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While echinocandins remain the preferred antifungal therapy for candidemia, bloodstream infections caused by C. auris, C. glabrata, or, at a lesser extent, C. parapsilosis may be difficult to treat with these antifungal agents. This is in view of the high propensity of the above-mentioned species to develop antifungal resistance or tolerance during treatment.Candida auris and other Candida species (C. albicans, C. glabrata, C. parapsilosis, C. tropicalis, and C. krusei) are important causes of bloodstream infection. Early or prolonged treatment with antifungal agents is often required. The inhibitory effect of antifungal agents in the patients' bloodstream may compromise the sensitivity of blood culture (BC) to diagnose and/or monitor patients with candidemia. Using a clinical BC simulation model, we compared antimicrobial drug-neutralizing BC media in BacT/Alert FA PLUS (FAP) or Bactec Plus Aerobic/F (PAF) bottles with non-neutralizing BC media in Bactec Mycosis IC/F (MICF) bottles to allow Candida growth in the presence of 100%, 50%, or 25% peak serum level (PSL) antifungal concentrations. In total, 117 organism/antifungal combinations were studied, and Candida growth was detected after incubating bottles into BacT/Alert VIRTUO or Bactec FX BC systems. Compared to control (without antifungal) bottles, both FAP and PAF bottles with 100% PSL antifungal concentrations allowed 100% recovery for C. auris, C. glabrata, and C. parapsilosis, whereas recovery was below 100% for C. albicans, C. krusei, and C. tropicalis. MICF bottles were less efficient at 100%, 50%, or 25% PSL antifungal concentrations, for all Candida species, except for C. auris. While azoles and amphotericin B did not hinder Candida growth in FAP or PAF bottles, echinocandins allowed C. auris, C. glabrata, and C. parapsilosis to grow in FAP, PAF, or MICF bottles. Overall, the maximum time to detection was 4.6 days. Taken together, our findings emphasize the reliability of BCs in patients undergoing anti

In keeping with the evolution of severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), the COVID-19 causative agent, PCR assays have been developed to rapidly detect SARS-CoV-2 variants, which have emerged since the first (Alpha) variant was identified. Based on specific assortment of SARS-CoV-2 spike-protein mutations (DH69/V70, E484K, N501Y, W152C, L452R, K417N, and K417T) among the major variants known to date, Seegene Allplex SARS-CoV-2 Variants I and Variants II assays have been available since a few months before the last (Omicron) variant became predominant. Using S gene next-generation sequencing (NGS) as the SARS-CoV-2 variant identification reference method, we assessed the results of SARS-CoV-2-positive nasopharyngeal swab samples from two testing periods, before (n = 288, using only Variants I) and after (n = 77, using both Variants I and Variants II) the appearance of Omicron. The Variants I assay allowed correct identification for Alpha (37/37), Beta/Gamma (28/ 30), or Delta (220/221) variant-positive samples. The combination of the Variants I and Variants II assays allowed correct identification for 61/77 Omicron variant-positive samples. While 16 samples had the K417N mutation undetected with the Variants II assay, 74/77 samples had both DH69/V70 and N501Y mutations detected with the Variants I assay. If considering only the results by the Variants I assay, 6 (2 Beta variant positive, 1 Delta variant positive, and 3 Omicron variant positive) of 365 samples tested in total provided incorrect identification. We showed that the Variants I assay alone might be more suitable than both the Variants I and Variants II assays to identify currently circulating SARS-CoV-2 variants. Inclusion of additional variant-specific mutations should be expected in the development of future assays.

We used nasopharyngeal swab samples of patients with a symptomatic (n = 82) or asymp-tomatic (n = 20) coronavirus disease 2019 (COVID-19) diagnosis to assess the ability of antigen detection tests to infer active (potentially transmissible) or inactive (potentially non-transmissible) infection by severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2). Using the subgenomic RNA (sgRNA) as an active replication marker of SARS-CoV-2, 48 (76.2%), 56 (88.9%), and 63 (100%) of 63 samples with sgRNA positive results tested positive with the SD BIOSENSOR STANDARD Q COVID-19 Ag (Standard Q), the SD BIOSENSOR STANDARD F COVID-19 Ag FIA (Standard F), or the Fujirebio LUMIPULSE G SARS-CoV-2 Ag (Lumipulse) assay, respectively. Conversely, 37 (94.9%), 29 (74.4%), and 7 (17.9%) of 39 samples with sgRNA negative results tested negative with Standard Q, Standard F, or Lumipulse, respectively. Stratifying results by the number of days of symptoms before testing revealed that most antigen positive/sgRNA positive results were among samples tested at 2–7 days regardless of the assay used. Conversely, most antigen negative/sgRNA negative results were among samples tested at 16–30 days only when Standard Q or Standard F were used. In conclusion, based on our findings, a negative antigen test, especially with the Lumipulse assay, or a positive antigen test, especially with the Standard F assay, may suggest, respectively, the absence or presence of replication-competent SARS-CoV-2.

We examined 29 autopsy cases (investigated between October 2020 and February 2021) whose postmortem swabs tested positive for SARS-CoV-2. Twenty-two of 29 cases died while hospitalized (H), while the remaining 7 cases were not hospitalized (NH). Since we included only cases in which the time since death was known (excluding unwitnessed NH deaths), the interval between death and postmortem swab(s) was registered, with a mean NH value of 5.50 days and a mean H value of 3.98 days. The mean age of NH was 65 years, while H were older (mean age: 73 years). Twenty-eight nasopharyngeal and 27 lungs postmortem swabs were obtained and real-time reverse transcriptase‒polymerase chain reaction assay for total and replicative SARS-CoV-2 RNA and mRNA detection was performed. Although the mean death-postmortem swabs interval was higher in NH than in H, the mean viral load of NH was higher than that of H (2.53 × 1011 copies/mL vs 9.31 × 108 copies/mL). In 13/29 cases (6 NH and 7 H), indicators of active replication were found. The relationship between the presence of replicative mRNA and death without hospitalization and that between the minimum cycle threshold value of SARS-CoV-2 RNA and the cycle threshold value of replicative SARS-CoV-2 mRNA were found to be statistically significant (with respective P values of 0.013 and 0.000). Therefore, especially in NH, full compliance with guidelines on biological safety in the autopsy room is essential, and no autopsy can be performed on infected cases in a structure that does not meet the established safety criteria.

We aimed to report a 32-month laboratory experience with the eazyplex® CSF direct panel assay for the rapid diagnosis of meningitis due to six most common bacterial species (Escherichia coli, Haemophilus influenzae, Listeria monocytogenes, Neisseria meningitidis, Streptococcus agalactiae, and Streptococcus pneumoniae). We included all cerebrospinal fluid (CSF) samples from patients admitted with a clinical suspicion of meningitis/encephalitis between May 2016 and December 2018 at our hospital. In addition to the eazyplex® assay, both Gram stain microscopy and culture were performed, and results were confirmed with 16S rRNA PCR/sequencing. Patients’ demographics and relevant clinical information were collected. Of 135 studied patients, 44 (32.6%) had a microbiologically documented diagnosis of meningitis. Overall, we identified 21 S. pneumoniae, 10 N. meningitidis, 6 L. monocytogenes, 3 E. coli, 2 Streptococcus pyogenes, 1 S. agalactiae, and 1 Citrobacter koseri as aetiological agents. The eazyplex® assay allowed identification in 40 (90.9%) cases, with four not identified cases due to microorganisms not included in the panel at the time of testing. Thirty-two (72.7%) cases had positive culture results, whereas 28 (63.6%) cases had positive Gram stain results. Notably, combining Gram stain and eazyplex® assay allowed identification in 100% of cases. After notification of rapid results, physicians modified the empiric antibiotic therapy, which became appropriate in three patients (all with L. monocytogenes meningitis). The eazyplex® CSF panel assay worked better than culture in detecting the most common agents of bacterial meningitis and accelerated the diagnosis leading to timely initiation or continuation of appropriate antibiotic therapy.

Objectives: Compared to RT-PCR, lower performance of antigen detection assays, including the Lumipulse G SARS-CoV-2 Ag assay, may depend on specific testing scenarios. Methods: We tested 594 nasopharyngeal swab samples from individuals with COVID-19 (RT-PCR cycle threshold [Ct] values ≤ 40) or non-COVID-19 (Ct values > 40) diagnoses. RT-PCR positive samples were assigned to diagnostic, screening, or monitoring groups of testing. Results: With a limit of detection of 1.2 × 104 SARS-CoV-2 RNA copies/ml, Lumipulse showed positive percent agreement (PPA) of 79.9% (155/194) and negative percent agreement of 99.3% (397/400), whereas PPAs were 100% for samples with Ct values of <18 or 18–<25 and 92.5% for samples with Ct values of 25–<30. By three groups, Lumipulse showed PPA of 87.0% (60/69), 81.1% (43/53), or 72.2% (52/72), respectively, whereas PPA was 100% for samples with Ct values of <18 or 18–<25, and was 94.4, 80.0, or 100% for samples with Ct values of 25–<30, respectively. Additional testing of RT-PCR positive samples for SARS-CoV-2 subgenomic RNA showed that, by three groups, PPA was 63.8% (44/69), 62.3% (33/53), or 33.3% (24/72), respectively. PPAs dropped to 55.6, 20.0, or 41.7% for samples with Ct values of 25–<30, respectively. All 101 samples with a subgenomic RNA positive result had a Lumipulse assay’s antigen positive result, whereas only 54 (58.1%) of remaining 93 samples had a Lumipulse assay’s antigen positive result. Conclusions: Lumipulse assay was highly sensitive in samples with low RT-PCR Ct values, implying repeated testing to reduce consequences of false-negative results.

The bacteremia level as well as the administration of antibiotics before blood collection may significantly affect the recovery of bacterial pathogens from pediatric blood cultures in BacT/Alert Virtuo or Bactec FX BC systems, which remain the common techniques to diagnose bacteremia in pediatric patients. We simulated pediatric blood cultures with low or intermediate bacteremia level to evaluate BacT/Alert PF Plus and Bactec Peds Plus blood culture bottles for resin-based inactivation of 16 antibiotic–bacterium combinations. Overall, 105/192 (54.7%) of BacT/Alert PF Plus bottles and 69/192 (36.0%) of Bactec Peds Plus bottles allowed organisms to grow when exposed to antibiotics. In particular, both BacT/Alert PF Plus and Bactec Peds Plus bottles proved to be effective with piperacillin/tazobactam and Pseudomonas aeruginosa or with oxacillin and methicillin-susceptible Staphylococcus aureus (100% growth), whereas no effectiveness was apparent with ceftriaxone and Escherichia coli, Streptococcus agalactiae, or Streptococcus pneumoniae or with cefepime and E. coli (0% growth). In some relevant instances (e.g., with vancomycin and methicillin-resistant S. aureus or Streptococcus pneumoniae), BacT/Alert PF Plus bottles were superior to Bactec Peds Plus bottles. Together, these findings underscore the potentiality of resin-containing bottles to enhance diagnosis of bacteremia in pediatric patients on antimicrobial therapy. This is particularly true with one of the evaluated BC systems and with simulated intermediate bacteremia level only.

Bacterial pneumonia is a challenging coronavirus disease 2019 (COVID-19) complication for intensive care unit (ICU) clinicians. Upon its implementation, the FilmArray pneumonia plus (FA-PP) panel’s practicability for both the diagnosis and antimicrobial therapy management of bacterial pneumonia was assessed in ICU patients with COVID-19. Respiratory samples were collected from patients who were mechanically ventilated at the time bacterial etiology and antimicrobial resistance were determined using both standard-of-care (culture and antimicrobial susceptibility testing [AST]) and FA-PP panel testing methods. Changes to targeted and/or appropriate antimicrobial therapy were reviewed. We tested 212 samples from 150 patients suspected of bacterial pneumonia. Etiologically, 120 samples were positive by both methods, two samples were culture positive but FA-PP negative (i.e., negative for on-panel organisms), and 90 were negative by both methods. FA-PP detected no culture-growing organisms (mostly Staphylococcus aureus or Pseudomonas aeruginosa) in 19 of 120 samples or antimicrobial resistance genes in two culture-negative samples for S. aureus organisms. Fifty-nine (27.8%) of 212 samples were from empirically treated patients. Antibiotics were discontinued in 5 (33.3%) of 15 patients with FA-PP-negative samples and were escalated/deescalated in 39 (88.6%) of 44 patients with FA-PP-positive samples. Overall, antibiotics were initiated in 87 (72.5%) of 120 pneumonia episodes and were not administered in 80 (87.0%) of 92 nonpneumonia episodes. Antimicrobial-resistant organisms caused 78 (60.0%) of 120 episodes. Excluding 19 colistin-resistant Acinetobacter baumannii episodes, AST confirmed appropriate antibiotic receipt in 101 (84.2%) of 120 episodes for one or more FA-PP-detected organisms. Compared to standard-of-care testing, the FA-PP panel may be of great value in the management of COVID-19 patients at risk of developing bacterial pneumonia in the ICU. IMPORTANCE Sinc

To support the clinical laboratory diagnosis of Pneumocystis jirovecii (PJ) pneumonia (PCP), an invasive fungal infection mainly occurring in HIV‐negative patients, in‐house or commercial PJ‐specific real‐time quantitative PCR (qPCR) assays are todays’ reliable options. The performance of these assays depends on the type of PJ gene (multi‐copy mitochondrial versus single‐copy nuclear) targeted by the assay. We described the development of a PJ‐PCR assay targeting the dihydrofolate reductase (DHFR)‐encoding gene. After delineating its analytical performance, the PJ‐PCR assay was used to test bronchoalveolar lavage (BAL) fluid samples from 200 patients (only seven were HIV positive) with suspected PCP. Of 211 BAL fluid samples, 18 (8.5%) were positive and 193 (91.5%) were negative by PJ‐PCR. Of 18 PJ‐PCR‐positive samples, 11 (61.1%) tested positive and seven (38.9%) tested negative with the immunofluorescence assay (IFA). All (100%) of the 193 PJ‐PCR‐negative samples were IFA negative. Based on IFA/PCR results, patients were, respectively, classified as having (n = 18) and not having (n = 182) proven (PJ‐ PCR+/IFA+) or probable (PJ‐PCR+/IFA−) PCP. For 182 patients without PCP, alternative infectious or non‐infectious etiologies were identified. Our PJ‐PCR assay was at least equivalent to IFA, fostering studies aimed at defining a qPCR‐based standard for PCP diagnosis in the future.

This study evaluated and compared the accuracy of BCID2 with that of BCID to identify bacterial species and relative antimicrobial resistance genes directly from positive BCs. We used archived samples from positive BCs (1.5 ml thereof mixed with 100 ml of dimethyl sulfoxide and stored at 280°C), which had prospectively been processed with the BD Bactec 9240 (BD Diagnostic Systems, Sparks, MD), BacTAlert 3D (bioMérieux), or BacTAlert Virtuo (bioMérieux) system at two hospital microbiology laboratories from January 2018 to August 2020.

Sir, Recently, EUCAST developed a rapid antimicrobial susceptibility testing (RAST) method for direct use on positive blood culture (BC) samples, which relies on EUCAST standard disc diffusion (DD) methodology and allows inhibition zone reading within 4–8 h of BC positivity.We herein report on EUCAST RAST results for BCs consecutively collected between June 2019 and December 2019 that contained E. coli (n = 134)or K. pneumoniae (n = 66) isolates.

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