:405.
  3. Hoffman PC, Fraser DW, Robicsek F, et al. Two outbreaks of sternal wound infection due to organisms of the Mycobacterium fortuitum complex. J Infect Dis 1981; 143:533.
  4. Wallace RJ Jr, Swenson JM, Silcox VA, et al. Spectrum of disease due to rapidly growing mycobacteria. Rev Infect Dis 1983; 5:657.
  5. Chang MA, Jain S, Azar DT. Infections following laser in situ keratomileusis: an integration of the published literature. Surv Ophthalmol 2004; 49:269.
  6. Kuritsky JN, Bullen MG, Broome CV, et al. Sternal wound infections and endocarditis due to organisms of the Mycobacterium fortuitum complex. Ann Intern Med 1983; 98:938.
  7. Park S, Suh GY, Chung MP, et al. Clinical significance of Mycobacterium fortuitum isolated from respiratory specimens. Respir Med 2008; 102:437.
  8. Lange C, Böttger EC, Cambau E, et al. Consensus management recommendations for less common non-tuberculous mycobacterial pulmonary diseases. Lancet Infect Dis 2022; 22:e178.
  9. Griffith DE, Girard WM, Wallace RJ Jr. Clinical features of pulmonary disease caused by rapidly growing mycobacteria. An analysis of 154 patients. Am Rev Respir Dis 1993; 147:1271.
  10. Dreisin RB, Scoggin C, Davidson PT. The pathogenicity of Mycobacterium fortuitum and Mycobacterium chelonei in man: a report of seven cases. Tubercle 1976; 57:49.
  11. Boxerbaum B. Isolation of rapidly growing mycobacteria in patients with cystic fibrosis. J Pediatr 1980; 96:689.
  12. Awe RJ, Gangadharam PR, Jenkins DE. Clinical significance of Mycobacterium fortuitum infections in pulmonary disease. Am Rev Respir Dis 1973; 108:1230.
  13. Varghese G, Shepherd R, Watt P, Bruce JH. Fatal infection with Mycobacterium fortuitum associated with oesophageal achalasia. Thorax 1988; 43:151.
  14. Rolston KV, Jones PG, Fainstein V, Bodey GP. Pulmonary disease caused by rapidly growing mycobacteria in patients with cancer. Chest 1985; 87:503.
  15. Redelman-Sidi G, Sepkowitz KA. Rapidly growing mycobacteria infection in patients with cancer. Clin Infect Dis 2010; 51:422.
  16. Park J, Cho J, Lee CH, et al. Progression and Treatment Outcomes of Lung Disease Caused by Mycobacterium abscessus and Mycobacterium massiliense. Clin Infect Dis 2017; 64:301.
  17. Shin SJ, Choi GE, Cho SN, et al. Mycobacterial genotypes are associated with clinical manifestation and progression of lung disease caused by Mycobacterium abscessus and Mycobacterium massiliense. Clin Infect Dis 2013; 57:32.
  18. Wentworth AB, Drage LA, Wengenack NL, et al. Increased incidence of cutaneous nontuberculous mycobacterial infection, 1980 to 2009: a population-based study. Mayo Clin Proc 2013; 88:38.
  19. Uslan DZ, Kowalski TJ, Wengenack NL, et al. Skin and soft tissue infections due to rapidly growing mycobacteria: comparison of clinical features, treatment, and susceptibility. Arch Dermatol 2006; 142:1287.
  20. Winthrop KL, Albridge K, South D, et al. The clinical management and outcome of nail salon-acquired Mycobacterium fortuitum skin infection. Clin Infect Dis 2004; 38:38.
  21. Winthrop KL, Abrams M, Yakrus M, et al. An outbreak of mycobacterial furunculosis associated with footbaths at a nail salon. N Engl J Med 2002; 346:1366.
  22. Vugia DJ, Jang Y, Zizek C, et al. Mycobacteria in nail salon whirlpool footbaths, California. Emerg Infect Dis 2005; 11:616.
  23. Stout JE, Gadkowski LB, Rath S, et al. Pedicure-associated rapidly growing mycobacterial infection: an endemic disease. Clin Infect Dis 2011; 53:787.
  24. Sergeant A, Conaglen P, Laurenson IF, et al. Mycobacterium chelonae infection: a complication of tattooing. Clin Exp Dermatol 2013; 38:140.
  25. Bechara C, Macheras E, Heym B, et al. Mycobacterium abscessus skin infection after tattooing: first case report and review of the literature. Dermatology 2010; 221:1.
  26. Drage LA, Ecker PM, Orenstein R, et al. An outbreak of Mycobacterium chelonae infections in tattoos. J Am Acad Dermatol 2010; 62:501.
  27. Kennedy BS, Bedard B, Younge M, et al. Outbreak of Mycobacterium chelonae infection associated with tattoo ink. N Engl J Med 2012; 367:1020.
  28. Centers for Disease Control and Prevention (CDC). Tattoo-associated nontuberculous mycobacterial skin infections--multiple states, 2011-2012. MMWR Morb Mortal Wkly Rep 2012; 61:653.
  29. Falsey RR, Kinzer MH, Hurst S, et al. Cutaneous inoculation of nontuberculous mycobacteria during professional tattooing: a case series and epidemiologic study. Clin Infect Dis 2013; 57:e143.
  30. Griffin I, Schmitz A, Oliver C, et al. Outbreak of Tattoo-associated Nontuberculous Mycobacterial Skin Infections. Clin Infect Dis 2019; 69:949.
  31. LeBlanc PM, Hollinger KA, Klontz KC. Tattoo ink-related infections--awareness, diagnosis, reporting, and prevention. N Engl J Med 2012; 367:985.
  32. Mateo L, Rufí G, Nolla JM, Alcaide F. Mycobacterium chelonae tenosynovitis of the hand. Semin Arthritis Rheum 2004; 34:617.
  33. Centers for Disease Control and Prevention (CDC). Nontuberculous mycobacterial infections after cosmetic surgery--Santo Domingo, Dominican Republic, 2003-2004. MMWR Morb Mortal Wkly Rep 2004; 53:509.
  34. Regnier S, Cambau E, Meningaud JP, et al. Clinical management of rapidly growing mycobacterial cutaneous infections in patients after mesotherapy. Clin Infect Dis 2009; 49:1358.
  35. Rimmer J, Hamilton S, Gault D. Recurrent mycobacterial breast abscesses complicating reconstruction. Br J Plast Surg 2004; 57:676.
  36. John T, Velotta E. Nontuberculous (atypical) mycobacterial keratitis after LASIK: current status and clinical implications. Cornea 2005; 24:245.
  37. Freitas D, Alvarenga L, Sampaio J, et al. An outbreak of Mycobacterium chelonae infection after LASIK. Ophthalmology 2003; 110:276.
  38. Sampaio JL, Junior DN, de Freitas D, et al. An outbreak of keratitis caused by Mycobacterium immunogenum. J Clin Microbiol 2006; 44:3201.
  39. Edens C, Liebich L, Halpin AL, et al. Mycobacterium chelonae Eye Infections Associated with Humidifier Use in an Outpatient LASIK Clinic--Ohio, 2015. MMWR Morb Mortal Wkly Rep 2015; 64:1177.
  40. Toy BR, Frank PJ. Outbreak of Mycobacterium abscessus infection after soft tissue augmentation. Dermatol Surg 2003; 29:971.
  41. Centers for Disease Control and Prevention (CDC). Mycobacterium chelonae infections associated with face lifts--New Jersey, 2002-2003. MMWR Morb Mortal Wkly Rep 2004; 53:192.
  42. Furuya EY, Paez A, Srinivasan A, et al. Outbreak of Mycobacterium abscessus wound infections among "lipotourists" from the United States who underwent abdominoplasty in the Dominican Republic. Clin Infect Dis 2008; 46:1181.
  43. Duarte RS, Lourenço MC, Fonseca Lde S, et al. Epidemic of postsurgical infections caused by Mycobacterium massiliense. J Clin Microbiol 2009; 47:2149.
  44. Leao SC, Tortoli E, Euzéby JP, Garcia MJ. Proposal that Mycobacterium massiliense and Mycobacterium bolletii be united and reclassified as Mycobacterium abscessus subsp. bolletii comb. nov., designation of Mycobacterium abscessus subsp. abscessus subsp. nov. and emended description of Mycobacterium abscessus. Int J Syst Evol Microbiol 2011; 61:2311.
  45. van Dissel JT, Kuijper EJ. Rapidly growing mycobacteria: emerging pathogens in cosmetic procedures of the skin. Clin Infect Dis 2009; 49:1365.
  46. Raad II, Vartivarian S, Khan A, Bodey GP. Catheter-related infections caused by the Mycobacterium fortuitum complex: 15 cases and review. Rev Infect Dis 1991; 13:1120.
  47. Ellis EN, Schutze GE, Wheeler JG. Nontuberculous mycobacterial exit-site infection and abscess in a peritoneal dialysis patient. A case report and review of the literature. Pediatr Nephrol 2005; 20:1016.
  48. Vera G, Lew SQ. Mycobacterium fortuitum peritonitis in two patients receiving continuous ambulatory peritoneal dialysis. Am J Nephrol 1999; 19:586.
  49. Franklin DJ, Starke JR, Brady MT, et al. Chronic otitis media after tympanostomy tube placement caused by Mycobacterium abscessus: a new clinical entity? Am J Otol 1994; 15:313.
  50. Sharma S, Tleyjeh IM, Espinosa RE, et al. Pacemaker infection due to Mycobacterium fortuitum. Scand J Infect Dis 2005; 37:66.
  51. Eid AJ, Berbari EF, Sia IG, et al. Prosthetic joint infection due to rapidly growing mycobacteria: report of 8 cases and review of the literature. Clin Infect Dis 2007; 45:687.
  52. Griffith DE, Aksamit T, Brown-Elliott BA, et al. An official ATS/IDSA statement: diagnosis, treatment, and prevention of nontuberculous mycobacterial diseases. Am J Respir Crit Care Med 2007; 175:367.
  53. Daley CL, Iaccarino JM, Lange C, et al. Treatment of Nontuberculous Mycobacterial Pulmonary Disease: An Official ATS/ERS/ESCMID/IDSA Clinical Practice Guideline. Clin Infect Dis 2020; 71:e1.
  54. Griffith DE, Philley JV, Brown-Elliott BA, et al. The significance of Mycobacterium abscessus subspecies abscessus isolation during Mycobacterium avium complex lung disease therapy. Chest 2015; 147:1369.
  55. Brown-Elliott BA, Woods GL. Antimycobacterial Susceptibility Testing of Nontuberculous Mycobacteria. J Clin Microbiol 2019; 57.
  56. Stone MS, Wallace RJ Jr, Swenson JM, et al. Agar disk elution method for susceptibility testing of Mycobacterium marinum and Mycobacterium fortuitum complex to sulfonamides and antibiotics. Antimicrob Agents Chemother 1983; 24:486.
  57. Swenson JM, Wallace RJ Jr, Silcox VA, Thornsberry C. Antimicrobial susceptibility of five subgroups of Mycobacterium fortuitum and Mycobacterium chelonae. Antimicrob Agents Chemother 1985; 28:807.
  58. Brown BA, Wallace RJ Jr, Onyi GO, et al. Activities of four macrolides, including clarithromycin, against Mycobacterium fortuitum, Mycobacterium chelonae, and M. chelonae-like organisms. Antimicrob Agents Chemother 1992; 36:180.
  59. Wallace RJ Jr, Brown BA, Onyi GO. Susceptibilities of Mycobacterium fortuitum biovar. fortuitum and the two subgroups of Mycobacterium chelonae to imipenem, cefmetazole, cefoxitin, and amoxicillin-clavulanic acid. Antimicrob Agents Chemother 1991; 35:773.
  60. Wallace RJ Jr, Brown-Elliott BA, Ward SC, et al. Activities of linezolid against rapidly growing mycobacteria. Antimicrob Agents Chemother 2001; 45:764.
  61. Wallace RJ Jr, Bedsole G, Sumter G, et al. Activities of ciprofloxacin and ofloxacin against rapidly growing mycobacteria with demonstration of acquired resistance following single-drug therapy. Antimicrob Agents Chemother 1990; 34:65.
  62. Brown-Elliott BA, Vasireddy S, Vasireddy R, et al. Utility of sequencing the erm(41) gene in isolates of Mycobacterium abscessus subsp. abscessus with low and intermediate clarithromycin MICs. J Clin Microbiol 2015; 53:1211.
  63. Griffith DE, Daley CL. Treatment of Mycobacterium abscessus Pulmonary Disease. Chest 2022; 161:64.
  64. Brown-Elliott BA, Killingley J, Vasireddy S, et al. In Vitro Comparison of Ertapenem, Meropenem, and Imipenem against Isolates of Rapidly Growing Mycobacteria and Nocardia by Use of Broth Microdilution and Etest. J Clin Microbiol 2016; 54:1586.
  65. Nash KA, Brown-Elliott BA, Wallace RJ Jr. A novel gene, erm(41), confers inducible macrolide resistance to clinical isolates of Mycobacterium abscessus but is absent from Mycobacterium chelonae. Antimicrob Agents Chemother 2009; 53:1367.
  66. Koh WJ, Jeong BH, Kim SY, et al. Mycobacterial Characteristics and Treatment Outcomes in Mycobacterium abscessus Lung Disease. Clin Infect Dis 2017; 64:309.
  67. Choi H, Jhun BW, Kim SY, et al. Treatment outcomes of macrolide-susceptible Mycobacterium abscessus lung disease. Diagn Microbiol Infect Dis 2018; 90:293.
  68. Koh WJ, Stout JE, Yew WW. Advances in the management of pulmonary disease due to Mycobacterium abscessus complex. Int J Tuberc Lung Dis 2014; 18:1141.
  69. Wallace RJ Jr, Swenson JM, Silcox VA, Bullen MG. Treatment of nonpulmonary infections due to Mycobacterium fortuitum and Mycobacterium chelonei on the basis of in vitro susceptibilities. J Infect Dis 1985; 152:500.
  70. Wallace RJ Jr, Tanner D, Brennan PJ, Brown BA. Clinical trial of clarithromycin for cutaneous (disseminated) infection due to Mycobacterium chelonae. Ann Intern Med 1993; 119:482.
  71. Diel R, Ringshausen F, Richter E, et al. Microbiological and Clinical Outcomes of Treating Non-Mycobacterium Avium Complex Nontuberculous Mycobacterial Pulmonary Disease: A Systematic Review and Meta-Analysis. Chest 2017; 152:120.
  72. Koh WJ, Jeong BH, Jeon K, et al. Oral Macrolide Therapy Following Short-term Combination Antibiotic Treatment of Mycobacterium massiliense Lung Disease. Chest 2016; 150:1211.
  73. Pasipanodya JG, Ogbonna D, Ferro BE, et al. Systematic Review and Meta-analyses of the Effect of Chemotherapy on Pulmonary Mycobacterium abscessus Outcomes and Disease Recurrence. Antimicrob Agents Chemother 2017; 61.
  74. Rubio M, March F, Garrigó M, et al. Inducible and Acquired Clarithromycin Resistance in the Mycobacterium abscessus Complex. PLoS One 2015; 10:e0140166.
  75. Gao YH, Guan WJ, Xu G, et al. Macrolide therapy in adults and children with non-cystic fibrosis bronchiectasis: a systematic review and meta-analysis. PLoS One 2014; 9:e90047.
  76. Serisier DJ, Martin ML, McGuckin MA, et al. Effect of long-term, low-dose erythromycin on pulmonary exacerbations among patients with non-cystic fibrosis bronchiectasis: the BLESS randomized controlled trial. JAMA 2013; 309:1260.
  77. Kwak N, Dalcolmo MP, Daley CL, et al. Mycobacterium abscessus pulmonary disease: individual patient data meta-analysis. Eur Respir J 2019; 54.
  78. Jarand J, Levin A, Zhang L, et al. Clinical and microbiologic outcomes in patients receiving treatment for Mycobacterium abscessus pulmonary disease. Clin Infect Dis 2011; 52:565.
  79. Jeon K, Kwon OJ, Lee NY, et al. Antibiotic treatment of Mycobacterium abscessus lung disease: a retrospective analysis of 65 patients. Am J Respir Crit Care Med 2009; 180:896.
  80. Rodriguez Díaz JC, López M, Ruiz M, Royo G. In vitro activity of new fluoroquinolones and linezolid against non-tuberculous mycobacteria. Int J Antimicrob Agents 2003; 21:585.
  81. Martiniano SL, Wagner BD, Levin A, et al. Safety and Effectiveness of Clofazimine for Primary and Refractory Nontuberculous Mycobacterial Infection. Chest 2017; 152:800.
  82. McGuffin SA, Pottinger PS, Harnisch JP. Clofazimine in Nontuberculous Mycobacterial Infections: A Growing Niche. Open Forum Infect Dis 2017; 4:ofx147.
  83. Yang B, Jhun BW, Moon SM, et al. Clofazimine-Containing Regimen for the Treatment of Mycobacterium abscessus Lung Disease. Antimicrob Agents Chemother 2017; 61.
  84. Kaushik A, Ammerman NC, Martins O, et al. In Vitro Activity of New Tetracycline Analogs Omadacycline and Eravacycline against Drug-Resistant Clinical Isolates of Mycobacterium abscessus. Antimicrob Agents Chemother 2019; 63.
  85. Pearson JC, Dionne B, Richterman A, et al. Omadacycline for the Treatment of Mycobacterium abscessus Disease: A Case Series. Open Forum Infect Dis 2020; 7:ofaa415.
  86. Morrisette T, Alosaimy S, Philley JV, et al. Preliminary, Real-world, Multicenter Experience With Omadacycline for Mycobacterium abscessus Infections. Open Forum Infect Dis 2021; 8:ofab002.
  87. Brown-Elliott BA, Wallace RJ Jr. In Vitro Susceptibility Testing of Tedizolid against Nontuberculous Mycobacteria. J Clin Microbiol 2017; 55:1747.
  88. Poon YK, La Hoz RM, Hynan LS, et al. Tedizolid vs Linezolid for the Treatment of Nontuberculous Mycobacteria Infections in Solid Organ Transplant Recipients. Open Forum Infect Dis 2021; 8:ofab093.
  89. Brown-Elliott BA, Wallace RJ Jr. In Vitro Susceptibility Testing of Bedaquiline against Mycobacterium abscessus Complex. Antimicrob Agents Chemother 2019; 63.
  90. Philley JV, Wallace RJ Jr, Benwill JL, et al. Preliminary Results of Bedaquiline as Salvage Therapy for Patients With Nontuberculous Mycobacterial Lung Disease. Chest 2015; 148:499.
  91. Kang N, Jeon K, Kim H, et al. Outcomes of Inhaled Amikacin-Containing Multidrug Regimens for Mycobacterium abscessus Pulmonary Disease. Chest 2021; 160:436.
  92. Henriette Zweijpfenning SM, Chiron R, Essink S, et al. Safety and Outcomes of Amikacin Liposome Inhalation Suspension for Mycobacterium abscessus Pulmonary Disease: A NTM-NET study. Chest 2022; 162:76.
  93. Jhun BW, Yang B, Moon SM, et al. Amikacin Inhalation as Salvage Therapy for Refractory Nontuberculous Mycobacterial Lung Disease. Antimicrob Agents Chemother 2018; 62.
  94. Story-Roller E, Maggioncalda EC, Cohen KA, Lamichhane G. Mycobacterium abscessus and β-Lactams: Emerging Insights and Potential Opportunities. Front Microbiol 2018; 9:2273.
  95. Pandey R, Chen L, Manca C, et al. Dual β-Lactam Combinations Highly Active against Mycobacterium abscessus Complex In Vitro. mBio 2019; 10.
  96. Dedrick RM, Guerrero-Bustamante CA, Garlena RA, et al. Engineered bacteriophages for treatment of a patient with a disseminated drug-resistant Mycobacterium abscessus. Nat Med 2019; 25:730.
  97. Ganapathy US, Dartois V, Dick T. Repositioning rifamycins for Mycobacterium abscessus lung disease. Expert Opin Drug Discov 2019; 14:867.
  98. Dedrick RM, Smith BE, Cristinziano M, et al. Phage Therapy of Mycobacterium Infections: Compassionate Use of Phages in 20 Patients With Drug-Resistant Mycobacterial Disease. Clin Infect Dis 2023; 76:103.
Topic 5347 Version 32.0

References

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