Abstract
Legionella spp. are the causative agent of Legionnaire’s disease and an opportunistic pathogen of significant public health concern. Identification and quantification from environmental sources is crucial for identifying outbreak origins and providing sufficient information for risk assessment and disease prevention. Currently there are a range of methods for Legionella spp. quantification from environmental sources, but the two most widely used and accepted are culture and real-time polymerase chain reaction (qPCR). This paper provides a review of these two methods and outlines their advantages and limitations. Studies from the last 10 years which have concurrently used culture and qPCR to quantify Legionella spp. from environmental sources have been compiled. 26/28 studies detected Legionella at a higher rate using qPCR compared to culture, whilst only one study detected equivalent levels of Legionella spp. using both qPCR and culture. Aggregating the environmental samples from all 28 studies, 2856/3967 (72%) tested positive for the presence of Legionella spp. using qPCR and 1331/3967 (34%) using culture. The lack of correlation between methods highlights the need to develop an acceptable standardized method for quantification that is sufficient for risk assessment and management of this human pathogen.
Introduction
Legionella is a genus of opportunistic pathogens of significant public health concern (Borges et al., Citation2012). It is the causative agent of Legionellosis, which collectively refers to Legionnaires disease and Pontiac fever (Fields et al., Citation2002). Legionnaires disease is a serious atypical bacterial pneumonia; whereas Pontiac fever is a self-limiting febrile illness (Neil & Berkelman, Citation2008). There have been no reports of human to human transmission of Legionella spp. (Khweek et al., Citation2013) and inhalation or aspiration of contaminated aerosols is the most commonly accepted mechanism of infection (Cianciotto, Citation2001). Community and nosocomial cases of Legionellosis are typically associated with cooling towers (Nhu Nguyen et al., Citation2006), hot water systems (Goetz et al., Citation1998; Leoni et al., Citation2005), potable water (Stout, Citation1992), spa pools (Benkel et al., Citation2000), decorative water fountains (Haupt et al., Citation2012) and potting mix (O’Connor et al., Citation2007).
Legionella spp. are difficult to control in environmental sources due to their resistance to disinfectants (Kim et al., Citation2002), association with biofilms (Murga et al., Citation2001) and parasitism of protozoan hosts (Thomas et al., Citation2004). Environmental surveillance and monitoring of Legionella spp. is crucial for evaluating risk and identifying control strategies (Cristino et al., Citation2012). This requires a quick and accurate method for detecting and enumerating Legionella spp. in environmental sources (Declerck et al., Citation2006).
Currently there are several methods for Legionella spp. detection and enumeration including: culture (Bopp et al., Citation1981); PCR (Mahbubani et al., Citation1990); qPCR (Behets et al., Citation2007); Fluorescent in situ hybridization (FISH) (Deloge Abarkan et al., Citation2007); solid phase cytometry (Aurell et al., Citation2004); optical wavelight spectroscopy (Cooper et al., Citation2009); Enzyme-Amplified Electrochemical Detection with DNA probe (Miranda-Castro et al., Citation2007) and Surface plasmon resonance immunosensor (Oh et al., Citation2003). Many national bodies have adopted guidelines which use culture methods as the standard; however, the development of more rapid techniques highlights the need for reassessment of guidelines. This review will compare culture and qPCR, which are the two methods which have gained prominence as the most widely used and accepted by analytical labs (Krøjgaard et al., Citation2011; Lee et al., Citation2011).
Detection of Legionella by culture methods
Due to the microbial complexity of environmental samples, isolating Legionella spp. by culture methods presents a range of challenges, which have been addressed by the development of specific agar formulations and sample treatments (Bopp et al., Citation1981). Sample collection protocols, sampling location and storage will not be specifically addressed in this manuscript; however, in its own right the sampling method used may exert a significant impact upon the likelihood of detection Legionella spp. in the environment (Asadi et al., Citation2011; CDC, Citation2005).
The requirement for sample treatment is generally contingent on the properties of the environmental source. Potable water and water from hot water reservoirs often requires less destructive sampling techniques than samples collected from cooling towers and potting mix as they are generally less microbially complex (Joly et al., Citation2006; Steele et al., Citation1990). For potable water, it may be sufficient to simply filter concentrated 1 L of water to ensure that sufficient microbial flora is present before plating (Fiume et al., Citation2005). Samples collected from chlorine treated water sources should be dosed with 0.5 ml of 0.1 N sodium thiosulfate per 1 L to neutralize residual disinfectants (CDC, Citation2005).
Samples from cooling towers, potting mix, reuse water distribution systems and non-potable sources generally require either heat or acid treatment to reduce the microbial content of the sample before plating (Bopp et al., Citation1981). As Legionella is thermally tolerant up to 63 °C (Fliermans et al., Citation1981) samples may be heat treated to reduce the content of competitive bacteria and fungi in a sample. Commonly this treatment comprises 30-min exposure to 50 °C (Leoni & Legnani Citation2001; Roberts et al., Citation1987). However, increasing exposure time or temperature further may reduce Legionella cultivability, particularly at temperatures above 60 °C (Rogers et al., Citation1994).
More commonly, water samples are acid treated using an adaptation of methods developed by Bopp et al. (Citation1981). In this procedure water samples are either filter concentrated, or centrifuged and resuspended to produce a concentrate of the original sample. This concentrate is then diluted in a HCl–KCl buffer of pH 2.2 and incubated for ∼15 min before plating. To further reduce interfering microbial growth, plates may be incubated under a microaerophillic in a candle jar or under a 2.5% CO2 atmosphere at 35 °C (CDC, Citation2005).
For laboratory culture, Legionella spp. requires relatively complex culture media in order to multiply. This requires specific additions to standard nutrient media, including l-cycseine, arginine, isoleucine, leucine, threonine, valine, methionine, phenylalanine, tyrosine and serine (Pine et al., Citation1979). The addition of trace elements iron, calcium, cobalt, copper, magnesium, manganese, molybdenum, nickel, vanadium and zinc has also been shown to stimulate the growth of Legionella species in culture (Reeves et al., Citation1981; Warren & Miller, Citation1979).
Several agar formulations exist, with slight differences in selectivity and growth characteristics for different Legionella species. Buffered charcoal yeast extract agar (BCYE) is the most commonly used for general growth and maintenance of Legionella spp. and contains 0.1% α-ketoglutarate and a range of supportive amino acids and micro-nutrients (CDC Citation2005; Feeley et al.; Citation1979; Pendland et al., Citation1997; Roberts et al., Citation1987; Ta et al., Citation1995). A modification of this agar containing 1% albumin (ABCYE) has been shown to slightly enhance the recovery and growth of L. micdadei and L. bozemanii (Morrill et al., Citation1990).
A range of antimicrobial compounds may be added to the BCYE agar base, which are designed to reduce the growth of competing bacteria and fungi without altering the growth of Legionella spp. This range of compounds incorporates vancomycin, polymyxin B and cycloheximide and will often include glycine to reduce the growth of glycine-sensitive Gram negative bacteria (Wadowsky & Yee, Citation1981). Treated environmental samples may also be plated onto selective media deficient in l-cysteine to serve as a negative control for Legionella spp. growth.
The most significant barriers to quantitative, reproducible enumeration of Legionella spp. using culture arises from two distinct problems; the growth of unwanted microorganisms which obscure identification (Bopp et al., Citation1981) and the presence of viable but non-culturable (VBNC) Legionella spp. (Shih & Lin Citation2006). Legionella which have replicated intracellularly (within marcophagic hosts) are morphologically distinct from other Legionella cells (Al-Bana et al., Citation2013). These cells have thickened outer membranes, greater resistance to environmental and chemical stresses, lower metabolic rates and readily enter a VBNC state in water. Hence, the number of Legionella cells detected using culture immediately post-parasitization of amoebic hosts may be distinctly lower than Legionella in different life-cycle stages. Chang et al. (Citation2007) also demonstrated that L. pneumophila became VBNC after starvation in nutrient-free water for 33–40 days. They also demonstrated that heat disinfection at temperatures 60 °C or higher for between 5 and 30 min caused L. pneumophila to become completely unculturable but a large number of cells remained viable as determined by LIVE/DEAD BacLight bacterial viability kit (Molecular Probe, Eugene, OR). Also the longer a cell had undergone starvation the greater resistance it had to chlorine disinfection and heat treatment. A similar study by Alleron et al. (Citation2008) demonstrated that L. pneumophila treated with 1–10 mg/L of monochloramine became unculturable on BCYE; however, 28.8–29.4% of cells were viable as determined by the LIVE/DEAD BacLight kit. Legionella recovery from environmental sources has been shown to be enhanced by passage through amoebic hosts (La Scola et al., Citation2001; Rowbotham, Citation1983). It has also been demonstrated that non-cultureable L. pneumophila can be resuscitated by co-culture with Acanthamoeba polyphaga (García et al., Citation2007) and Acanthamoeba castellanii (Steinert et al., Citation1997). However, this adds further time to the isolation process and at best only allows for a qualitative presence/absence assessment of Legionella’s presence in a sample. Sample holding time also exerts a significant impact upon Legionella recovery by culture, with enumerated Legionella changing by up to 50% within 6 h and up to 2 log difference after 24 h (McCoy et al., Citation2012).
If culturable Legionella is present in its slow growth rate often leads to plate overgrowth by competing organisms with more rapid generation times (Alary & Joly, Citation1992; Bopp et al., Citation1981; Steele, Citation1990). Plates often require at least 5–7 days before Legionella colonies become visible, at which point the density of competing organisms often either renders enumeration infeasible or completely obscures the surface of culture plates (Bopp et al., Citation1981, Leoni & Legnani, Citation2001). Once Legionella colonies are visible, their positive identification is often primarily visual and/or confirmed by latex agglutination (Sathapatayavongs et al., Citation1983). However, these methods in their own right present limitations, as noted by Carvalho et al. (Citation2007) who demonstrated that from 20 colonies presenting the characteristic Legionella “ground glass” appearance on BYCE GVPC agar, all 20 were negative when tested using latex agglutination and fluorescent antibody assays but were confirmed to be Legionella when sequencing of 16S rDNA was carried out.
As Legionella is a pathogen of public health concern, the consequences of reporting false negatives or underreporting the concentration of Legionella may be serious. The potential harm caused by the failure to detect and treat systems containing Legionella may be ultimately deemed greater than the cost of presumptively treating/cleaning systems where results are equivocal. As culture results tend to underestimate the presence of Legionella in water systems, it may be better suited to use culture as an adjunct to molecular detection rather than an alternative.
qPCR enumeration
qPCR is an alternative method for rapid Legionella spp. enumeration from environmental samples (Joly et al., Citation2006). It simultaneously amplifies and quantifies a target DNA sequence (Templeton et al., Citation2003), giving the number of genome units (GU) per liter. An equivalence with the number of CFU has not been established and the results obtained are highly dependent upon the method used and the sample composition (Wellinghausen et al., Citation2001).
The rapid turn-around time and sensitivity of qPCR is advantageous when compared to traditional culture methods (Yaradou et al., Citation2007). The main limitation is a tendency to overestimate due to the amplification of non-viable or “dead” cells (Delgado-Viscogliosi et al., Citation2009). DNA within environmental samples can be very stable and may persist for extended lengths of time (Nocker et al., Citation2007). Josephson et al. (Citation1993) demonstrated that in biofilm DNA from non-viable cells persisted from days to weeks depending on the microbial consortium present. Some studies have shown that pre-treatments with ethidium monoazide (EMA) and propium monoazide (PMA) prior to DNA extractions enable amplification of viable cells only (Chang et al., Citation2009; Chen & Chang Citation2010; Delgado-Viscogliosi et al., Citation2009; Qin et al., Citation2012). When exposed to light, EMA and PMA bind to DNA that is not protected by a cell membrane and prevents its amplification, and hence enumeration by qPCR. These methods have not been optimized for differing sample types and their reliability and accuracy of these results is still debated (Hein et al., Citation2006). Pisz et al. (Citation2007) demonstrated that EMA was not effective in preventing the amplification of non-viable cells within biofilm samples, and suggested that the presence of extracellular polymeric substances could interfere with either the DNA binding or the photo-activation of EMA. Other studies have shown EMA to penetrate intact cells, with the extent of EMA-uptake-dependant on bacterial species and EMA concentration (Flekna et al., Citation2007; Kobayashi et al., Citation2009). Conversely, if the concentration of EMA or PMA is too low, insufficient free DNA will be bound resulting in further confounding unknowns (Fittipaldi et al., Citation2011).
Another difficultly with qPCR is the presence of environmental compounds inhibiting the qPCR reaction (Brooks et al., Citation2004). However, conducting 1:10 dilutions of DNA extracts has been shown to be effective at reducing inhibitors and enabling quantification of target DNA (Ballard et al., Citation2000). There are also a range of commercially available kits which contain components that may aid in the removal of qPCR inhibitors from DNA extracted from target samples (Wilson, Citation1997).
Comparison of current literature
Publications from the last 10 years which concurrently used culture and qPCR to detect Legionella spp. from environmental samples are collated in . Papers published in English and between the years 2003 and 2013 were included. Clinical and in situ experiments, including artificially spiked environmental samples were not included in this review.
Table 1. Comparison of published studies from 2003 to 2013 using qPCR and culture enumeration of Legionella spp. from environmental samples.
Some studies which compared PCR (not qPCR) and culture are included in if they provide a particular point of interest; however, for the basis of comparison the authors will only use the results obtained from the 28 studies which specifically enumerated Legionella using both culture and qPCR concurrently.
When the results of these studies are aggregated it becomes apparent that culture is more likely to underreport the presence of Legionella in water samples, with only 1/28 studies reporting higher detectable Legionella using culture and qPCR. In contrast, 25/28 studies reported higher detectable levels of Legionella using qPCR and one study reported equivalent results using both methods. On a sample per sample basis, samples analyzed concurrently by qPCR and culture were approximately 50% more likely to return a positive result by qPCR, with 2856/39 673 (72%) of all samples positive by qPCR and 1331/3967 (34%) of samples positive by culture.
The study by Levi et al. (Citation2003), which reported higher detectable Legionella using culture compared to qPCR could be explained by the high limit of detection (800CFU/L) of the qPCR method used in this study. Four of the studies noted the presence of qPCR inhibitors (Behets et al., Citation2007; Brooks et al., Citation2004; Parthuisot et al., Citation2010; Yaradou et al., Citation2007), which either could not be removed or required additional sample processing to allow for qPCR detection. Whilst one study noted complete disparity between results, reporting five samples as positive by culture, five samples positive by qPCR, but no samples returning a positive result by both culture and qPCR (Hsu et al., Citation2009).
Conclusion
Legionella spp. continues to exist as a public health concern; an ongoing risk assessment focus and an obstacle for cooling tower operators and facility managers. The discrepancies between testing procedures highlights the requirement for adopting a standard method for Legionella spp. detection in environmental samples. This review identifies the numerous inconsistencies between culture and qPCR enumeration, with studies from the last decade reporting a 50% difference between methods. International consensus is required to develop a universality accepted testing protocol to ensure consistency of results for both research purposes and risk assessment and management legislation.
Declarations of interest
The authors report no declarations of interest.
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