A 16‐Year‐Old with a Mycoplasma hominis Empyema Post‐Lung Transplantation: A Case Report

ABSTRACT

Background

This is the first reported case of empyema due to Mycoplasma hominis in a pediatric transplant recipient.

Methods

A 16‐year‐old Indigenous Canadian boy developed acute respiratory distress 29 days post‐bilateral lung transplantation for chronic lung disease and pulmonary hypertension secondary to extreme prematurity and an atrial septal defect. Pre‐transplant donor bronchial cultures grew Candida albicans and methicillin‐sensitive Staphylococcus aureus , so he received 14 days of cefazolin. Post‐transplant prophylaxis included azithromycin, voriconazole, micafungin, TMP‐SMX, and valacyclovir. Immunosuppression included anti‐thymocyte globulin induction, followed by tacrolimus, mycophenolate mofetil, and prednisone. The patient developed a large right pleural effusion over the course of 24 h requiring intensive care and high‐flow supplemental oxygen. Pleural thoracentesis revealed a neutrophil‐predominant exudative empyema. Routine cultures were negative; M. hominis was detected by PCR and specialized media. The patient completed 28 days of clindamycin and doxycycline and made an uneventful recovery.

Results

M. hominis and Ureaplasma species are donor‐derived pathogens that can cause significant morbidity, including sternal wound infection, mediastinitis, pericarditis, and empyema. Post‐lung transplant M. hominis infections occur in 2%–5% of cases. Diagnostic challenges, low clinical suspicion, and rising resistance contribute to poor outcomes and inappropriate antibiotic use. Although this patient's ammonia level was normal, hyperammonemia syndrome also remains a rare but serious complication of Ureaplasma urealyticum and M. hominis infections.

Conclusion

Early screening, PCR testing, and prompt combination empiric therapy are crucial for improving outcomes in M. hominis infections.

1. Background

Lung transplantation is a life‐saving treatment for patients with end‐stage lung disease [1]. Although surgical advancements have improved survival rates, infections remain a major source of morbidity and the leading cause of death within the first year after transplantation [2].

Mycoplasma hominis is a part of the normal genitourinary microbiota but can also colonize the upper respiratory tract, likely because of sexual activity [3]. In the post‐lung transplant period, infections are considered donor‐derived. The incidence of M. hominis infection post‐lung transplant has been reported to be 2%–5% in adult studies. These infections pose significant diagnostic challenges and can cause substantial morbidity [4]. Although M. hominis infections are described in adult transplant recipients [4, 5, 6, 7, 8], limited evidence in pediatric patients likely contributes to underreporting and underrecognition. To our knowledge, this is the first reported case of M. hominis empyema in a pediatric transplant recipient [8].

2. Case Description

In November 2024, a 16‐year‐old fully immunized boy from an Indigenous community in Canada with known methicillin‐resistant Staphylococcus aureus (MRSA) colonization developed acute respiratory distress 29 days after sequential bilateral lung transplantation for chronic lung disease secondary to extreme prematurity and pulmonary arterial hypertension from an atrial septal defect. He was Cytomegalovirus Donor‐/Recipient‐ and Epstein–Barr Virus Donor‐/Recipient+.

The patient had been hospitalized for 211 days pre‐transplant because of decompensated pulmonary hypertension, requiring atrial septal defect repair and extracorporeal lung bypass with an interventional lung assist device as a bridge to transplantation.

The patient's donor was a young adult who died from a traumatic event and who was sexually active. Pre‐transplant donor bronchial cultures grew Candida albicans and methicillin‐sensitive Staphylococcus aureus (MSSA), so the patient received 14 days of cefazolin post‐transplant. He also received azithromycin to prevent chronic lung allograft dysfunction, along with prophylactic voriconazole, micafungin, trimethoprim‐sulfamethoxazole, and valacyclovir. His immunosuppression consisted of anti‐thymocyte globulin induction and maintenance therapy with tacrolimus, mycophenolate mofetil, and prednisone. There was no history of travel, animals, or tuberculosis (TB) infection or exposure. Prior to his respiratory deterioration, the preceding 29 days post‐transplant were largely uneventful, and he was doing well from an infectious diseases perspective. He was admitted to the hospital with ongoing issues, including hypertension, oxygen dependency for comfort, pain, fluid, and electrolyte management.

Twelve hours after the onset of respiratory distress, he was transferred to the Pediatric Cardiac Intensive Care Unit (PCICU) after being started on piperacillin‐tazobactam. Although afebrile, he was tachypneic and required 30 L/min of high‐flow oxygen (50% FiO₂) and showed severe work of breathing with nasal flaring, subcostal retractions, and diminished air entry on the right side and both lung bases.

Laboratory tests revealed a C‐reactive protein of 145 mg/L (reference (ref): < 8 mg/L) and a white blood cell count of 22.1 × 10⁹/L (ref: 4.5–13.0 × 10⁹/L). His venous blood gas showed a pH of 7.36 (ref: 7.35–7.45), lactate of 1.4 mmol/L (ref: 0.5–2.2 mmol/L), CO₂ of 43 mmHg (ref: 35–50 mmHg), and a base excess of −1 mmol/L (ref: −4 to 1 mmol/L).

A chest radiograph from the day prior to his respiratory deterioration showed small bilateral pleural effusions (Figure 1). Upon PCICU admission, imaging revealed a large right pleural effusion, bilateral atelectasis, and patchy airspace opacities (Figure 2). A right pleural thoracentesis performed 18 h after symptom onset and 6 h after the first dose of piperacillin‐tazobactam drained 810 mL of serosanguinous cloudy fluid. Analysis revealed > 45,000 × 10⁹/L leukocytes (93% neutrophils), 45 g/L protein, and 2048 U/L lactate dehydrogenase. The Gram stain showed 3+ red blood cells and 4+ leukocytes, but no bacteria. His routine bacterial and mycobacterial cultures were ultimately negative. No acid‐fast bacilli were seen. Vancomycin was also started. The patient remained afebrile with no further respiratory deterioration.

FIGURE 1.

Chest radiograph prior to respiratory deterioration.

FIGURE 2.

Chest radiograph at time of PCICU admission.

A diagnosis of TB was considered because of the patient's Indigenous background and having lived on a Northern reserve, but the rapid symptom progression and neutrophil‐predominant pleural fluid made this unlikely. Fungal etiologies were deemed less likely because of his antifungal prophylaxis. The pleural fluid parameters were consistent with empyema, but the absence of bacteria on Gram stain raised suspicion for atypical organisms, such as donor‐derived Mycoplasma and Ureaplasma species, which lack a cell wall. Polymerase chain reaction (PCR) and cultures were requested for M. hominis and Ureaplasma species on the pleural fluid, and clindamycin and doxycycline were added 12 h after PCICU transfer. Serum ammonia was normal on day 2 of PCICU admission and was not repeated as there were never concerns about the patient's neurologic status. On day 6 of PCICU admission, PCR results on pleural fluid detected M. hominis , which was later cultured on specialized media. Susceptibility testing (CLSI M43 broth microdilution method) from the University of Alabama Diagnostic Mycoplasma Laboratory [9], available 21 days into treatment, confirmed sensitivity to clindamycin, tetracycline, and moxifloxacin.

The patient recovered uneventfully and completed 28 days of clindamycin and doxycycline. During the first week of infection, he required treatment for antibody‐mediated rejection with plasmapheresis, intravenous immunoglobulin (IVIG), and a dose of rituximab. Pulse methylprednisolone was given over 3 days, followed by a steroid wean with prednisone, further increasing his net state of immunosuppression. He was discharged from the hospital 79 days post‐transplant.

3. Discussion

Infections are a significant source of morbidity and the leading cause of mortality within the first year post‐lung transplant [2]. Pleural effusions occur in approximately 25% of patients within the first 90 days following transplantation, typically resulting from increased alveolar permeability, perioperative pleural inflammation, postoperative atelectasis, and impaired lymphatic drainage [10]. Among these cases, 3%–8% develop empyema [11], a serious complication that can lead to systemic infection, decrease one‐year survival rates [12] and require aggressive treatment [11].

Diagnosing empyemas in lung transplant recipients is particularly challenging because of muted symptoms caused by immunosuppression [11]. Additionally, post‐transplant pleural effusions often resemble empyemas biochemically, even when sterile [11]. Traditional pleural markers used to distinguish transudative from exudative effusions are less reliable in the transplant population, as most early post‐transplant effusions are exudative by nature [10]. A pleural fluid neutrophil count of greater than 21% has been shown to differentiate infected from noninfected effusions, with 70% sensitivity and 79% specificity [11]. However, noninfected effusions can also present with neutrophilic or lymphocytic predominance [10], highlighting the importance of obtaining cultures early on in the diagnostic process [10].

Independent patient‐related risk factors for empyema include diabetes, prior cardiothoracic surgery, female donor, prolonged ischemic time, and the number of perioperative red blood cell transfusions [13].

Most post‐transplant empyemas are monomicrobial, with fungal pathogens, particularly Candida albicans, being most common [11]. Polymicrobial infections occur in approximately 14.3% of cases [11].

Donor‐derived infections involving M. hominis and Ureaplasma infections have emerged as a notable cause of morbidity. These organisms can lead to bronchial anastomotic infection, sternal wound infection, mediastinitis, pericarditis, and empyema [5]. Colonization rates vary among studies; however, M. hominis is a highly prevalent urogenital organism and can also colonize the upper respiratory tract, likely associated with sexual activity [6, 14]. Colonization rates among donors also vary. In one retrospective review from 2015 to 2019, out of 105 donors screened with PCR, 12 (11.4%) tested positive for M. hominis and Ureaplasma infections [15]. In a prospective screening study at a single site from 2020 to 2021, 18% (18 of 99 donors) of post‐lung transplant bronchoalveolar lavage (BAL) fluid samples were positive on culture or PCR for these species [4]. There are no pediatric data, but the incidence of M. hominis infection post‐lung transplant in adults ranges from 2% to 5% [4, 5, 16]. Detection rates may be increasing because of greater awareness, more specific testing, and a rising prevalence of colonization in the general population [5, 17]. Donor risk factors for Mycoplasma and Ureaplasma infections in lung transplant recipients include young age and high‐risk sexual activity [3]. Clinical manifestations typically appear a median of 19 days (range, 4–90) post‐transplant [15].

Delays in diagnosing M. hominis and Ureaplasma species post‐lung transplant are common, especially if the diagnosis is not considered, leading to poor outcomes and inappropriate antibiotic use [6]. These fastidious organisms lack a cell wall, rendering them undetectable by Gram stain. Additionally, M. hominis is fragile and sensitive to drying, temperature fluctuations, and toxic metabolites, so it can easily die during transport to reference laboratories, complicating speciation and susceptibility testing [6].

Selective culture media increase yield, but cultures are labor‐intensive and costly.

Generally, growth of Mycoplasmas species pathogenic for humans requires the presence of serum and growth factors such as yeast extract [18, 19]. No single formulation is ideal for all species because of different properties, optimum pH, and substrate requirements [18, 19]. Broth and agar are usually incubated for 7 days [5, 20]. Overgrowth from competing bacteria and the pinpoint nature of M. hominis colonies contribute to the organism being easily missed by laboratory staff [21]. Agar plates should be examined using a stereomicroscope so as not to miss colonies [18, 21]. At our center, respiratory Mycoplasma agar in scintillation vials, respiratory Mycoplasma agar plate (RM agar), respiratory glucose broth, bromothymol blue broth with arginine (B'arg broth), bromothymol blue broth with urea (B broth), and genital Mycoplasma agar (GM agar) were used (M. Diggle, personal communication, February 7, 2025). Diagnostic yield greatly increases with selective Mycoplasma culture media and molecular testing, which have high sensitivity and specificity [21]. For M. hominis PCR testing, samples are sent out to our national reference laboratory, the National Microbiology Laboratory in Winnipeg, Canada, where the Allplex STI Essential Assay Q on the Seegene platform is most commonly used [22].

Treatment of M. hominis remains challenging because of its lack of a cell wall, making it resistant to beta‐lactams and vancomycin, which are often a part of routine post‐transplant antibiotic regimens. Additionally, M. hominis is resistant to macrolides, aminoglycosides, and TMP‐SMX. Protein synthesis inhibitors such as doxycycline, moxifloxacin, and clindamycin are commonly used; however, resistance to these agents is rising. In the context of limited antimicrobial choice, antibiotic resistance is a large concern. A German study showed resistance to tetracyclines increased from 2.1% in 1983 to 14.5% in 2004, and to fluoroquinolones from 0% to 15.4% [23]. PCR‐based assays have been used to identify key antimicrobial resistance genes in M. hominis and Ureaplasma species [24]. Treatment generally involves combination therapy for 2–6 weeks, depending on the clinical syndrome [5]. Dual antibiotics are recommended, given the delayed susceptibility results and significant resistance rates. The incidence of sequelae or of recurrent disease is not known.

Our local protocol includes PCR for M. hominis and Ureaplasma species on routine BAL 1–3 days post‐transplant [7]; however, this test was unfortunately missed in this patient. If positive, doxycycline prophylaxis is given for 7–14 days. The choice of antibiotics is based on increasing gyrA and gyrB fluoroquinolone resistance and M. hominis ' resistance to macrolides [7].

Hyperammonemia syndrome (HS) is a serious complication marked by encephalopathy and elevated serum ammonia levels. It affects up to 5% of lung transplant recipients with M. hominis or Ureaplasma infection, with a mortality rate of up to 80% [4]. HS was previously thought to result from an inborn error of urea metabolism, unmasked by calcineurin inhibition [3]. However, recent evidence has shown a strong association with primarily Ureaplasma urealyticum infections, as well as M. hominis [3]. Targeted antibiotic therapy can reverse the clinical syndrome [3].

Although this patient's ammonia level remained normal, HS continues to be a rare but potentially fatal complication of Ureaplasma urealyticum and M. hominis infections. Early monitoring post‐transplant of ammonia levels is standard practice at our centre. Any unexplained sudden neurological change, such as lethargy or severe encephalopathy, should prompt ammonia‐level assessment in any immunocompromised patient because of their heightened risk for these infections [7].

4. Conclusions

Following lung transplantation, M. hominis and Ureaplasma are considered donor‐derived infections and can lead to serious complications if not promptly treated. Early diagnosis is challenging because of the difficulty of culturing and identifying these organisms, but PCR testing can significantly improve detection. Known donor‐related risk factors include young age and high‐risk sexual history.

In cases of unexplained respiratory distress and effusion after lung transplantation, clinicians should maintain a high index of suspicion for M. hominis or Ureaplasma infections. Early empiric therapy and serum ammonia testing are crucial to minimize complications. Given the diagnostic challenges, early screening, targeted testing, and prompt combination therapy are essential for improving patient outcomes. With the wide range of practice variations in the screening of M. hominis and Ureaplasma infections post‐lung transplant, we suggest that transplant societies develop guidelines to standardize screening, prophylaxis, as well as diagnostic and treatment approaches across centers.

Author Contributions

All authors contributed to the patient's care and conceived the report. All authors reviewed the manuscript, provided critical feedback, finalized, and approved the final manuscript.

Conflicts of Interest

Dr. Joan Robinson has received honoraria paid by Alberta Pharmacists' Association, Alberta ID Conference, and Canadian Pediatric Society for talks on RSV and on Pneumonia in 2023 and 2024. Dr. Catherine Burton has research grant funding for a variety of studies related to vaccination from the Public Health Agency of Canada, Canadian Donation and Transplantation Research Program, Canadian Institutes of Health Research, Canadian Immunity Task Force, Women and Children's Health Research Institute, and Moderna. Dr. Dima Kabbani has received research grants and conducted clinical trials with/from: AVIR Pharma, PULMOCIDE, F2G, and Moderna. Dr. Geraldine Huynh declares no conflicts of interest.

Huynh G., Burton C., Kabbani D., and Robinson J., “A 16‐Year‐Old with a Mycoplasma hominis Empyema Post‐Lung Transplantation: A Case Report,” Pediatric Transplantation 29, no. 7 (2025): e70214, 10.1111/petr.70214.

Data Availability Statement

Data sharing is not applicable to this article.