Quantitation of Pseudomonas aeruginosa in wound biopsy samples: from bacterial culture to rapid ‘real-time’ polymerase chain reaction

Early diagnosis of wound colonisation or prediction of wound sepsis provides an opportunity for therapeutic intervention. There is need for qualitative and quantitative tests that are more rapid than bacterial culture. Pseudomonas aeruginosa results in high morbidity and mortality rates, is inherently resistant to common antibiotics, and is increasingly being isolated as a nosocomial pathogen. We developed three PCR-based methods to detect and quantify P aeruginosa in wound biopsy samples: conventional PCR, enzyme-linked immunosorbent assay (ELISA)-PCR, and RTD-PCR with rapid thermal cycling (LightCycler^™ technology), all based on the amplification of the outer membrane lipoprotein gene oprL. We compared the efficacy of these methods to bacterial culture by quantitatively measuring levels of P aeruginosa in serial dilutions, in reconstituted skin samples and 21 burn wound biopsy samples.

The PCR mixture contained sterile distilled water, PCR buffer, deoxynucleotide mixture or digoxigenin labelling mix, MgCl₂, diluted template, primers PAL1 and PAL2, and AmpliTaQ DNA polymerase. The amplification was performed in a GeneAmp^® PCR System 2400. An aliquot of the reaction mixture was put on an agarose gel for electrophoresis and visualisation of the PCR product. An image of the gel was made using a digital camera. Image analysis software was used to calculate the band mass of the experimental bands.

An aliquot of the digoxigenin labelling reaction was denatured and then hybridized with the biotinylated capture probe PrL. Some of the resultant solution was transferred to a well of a streptavidin-coated microtitre plate (MTP) and incubated for 3 h at 45°C. The solution was discarded. Peroxidase conjugated antidigoxigenin was added and the MTP was incubated for 30 min at 37°C. The solution was discarded and ABTS substrate was added. The MTP was incubated for 30 min at 37°C. Absorbance was read at 405 nm.

The RTD-PCR mixture contained PCR grade sterile water, diluted template DNA, primers PAL1 and PAL2, 3' fluorescein (FL)-labelled probe oprL-FL, 5' LC Red 640-labelled and 3' phosphorylated probe oprL-LC, MgCl₂, and LC DNA Master Hybridisation Probes, containing Taq DNA polymerase, reaction buffer, dNTP mix with dUTP instead of dTTP, and MgCl₂. The amplification was performed in a LightCycler^™. The fluorescence signal of LC Red 640 was measured during the annealing phase. The measured fluorescence data was processed with analysis software.

The three methods showed a good concordance with the culture results. Conventional PCR was at least 100 times less sensitive than bacterial culture and had a low dynamic range (2 logs). With a lower detection limit of 10³ CFU/g tissue, ELISA-PCR was ten times more sensitive than conventional PCR. The dynamic range, however, did not increase. ELISA-PCR is very time consuming (8 h). The RTD-PCR produced a linear quantitative detection range of 7 logs with a lower detection limit of 10³ CFU/g tissue. More important, however, was that the time from sample collection to result was less than 1 h. Two biopsy specimens scored significantly higher in ELISA-PCR and RTD-PCR than in bacterial culture. This could indicate that DNA from dead bacteria was amplified. One out of ten culture positive biopsy samples was found negative by all PCR-based methods. Topical antimicrobial agents possibly inhibited PCR. These results show that RTD-PCR has potential for the rapid quantitative detection of pathogens in critical care patients, enabling early and individualized treatment. Further study is required to assess the reliability of this new technology, and its impact on patient outcome and hospital costs.

The PCR was completed in 200-μl microcentrifuge tubes. The PCR mixture (50 μl final volume) contained the following: 26.5 μl sterile distilled water; 5 μl 10×PCR buffer (500 mmol/l KCl and 100 mmol/l Tris-HCl: pH 8.3); 5 μl of a deoxynucleotide mixture (dGTP, dTTP, dATP and dCTP; 2 mmol/l each) or 5 μl digoxigenin-labelling mix from Boehringer-Mannheim (Brussels, Belgium; 2 mmol/l dGTP, dATP and dCTP, and 1.9 mmol/l digoxigenin-dUTP); 6 μl MgCl₂ (2.5 mmol/l); 5 μl diluted template DNA; 1 μl primer PAL1 (25 μmol/l); 1 μl primer PAL2 (25 μmol/l); and 0.5 μl AmpliTaQ DNA polymerase (5 U/μl). The primers had the following sequences: PAL1, 5'-ATGGAAATGCTGAAATTCGGC-3'; and PAL2, 5'-CTTCTTCAGCTCGACGCGACG-3'. All PCR reagents were purchased from PE Applied Biosystems (Nieuwerkerk a/d Ijssel, The Netherlands).

A reaction mixture containing all of the ingredients except the template was made. The amplification was performed in a GeneAmp^® PCR System 2400 (PE Applied Biosystems). The amplification program was set at 35 cycles of denaturation at 94°C for 30 s or 45 s (digoxigenin labelling), annealing at 57°C for 30 s or 1 min (digoxigenin labelling), and elongation at 72°C for 30 s or 2 min (digoxigenin labelling).

Some of the digoxigenin labelling reaction (35 μl) was transferred to a sterile microcentrifuge tube (Eppendorf). Denaturation solution (40 μl) was added and the tube was incubated for 10 min at room temperature. The tube was filled up to 500 μl with hybridization solution, which contained 50 pmol/ml biotinylated capture probe PrL with the following sequence: 5'-AAGCCGGAAGCCATGCGCG-CT-3'. Some of the resultant solution (200 μl) was transferred to a well of a streptavidin-coated microtitre plate (MTP). The MTP was incubated for 3 h at 45°C on a MTP shaker. The solution was discarded and the well was washed five times with 250 μl washing solution. A quantity (200 μl) of a peroxidase conjugated antidigoxigenin solution (10 mU/ml) was added, and the MTP was incubated for 30 min at 37°C on a MTP shaker. The solution was discarded and the well was washed five times with washing solution. ABTS substrate solution (200 μl) was added and the MTP was incubated for 30 min at 37°C on a MTP shaker. Absorbance was read at 405 nm. All digoxigenin detection reagents were purchased from Boehringer-Mannheim.

The RTD-PCR mixture (20 μl final volume) contained the following: 3.8 μl PCR grade sterile water; 3 μl diluted template DNA; 2 μl primer PAL1 (5 μmol/l); 2 μl primer PAL2(5 μmol/l); 2 μl of the 3' fluorescein-labelled probe oprL-FL (2 μmol/l); 4 μl of 5' LC Red 640-labelled and 3' phosphorylated probe oprL-LC (2 μmol/l); 1.2 μl MgCl₂ (25 mmol/l); 2 μl LC DNA Master Hybridisation Probes containing Taq DNA polymerase; reaction buffer; dNTP mix with dUTP instead of dTTP; and 10 mmol/l MgCl₂. All reagents were purchased from Roche Molecular Biochemicals (Brussels, Belgium). The probes were manufactured by the TIB MOLBIOL synthesis laboratory in Berlin, Germany, and had the following sequences: oprL-FL, 5'-TGCGATCACCACCTTCTACTTCGAGT-FL-3'; and oprL-LC, 5'-LC Red 640-CGACAGCTCCGACCTGAAG-p-3'.

Samples were spun into glass capillary tubes, capped and placed in the LightCycler^™. After an initial denaturation at 95°C for 30 s, amplification was performed for 45 cycles of denaturation at 95°C for 2 s, annealing at 59°C for 10 s and elongation at 72°C for 10 s. The fluorescence signal of LC Red 640 was measured during the annealing phase. The measured fluorescence data was processed with analysis software.

With a lower detection level between 10⁴ and 10⁵ CFU/g, wound biopsy tissue conventional PCR was at least 100 times less sensitive than bacterial culture (Table 1). This problem was related to the limited amount of sample, as compared with bacterial culture, that can be incorporated into one PCR reaction. Concentration of DNA or division of the sample over multiple PCR reactions could resolve this problem, but would result in a more elaborate method. Conventional PCR had a low dynamic range; bands that were very close to the background level gave poor results, and bands that were saturated also skewed the results (Fig. 1).

Ten out of twenty-one burn wound biopsy samples turned out to be culture positive; five of them, with a density lower than 10⁴ CFU/g, were not detected by conventional PCR. All culture-negative samples were also negative in PCR.

With a detection limit of 10³ CFU/g tissue, ELISA-PCR was at least 10 times more sensitive than conventional PCR (Table 1). However, the linear response range did not increase, but shifted to lower concentrations along with the detection limit (Fig. 1). This technique is very time-consuming and involves handling that increase the risk of contamination of PCR product (Fig. 2). Only one culture-positive wound biopsy, out of 10, containing 3000 CFU/g tissue, was found to be negative by ELISA-PCR. Topical antimicrobial agents possibly inhibited PCR.

Two biopsy specimens with approximately 200 and 400 CFU/g tissue, as determined by bacterial culture, scored significantly higher in ELISA-PCR: 1200 and 1450 CFU/g tissue, respectively. This could indicate that DNA from nonproliferating or dead bacteria was amplified. The question regarding whether quantitative estimates of bacteria, determined using PCR-based methods, correlate with the number of viable bacteria has been raised before [30]. For this reason, PCR-based tests are probably not appropriate for the monitoring of treatment efficacy. They should be considered as complementary indicative tests in critical care settings. On the other hand, PCR-based methods could detect bacteria that were inhibited in their growth by residual concentrations of antimicrobials in the sample [31], overgrown by other bacteria present in the sample, autolyzed during incubation [32], or difficult to cultivate in vitro [33], and are thus under-diagnosed by bacterial culture. PCR-ELISA has merit with regard to its sensitivity, but is too time-consuming and expensive to implement in a clinical laboratory (Fig. 2).

RTD-PCR produced a linear quantitative detection range of 7 logs with a lower detection limit of 10³ CFU/g tissue, or a few copies per reaction (Table 1). RTD-PCR uses a kinetic approach rather than an end-point approach. Kinetic quantification in the log phase of the reaction, the phase of constant efficiency, clearly generates a larger linear response than end-point detection in the plateau phase of the reaction (Fig. 1). Also, there is no need for any post-PCR sample manipulation, eliminating PCR contamination concerns. The recent introduction of a second dye for the LightCycler^™ system offers the prospect of an internal control, addressing the problem of tube-to-tube variations and PCR inhibition, and thus improving the reliability of the results. However, the greatest merit of Light-Cycler^™ RTD-PCR is its rapidity. Rapid thermal cycling reduced the time of amplification, detection and analysis of DNA from several hours to 30 min (Fig. 2). The RTD-PCR results for the burn wound biopsy samples were concordant with those obtained using ELISA-PCR; the false-negative result and the two over-diagnosed biopsies were also observed using RTD-PCR. The use of capillaries instead of tubes permits smaller reaction volumes, thus lowering the reagent costs. The set-up costs, however, may be beyond the capabilities of some laboratories (Table 1).