Clinical laboratory workers encounter a variety of occupational hazards, including exposure to infectious agents. The routes of pathogen exposure associated with laboratory work include ingestion, inhalation, direct inoculation, and contamination of skin and mucous membranes (1). The accidental inoculation of infectious materials (i.e., via contaminated needles, broken glass, or other sharps) is the leading cause of laboratory-associated infections (1).
In the 1980s, the emergence of the HIV epidemic created an appreciation for biosafety and laboratory-acquired infections, ultimately leading to both practice guidelines and legislation to reduce the risk of exposure of laboratory workers to bloodborne pathogens; this was implemented by the adoption of Universal Precautions in 1987, and later, in 1996, these became a component of Standard Precautions (2). The actual incidence and risk of laboratory-acquired infections is very difficult to quantify in the absence of standardized reporting systems, as well as the challenge of attributing a specific exposure to acquisition of infection. The most comprehensive studies to date attempting to estimate the incidence and epidemiology of laboratory-acquired infections were completed prior to 1980, and it is difficult to generalize them to today's laboratory environment, for a variety of reasons—standards for personal protective equipment have changed, the availability of vaccines, the epidemiology of bloodborne infections has changed, and the way laboratory testing is performed has changed (with a shift away from primarily manual methods to more automated methods). That said, since 1999, only 1 confirmed case of laboratory-acquired HIV infection has been reported in the US; this case was secondary to a needle stick injury sustained in an individual working with a live HIV culture (2).
The fear associated with the recent Ebola virus (EV)3 epidemic triggered a renewed interest in occupationally acquired infections in healthcare workers in the US, including the safety of laboratory workers in handling samples from persons under investigation (PUIs) for EV disease (EVD). Individuals at risk for EVD are also at risk for several other infectious diseases with overlapping symptom profiles (such as malaria, influenza, and bacteremia) thus obligating a number of diagnostic laboratory tests. In addition, the clinical management of patients with EVD requires ongoing laboratory testing to optimize care (such as complete blood count, coagulation testing, electrolyte analysis, etc.) (3). Laboratory testing for suspect or confirmed EVD patients is unfamiliar to most healthcare workers in the US, and thus determining the safest approach to this testing generated anxiety and controversy.
The CDC recommended that laboratories perform a risk assessment to minimize the risk to laboratory staff, and determine what equipment would be available or needs to be acquired for providing critically important testing (4). The infectious dose for EV is estimated to be 10 viral particles, and patients with EVD may have EV viral loads of ≥108 plaque-forming units/mL (5). Thus, in the absence of data regarding the potential risk of laboratory testing using automated analyzers, many laboratories have opted to utilize point-of-care testing devices or conduct testing for PIUs in a separate lab, outside of the routine laboratory workflow (6). Even among designated US Ebola Treatment Centers, the approach to laboratory testing is highly variable, with some centers performing testing in a Biosafety Level-3 laboratory, some within a patient isolation unit, and some within the clinical laboratory of the hospital (6).
In this issue of Clinical Chemistry, to gather data on the frequency of contamination of laboratory equipment with blood-borne pathogens, Bryan et al. conducted an important evaluation of total laboratory automation (TLA) used in the routine clinical laboratory. Quantitative PCR assays were used to evaluate the extent of TLA contamination after blood samples were processed through routine workflow (7). Because it was not practical to test EV directly, contamination with hepatitis B virus (HBV) and hepatitis C virus (HCV) were used as surrogates to measure contamination events (7). To establish a baseline level of instrument contamination, the authors evaluated numerous components of the TLA at multiple time points. Out of 79 baseline swab samples, 10 (12.6%) and 8 (10.1%) were positive for HBV and HCV, respectively. The contaminated sites were associated with visible flecks of dried blood, and generally located near the decapper portion of the instrument. The authors then tested TLA surfaces and clean glass slides placed throughout the TLA after sample processing of high titer HCV samples (mean of 5.8 × 107 IU/mL). The purpose of the clean glass slides was to directly assess contamination events specifically attributed to the high titer HCV samples, thereby eliminating detection of baseline contamination events. The authors detected HCV contamination on 1 glass slide out of the 54 that were tested. Of note, although HBV and HCV were detected in this study, it is unknown if the viral particles were still viable/infectious. No contamination events were identified when sites away from the TLA were sampled, indicating that cross-contamination from the TLA to other sites did not occur in the sites examined. Although the authors examined only HBV and HCV, we would anticipate similar contamination patterns with other bloodborne pathogens, such as EV.
The results from this important study reveal that background contamination of TLA with HBV and HCV occurs during routine clinical use. This is an important finding, especially considering HBV viability in dried blood has been demonstrated for up to 7 days at room temperature (1) and HBV can be present in very high concentrations in clinical samples (up to 109 IU/mL). Although HBV was the most commonly reported laboratory-acquired infection, the incidence of such infections has decreased drastically in the era of standard precautions and HBV vaccination (8). Additionally, although HCV seropositivity is slightly higher in healthcare workers compared to the general public, laboratory-acquired HCV infection appears to be rare, with only single case reports (8–10).
The evolving guidance from the CDC on the testing of PUIs for EVD stresses the importance of performing a risk assessment to identify potential exposure sources and to mitigate those events (4). It should be emphasized that infection with EV, similar to HIV, HBV, and HCV, requires direct exposure to EV-contaminated blood or bodily fluids, so proper personal protective equipment is essential when handling clinical samples (4). Additionally, the CDC recommendations state that laboratories should minimize processes that would generate aerosols that would expose laboratory staff to EV, by the use of engineering controls and safety equipment when available (4). There are several reports of hospitalized patients in the Netherlands (11), the US (12), and South Africa (13) with initially undiagnosed EV or the related Marburg virus infections that had extensive contact with healthcare workers, including laboratory staff working with clinical samples from these patients. In these reports, none of the healthcare workers contracted EV or Marburg virus from these patients, which could be attributed to adherence to proper use of personal protective equipment and standard precautions.
It is incumbent on healthcare facilities to provide the tools and training to be able to properly evaluate PUIs for EVD, as well as other infections that could mimic EVD (i.e., meningitis, malaria). These other infections are potent and can be deadly if they are not treated promptly and properly. To this point, a recent report describes 3 instances between 2014–2015, in which malaria testing was significantly delayed in PUIs for EVD (14). One patient was initially empirically treated for malaria without having had malaria testing (which is counter to CDC recommendations), and in another patient the level of parasitemia was not evaluated, which is an important factor in determining which antimalarial treatment should be administered.
Importantly, the investigation by Bryan et al. (7) illustrates that contamination events do occur with pathogens (i.e., HBV and HCV) that are frequently encountered, likely on a daily basis. Although contamination of laboratory equipment with bloodborne pathogens may be common, laboratory-acquired infection with these agents is not. This underscores the importance of adherence to standard precautions when handling all patient samples, and treating every clinical sample as though it may contain an infectious agent.
Important questions for clinical laboratories that are not addressed by this investigation include what methods are needed to adequately decontaminate laboratory equipment, the frequency with which these methods could or should be deployed, and the risk that these methods may pose to laboratory workers. Laboratories should work closely with device manufacturers to understand best practices for instrument decontamination.
EV and other emerging pathogens will continue to be encountered in the clinical laboratory. It is the joint responsibility of laboratorians and laboratory leadership to create a culture of safety and adherence to safety protocols, which are essential to reduce the risk of laboratory-acquired infections.
↵3 Nonstandard abbreviations:
- Ebola virus;
- persons under investigation;
- Ebola virus disease (EVD);
- total laboratory automation;
- hepatitis B virus;
- hepatitis C virus.
(see article on page 973)
Author Contributions: All authors confirmed they have contributed to the intellectual content of this paper and have met the following 3 requirements: (a) significant contributions to the conception and design, acquisition of data, or analysis and interpretation of data; (b) drafting or revising the article for intellectual content; and (c) final approval of the published article.
Authors' Disclosures or Potential Conflicts of Interest: No authors declared any potential conflicts of interest.
- Received for publication April 26, 2016.
- Accepted for publication April 26, 2016.
- © 2016 American Association for Clinical Chemistry