Human cytomegalovirus remains an important cause of morbidity and mortality among lung and heart transplant patients. Prophylactic treatment with ganciclovir (GCV) has been used justifiably to reduce the incidence of CMV disease. However GCV prophylaxis incurs expense, drug toxicity and the potential for the development of resistant strains. Alternatively preemptive GCV treatment can reduce CMV disease but this requires a sensitive diagnostic test that can be used as an accurate predictor of CMV disease (1).
The CMV antigenemia assay quantitates the level of replicating virus by assessing the expression of the pp65 antigen in PMNLs and has been used to effectively monitor CMV levels in a preemptive strategy to control CMV infection (2,3). This assay requires samples to be processed within a few hours, is labour intensive and requires skilled interpretation (2). The hybrid capture CMV DNA assay (4,5) offers the advantages of processing that can be delayed for up to 8 hr in a commercially available standardized quantitative assay, but lacks the sensitivity of DNA amplification methods.
Sensitive DNA amplification techniques offer the potential for detecting very low levels of CMV and have been described for use in predicting CMV disease (6,7). However, in a qualitative format the predictive value is limited (8,9,10) and quantitative viral load measurements are required for accurate prediction of disease. Although quantitative CMV PCR assays have been developed (11–16) many of these assays are technically complex and would be difficult to maintain in a routine diagnostic setting. Quantitative CMV PCR assays have been described to determine the relationship between viral load and probability of the progression of CMV disease, and these approaches have used a variety of clinical samples including urine, whole blood, plasma and polymorphonuclear leucocytes (9,15,17,18). Significant viral load levels varied dependent on the patient group studied as well as the sample type examined (19).
Competitive PCR assays involving the co-amplification of an exogenous target are the most successful strategy currently described and have been developed in a convenient microtitre plate format (15), although the linear dynamic range of these assays is often limited.
Automated PCR systems that detect the accumulation of amplified product in real-time have recently been described (20,21). The Perkin-Elmer Applied Biosystems (PE-ABI) 7700 Sequence Detection System employs a closed tube fluorescence-based format that eliminates postPCR processing and provides accurate real-time quantitative PCR. TaqMan™ assays have been described for the detection of several pathogenic organisms including hepatitis C (22), Salmonella spp (23)., Listeria monocytogenes (24), and Mycobacterium tuberculosis(25). The TaqMan™ assay utilises a dual-labeled fluorescent probe, the 5′ end of which is labeled with one of several reporter dyes such as FAM (6, carboxyfluorescein) and the 3′ end with the fluorescent dye TAMRA (6 carboxy-tetramethylrhodamine) (26). On the intact probe TAMRA dye quenches the signal from the reporter dye. During amplification, the probe hybridises to the target sequence bounded by the primers. The probe is subsequently digested by the 5′ exonuclease activity of Taq DNA polymerase during primer extension releasing the reporter dye and resulting in a relative increase in fluorescent signal.
We report here the development of a quantitative CMV assay using the PE-ABI TaqMan™system to measure the CMV viral load in whole blood samples. The assay was evaluated prospectively in a cohort of lung and heart transplant patients over a period of nine months. The aim of this study was to compare CMV viral load quantification by antigenemia with TaqMan™ CMV QPCR in this cohort of patients.
From March - December 1998 a prospective evaluation was carried out to monitor the CMV viral loads determined by TaqMan™ CMV PCR compared with CMV antigenemia. In total 25 patients were included in the study group, 12 were single-lung and 13 were heart transplant recipients. The CMV serological status of the donor and recipient (D+ or -, R+ or -) was determined prior to transplantation (CMV Scan, Becton Dickinson), and is summarized in Table 1. Surveillance samples were collected on the first postoperative day and then weekly for six weeks, two weekly until week twelve, and then monthly thereafter. More frequent samples were taken following an antigenemia or PCR positive sample or on suspicion of symptomatic CMV infection.
All patients had received intravenous methylprednisolone and azathioprine 4 mg/kg as induction immunosuppression. In addition all heart recipients, heart-lung recipients and single lung transplant recipients received rabbit anti-thymocyte globulin, 1 mg/kg for three days, for induction of immunosuppression. For maintenance immunosuppression all received cyclosporin, azathioprine 2 mg/kg, and prednisolone.
EDTA blood samples were collected for TaqMan™ CMV QPCR and CMVantigenemia. For PCR analysis DNA was extracted from 200 μl of EDTA blood using the Gentra blood extraction kit (Flowgen, Lichfield, UK) according to the manufacturer’s protocol and recovered in a 200 μl volume of elution buffer. For the antigenemia cytospin preparations of dextran enriched PMNLs were obtained, and incubated with the monoclonal antibody directed against the pp65 antigen (mouse monoclonal antibody IC3 anti-CMV pk 65, Biosoft, Bucks, UK) followed by immunofluorescence staining, and the number of antigen positive cells per 200 000 PMNLs counted using fluoresence microscopy (2).
The specificity of the TaqMan™ CMV QPCR assay was assessed by amplifying extracted DNA from virus culture isolates of herpes simplex 1 (n=6), herpes simplex 2 (n=6), varicella zoster (n=4), sera from patients positive for hepatitis C (n=2), HIV (n=1) and hepatitis B (n=3), human herpes virus 6 (n=1) and Epstein-Barr virus DNA (Sigma, Poole UK). DNA was extracted from virus isolates and sera using Dnazol (Life Technologies, Glasgow UK) according to the manufacturer’s instructions. Twenty CMV seronegative EDTA blood samples were also tested.
Amplification using the PE-ABI 7700 Sequence Detection System
The primers and probe for the TaqMan™ assay were designed from the glycoprotein B gene sequence using the Primer Express software program (PE-ABI) (Genbank Accession X04606) and were synthesised by Oswel DNA services (Southhampton UK). The forward and reverse primer sequences were as follows: 5′CTGCGTGATATGAACGTGAAGG-3′, 5′-ACTGCACGTACGAGCTGTTGG-3′. The TaqMan™ probe (5′-CGCCAGGACGCTGCTACTCACGA-3′) was labeled at the 5′ end with FAM and the 3′ end was labeled with the fluorescent quencher TAMRA and phosphorylated to prevent probe extension during amplification.
A volume of 2 μl of extracted samples was added to 23 μl of PCR reaction mix prepared from TaqMan™ Universal master mix, 200 nM of each of two primers and 100 nM of FAM labeled probe. DNA amplification was carried out on the PE-ABI 7700 Sequence Detection System by heating at 50°C for 5 min, 95°C for 10 min followed by 45 cycles of a two-stage temperature profile of 95°C for 15 seconds and 60°C for 1 min. Data points collected following primer extension were analyzed at the end of thermal cycling. A threshold value is determined as 10 standard deviations above the mean of the background fluorescence emission for all wells between cycle 3 and 15. The cycle number at which the fluorescence signal from a positive sample crosses this threshold is recorded. Quantitation standards were included in every assay. 10-fold dilutions of the plasmid standard containing a cloned insert of the target sequence were prepared in water ranging from 10 to 106 copies per PCR reaction and amplified in duplicate. The cycle number at which the standards became positive and crossed the calculated threshold was used to generate a standard curve from which the viral load from patient samples could be extrapolated (Fig. 1).
Generation of a Quantitative Standard
The PCR product generated from primers amplifying part of the glycoprotein B gene was cloned using the Invitrogen TOPO (R&D systems Oxford, U.K.) cloning system. From a selected plasmid containing the cloned glycoprotein B PCR product a purified plasmid preparation was obtained (Pureprep Kit, Pharmacia, Milton Keynes. U.K.). Optical density of the plasmid preparation was determined spectrophotometrically (GeneQuant, Pharmacia) and the DNA concentration calculated.
Regression analysis and Pearson correlation was used to compare antigenemia and TaqMan™ CMV PCR viral load values.
Sensitivity and Specificity
A TaqMan™ quantitative PCR assay targeting the glycoprotein B gene of CMV was developed. The quantitative standards enabled real-time quantitative measurements over a wide linear range from 10 to 106 genome copies per PCR reaction (Fig. 1). Ten copies of the cloned plasmid standard could be reproducibly detected. The theoretical limit of sensitivity was 500 copies per ml of blood based on a sampling volume of 2 μl of EDTA blood following extraction. Primers and probe were specific for the glycoprotein B target gene and negative results were obtained for the range of viruses described in the Methods. All twenty seronegative blood samples tested were also negative in the TaqMan™ assay.
Overall 362 samples were tested from 25 patients of which all 8 of the D+R+ patients and 4 of the 7 D+R- patients were positive for both antigenemia and TaqMan™ CMV QPCR viral load (Table 1). The remaining 10 D-R+ and D-R- patients were negative by both assays. Of the 362 samples 128 were positive and 188 were negative by both assays (Table 2). The sensitivity of the TaqMan™ assay compared to the antigenemia was 84.2% and the specificity was 89.5%. Of the antigenemia positive PCR negative samples 12 were positive with only one positive PMNL per 200 000 with the remainder with 6 or less positive cells. Similarly the 22 samples PCR positive and antigenemia negative had relatively low values ranging from log 1.6 to 3.9 genome copies per ml.
The reproducibility of the extraction procedure and the TaqMan™ amplification and detection was assessed using selected samples representative over the whole of the linear range of the assay. The standard deviation (SD) is shown for these samples (Fig. 2). Reproducibility between four separate extractions from the same sample showed the greatest variation at the lower concentrations with a SD of 0.24 log genome copies per ml blood for sample 4. For all samples with mean viral loads above log 2.82, SD values were all below log 0.1. Differences between successive patient samples of 0.5 log or greater were considered significant differences.
Correlation between Antigenaemia and TaqMan™ CMV QPCR
Comparison of antigenemia and TaqMan™ CMV QPCR viral load estimates for all samples tested (Fig. 3) demonstrated a statistically significant relationship (R=0.843, P =0.001). Analysis of the D+R- patients alone showed the correlation to be higher (R=0.8565, P =0.001), while the D+R+ patients showed a slightly lower correlation (R=0.6868, P =0.001) (data not shown).
The D+R+ patients had on average lower viral load measured by TaqMan™ CMV QPCR and antigenemia than the D+R- patients (Table 1). At lower levels the variability was greatest (Fig. 2.) which would adversely affect the correlation between the two assays.
Although the correlation between the two assays was highly significant TaqMan™ CMV QPCR viral load values show a spread of values over the range of antigenemia values. Serial monitoring of patients show comparisons of TaqMan™ CMV QPCR viral load and antigenemia values are closely paralleled (Fig. 4), although very often not exactly synchronised and therefore would contribute to the spread of viral load values compared to antigenemia.
Transplant patients monitored with the antigenemia test has established a clinical threshold of 50 pp65 positive cells per 200 000 PMNLs predictive of the development of CMV disease (2). Positive values only were compared for both the antigenemia and TaqMan™ CMV QPCR in Figure 5. Data points were simplified by plotting the mean values for selected ranges of antigenemia values from 1–10, 11–20, 21–40, 41–80, 130–500, and 600–2500 with the corresponding mean values of genome copies per ml determined by TaqMan™ CMV QPCR. The plot (Fig. 5) again shows the highly significant relationship between antigenemia and TaqMan™ CMV QPCR viral load estimates (R=0.9794, P =0.001) and from the equation defining the linear regression line the TaqMan™ CMV QPCR viral load equivalent of 50 PMNL positive cells per 200 000 is log 4.6 genome copies per ml of blood.
Correlation between CMV Disease and TaqMan™ CMV QPCR Viral load
Prophylaxis and preemptive GCV treatment based on antigenemia levels has been used to treat heart/lung transplant patients in this study, however despite this, symptoms attributable to CMV disease were recorded in 3 of the 25 patients. Of the D+R- group, Patient 11 recorded a pyrexia due to CMV infection, and Patient 10 developed extensive retinitis (Table 1). Patient 9 had no recorded symptoms due to CMV despite having very high viral load levels by antigenemia and TaqMan™ CMV QPCR. Of the 8 D+R+ patients a pyrexia was attributed to CMV in patient 1. The other 7 were asymptomatic with peak viral load values approaching the clinical threshold for the antigenemia and TaqMan™ CMV QPCR in patient 4 only. All of these patients received GCV treatment.
The effectiveness of GCV treatment can be effectively monitored using TaqMan™ CMV QPCR viral load and in Figure 4A and 4 B the patient’s response to GCV treatment is closely paralleled by antigenemia and QPCR viral load values.
This study reports the development of quantitative CMV PCR assay from extracted whole blood samples using the PE-ABI automated Sequence Detection System and TaqMan™ assay. The TaqMan™ real-time PCR enabled quantification of CMV viral load over a wide linear range.
Viral load levels estimated from extracted whole blood using the TaqMan™ CMV QPCR showed a highly significant linear correlation with antigenemia levels. Analysis of the relationship between CMV antigenemia levels and TaqMan™ CMV PCR in this study established an equivalent threshold value of log 4.6 genome copies per ml of blood. Disease due to CMV was recorded in three patients in this study group, two of which had viral loads above the predictive thresholds for both the antigenemia and TaqMan™ CMV QPCR assays. The third patient who had a pyrexia attributed to CMV had antigenemia and PCR viral load values below the threshold values. Conversely Patient 9 showed no signs of CMV disease despite very high levels of antigenemia and TaqMan™ CMV QPCR viral load. Intervention with GCV treatment in these patients would both influence the measured TaqMan™ CMV QPCR viral load and antigenemia levels and the development of CMV disease. Patients negative by antigenemia throughout the study were also TaqMan™ CMV QPCR negative indicating the negative predictive value is similar to that established for the antigenemia assay (2) which is important in preventing unnecessary expensive and toxic treatment.
The clinical threshold for the TaqMan™ CMV QPCR established by comparison with antigenemia levels for this group of transplant patients, may not be applicable in different transplant programmes such as bone marrow transplant recipients where much lower viraemia levels as determined by antigenemia assay have been shown to be associated with CMV disease (27). Similarly, viral load levels may not be comparable to estimates for extracted DNA from plasma or buffy coats (7). Other factors such as patient serostatus should be judged in context with the quantitated viral load. For example in this centre it is current practice to initiate GCV therapy with low level antigenemia levels in lung transplant patients who are at risk of primary infection. While absolute threshold values are often useful, the relative changes in viral load can be equally valuable in assessing the relative risk of a patient. The inherent variability of this PCR assay is therefore important in determining significant changes. Repeated extracts from selected samples with low viral loads showed the greatest variation with a standard deviation of log 0.24 genome copies per ml blood. Therefore samples showing variation greater than 0.5 log were considered to be significant changes in this population. Analysis of the kinetics of the TaqMan™ CMV QPCR from longitudinal monitoring of patients showed viral load levels closely paralleled antigenemia. This information can be used in context with the relative risk of infection as an indicator to treat and subsequently monitor the success of the treatment. TaqMan™ CMV QPCR may be used to effectively monitor response to and the length of treatment.
By comparison with antigenemia TaqMan™ CMV QPCR has important practical advantages. Delays in sample processing can result in significant decrease in antigenemia levels (28), while blood samples for PCR testing may be stored for up to 72 hr without deterioration (29,30). The automated sequence detection system employing the TaqMan™ assay provides a robust platform suitable for the demands of a routine diagnostic laboratory. Consumable costs for extraction and TaqMan™ amplification are estimated at £2.00 per reaction however the capital outlay of £70 000 for the Applied Biosystems 7700 system is considerable. The rapid extraction and automated amplification and detection allows same day testing necessary for early initiation of anti-viral therapy and the accurate quantification allows clinically useful assessment of the effect of antivirals or other intervention. The 96 well format also provides the potential for rapid and high throughput testing which would enable a centralised strategy for screening and viral load monitoring of CMV.
The TaqMan™ CMV QPCR therefore offers an alternative to antigenemia in directing preemptive treatment and monitoring CMV infection in lung and heart transplant patients.
1. Rubin RH. Preemptive therapy in immunocompromised hosts. N Engl J Med 1991; 324: 1057–1059.
2. Egan JJ, Barber L, Lomax J, et al. Detection of human cytomegalovirus antigenemia: a rapid diagnostic technique for predicting cytomegalovirus infection/pneumonitis in lung and heart transplant recipients. Thorax 1995; 50: 9–13.
3. Gerna G, Zipeto D, Parea M, et al. Monitoring of human cytomegalovirus infections and ganciclovir treatment in heart transplant recipients by determination of viremia, antigenemia, and DNAemia. J Infect Dis 1991; 164: 488–498.
4. Hebart H, Gamer D, Loeffler J. Evaluation of Murex CMV DNA hybrid capture assay for detection and quantitation of cytomegalovirus infection in patients following allogeneic stem cell transplantation. J Clin Microbiol 1998; 36: 1333–1337.
5. Barrett-Muir WY, Aitken C, Templeton K, Raftery M, Kelsey SM, Breuer J. Evaluation of the murex hybrid capture cytomegalovirus DNA assay versus plasma PCR and shell vial assay for diagnosis of human cytomegalovirus viremia in immunocompromised patients. J Clin Microbiol 1998; 36: 2554–2556.
6. Aitken C, Barrett-Muir W, Millar C, et al. Use of molecular assays in diagnosis and monitoring of cytomegalovirus disease following renal transplantation. J Clin Microbiol 1999; 37: 2804–2807.
7. Tong CY, Cuevas L, Williams H, Bakran A. Use of laboratory assays to predict cytomegalovirus disease in renal transplant recipients. J Clin Microbiol 1998; 36: 2681–2685.
8. Boeckh M, Gallez-Hawkins GM, Myerson D, Zaia JA, Bowden RA. Plasma polymerase chain reaction for cytomegalovirus DNA after allogeneic marrow transplantation: comparison with polymerase chain reaction using peripheral blood leukocytes, pp65 antigenemia, and viral culture. Transplantation 1997; 64: 108–113.
9. Niubo J, Perez JL, Manito N, Garcia A, Roca J, Martin R. The polymerase chain reaction as a marker of cytomegalovirus infection in heart transplant recipients [in Spanish]. Med Clin (Barc) 1999; 112: 121–124.
10. Barber L, Egan JJ, Lomax J, et al. Comparative study of three PCR assays with antigenemia and serology for the diagnosis of HCMV infection in thoracic transplant recipients. J Med Virol 1996; 49: 137–144.
11. Fox JC, Griffiths PD, Emery VC. Quantification of human cytomegalovirus DNA using the polymerase chain reaction. J Gen Virol 1992; 73: 2405–2408.
12. Zipeto D, Baldanti F, Zella D, et al. Quantification of human cytomegalovirus DNA in peripheral blood polymorphonuclear leukocytes of immunocompromised patients by the polymerase chain reaction. J Virol Methods 1993; 44: 45–55.
13. Imbert-Marcille BM, Cantarovich D, Ferre-Aubineau V, Richet B, Soulillou JP, Billaudel S. Usefulness of DNA viral load quantification for cytomegalovirus disease monitoring in renal and pancreas/renal transplant recipients. Transplantation 1997; 63: 1476–1481.
14. Roberts TC, Brennan DC, Buller RS, et al. Quantitative polymerase chain reaction to predict occurrence of symptomatic cytomegalovirus infection and assess response to ganciclovir therapy in renal transplant recipients. J Infect Dis 1998; 178: 626–635.
15. Barber L, Egan JJ, Turner AJ, et al. The development of a quantitative PCR ELISA to determine HCMV DNAaemia levels in heart, heart/lung and lung transplant recipients. J Virol Methods 1999; 82: 85–97.
16. Ferreira-Gonzalez A, Fisher RA, Weymouth LA, et al. Clinical utility of a quantitative polymerase chain reaction for diagnosis of cytomegalovirus disease in solid organ transplant patients. Transplantation 1999; 68: 991–996.
17. Cope AV, Sweny P, Sabin C, Rees L, Griffiths PD, Emery VC. Quantity of cytomegalovirus viruria is a major risk factor for cytomegalovirus disease after renal transplantation. J Med Virol 1997; 52: 200–205.
18. Gor D, Sabin C, Prentice HG, et al. Longitudinal fluctuations in cytomegalovirus load in bone marrow transplant patients: relationship between peak virus load, donor/recipient serostatus, acute GVHD and CMV disease. Bone Marrow Transplant 1998; 21: 597–605.
19. Boeckh M, Boivin G. Quantitation of cytomegalovirus: methodologic aspects and clinical applications. Clin Microbiol Rev 1998; 11: 533–554.
20. Gibson UE, Heid CA, Williams PM. A novel method for real time quantitative RT-PCR. Genome Res 1996; 6: 995–1001.
21. Heid CA, Stevens J, Livak KJ, Williams PM. Real time quantitative PCR. Genome Res 1996; 6: 986–994.
22. Martell M, Gomez J, Esteban JI, et al. High-throughput real-time reverse transcription-PCR quantitation of hepatitis C virus RNA. J Clin Microbiol 1999; 37: 327–332.
23. Chen S, Yee A, Griffiths M, et al. The evaluation of a fluorogenic polymerase chain reaction assay for the detection of Salmonella
species in food commodities. Int J Food Microbiol 1997; 35: 239–250.
24. Bassler HA, Flood SJ, Livak KJ, Marmaro J, Knorr R, Batt CA. Use of a fluorogenic probe in a PCR-based assay for the detection of Listeria
monocytogenes. Appl Environ Microbiol 1995; 61: 3724–3728.
25. Desjardin LE, Chen Y, Perkins MD, Teixeira L, Cave MD, Eisenach KD. Comparison of the ABI 7700 system (TaqMan) and competitive PCR for quantification of IS6110
DNA in sputum during treatment of tuberculosis. J Clin Microbiol 1998; 36: 1964–1968.
26. Livak KJ, Flood SJ, Marmaro J, Giusti W, Deetz K. Oligonucleotides with fluorescent dyes at opposite ends provide a quenched probe system useful for detecting PCR product and nucleic acid hybridization. PCR Methods Appl 1995; 4: 357–362.
27. Gozlan J, Laporte JP, Lesage S, et al. Monitoring cytomegalovirus infection and disease in bone marrow recipients by reverse transcription-PCR and comparison with PCR and blood and urine cultures. J Clin Microbiol 1996; 34: 2085–2088.
28. Boeckh M, Woogerd PM, Stevens-Ayers T, Ray CG, Bowden RA. Factors influencing detection of quantitative cytomegalovirus antigenemia. J Clin Microbiol 1994; 32: 832–834.
29. Roberts TC, Buller RS, Gaudreault-Keener M, et al. Effects of storage temperature and time on qualitative and quantitative detection of cytomegalovirus in blood specimens by shell vial culture and PCR. J Clin Microbiol 1997; 35: 2224–2228.
30. Schäfer P, Tenschert W, Gutensohn K, Laufs R. Minimal effect of delayed sample processing on results of quantitative PCR for cytomegalovirus DNA in leukocytes compared to results of an antigenemia assay. J Clin Microbiol 1997; 35: 741–744.