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Initial evaluations of the Cobas AmpliPrep/Cobas TaqMan human immunodeficiency virus type 1 (HIV-1) test (CAP/CTM) demonstrated good performance but, afterwards, reports about underquantification were published. We investigated whether the problem was solved with a second version of this assay, the Cobas AmpliPrep/Cobas TaqMan HIV-1 test, version 2.0 (CAP/CTM v2.0). The remaining plasma of 375 consecutive HIV-1 positive samples with a viral load of ≥4,000 copies/ml was collected in three laboratories. The samples were diluted and retested with our routine method Cobas AmpliPrep/Cobas Amplicor HIV-1 monitor test v1.5 in ultrasensitive mode (CAP/CA PHS), as well as with the CAP/CTM and CAP/CTM v2.0 tests. An absolute difference between the results of two methods of ≥0.71 log10 copies/ml was defined as moderately discrepant, and an absolute difference of ≥0.93 log10 copies/ml was defined as severely discrepant. In addition, criteria for considering the new methods equivalent to the routine method were formulated. (i) For CAP/CTM compared to CAP/CA PHS, 36 (9.5%) and 20 (5.3%) samples were, respectively, considered moderately and severely underquantified by CAP/CTM. The mean difference between CAP/CTM and CAP/CA PHS was −0.32 log10 copies/ml. Eight of nineteen of the severely underquantified samples were from patients infected with HIV-1 subtype B strain. (ii) For CAP/CTM v2.0 compared to CAP/CA PHS, no sample was moderately or severely underquantified by CAP/CTM v2.0. A mean difference of 0.08 log10 copies/ml was found with CAP/CTM v2.0 compared to CAP/CA PHS. The underquantification problem of the CAP/CTM kit was clearly demonstrated. The criteria for the equivalence of CAP/CTM v2.0 to the routine test CAP/CA PHS were fulfilled.
Since the mid-1990, the human immunodeficiency virus type 1 (HIV-1) viral load (VL) assay is a major tool in the follow-up of HIV-1-infected individuals to predict progression of HIV disease and to monitor antiviral treatment response (5, 7, 14). Several patented methodologies using a variety of techniques, from reverse transcriptase PCR to the branched DNA assay, are commercially available for quantitative HIV-1 RNA testing in diagnostic laboratories.
VL assays were developed in industrialized countries, where HIV-1 subtype B predominates. In contrast, subtype B is a minor variant in developing countries and dissemination of non-B subtypes has also started in Western countries, especially in Belgium, where many viruses of African origin are circulating (17). Among HIV-1-infected patients attending Universitair Ziekenhuis Brussel (UZB), as many as 59% of the strains belong to non-B subtypes (6). This high genetic diversity of HIV-1 is a challenge for the quantification of plasma HIV-1 RNA. Indeed, in the past, several studies have reported the failure of commercial assays for viral load monitoring in patients infected with non-B subtypes (1, 2, 4, 19). This finding led Roche Diagnostics, Ltd. (Rotkreuz, Switzerland), the manufacturer of the widely used Cobas Amplicor HIV-1 monitor test version 1.0 assay, to modify this assay in 1997. By the addition of new primers, the assay could cover a broader range of viral diversity (Cobas Amplicor HIV-1 monitor test version 1.5 PHS/PHM).
In the early years of 2000, automation in sample extraction (for example, the Cobas AmpliPrep of Roche Diagnostics and M1000 of Abbott Diagnostics, Abbott Park, IL) facilitated high-volume testing and made viral load testing more robust.
With the ongoing introduction of new technologies, classical endpoint amplification techniques became more and more replaced by kinetic real-time fluorescence based tests that combine amplification and detection in one step. A TaqMan-based real-time technique (10) is simple, rapid, sensitive, specific, and reproducible and makes further automation possible. Moreover, the risk of contamination is lower due to the closed tube configuration. The Roche Cobas AmpliPrep/Cobas TaqMan (CAP/CTM) quantitative HIV-1 assay requires little manual intervention between the initial addition of the sample to the assay tube and the generation of the quantitative result. Initial evaluations of this assay were good (11, 15, 16, 18), but with a trend to lower viral load values on average than those obtained with the Cobas Amplicor HIV-1 monitor version 1.5 PHS/PHM assay (9). Afterward, reports about serious underquantification became available (3, 9, 20). The incidence of underquantification was low and seemed to be prominent only in selected populations.
To overcome the issue of underquantification, the CAP/CTM assay was modified to Cobas AmpliPrep/Cobas TaqMan HIV-1 test, version 2.0 (CAP/CTM v2.0). A dual-target strategy was chosen: besides gag primers and a FAM-labeled gag probe, additional ltr primers and a FAM-labeled ltr probe were included in the assay. The two targets, gag and ltr, are amplified with the same efficiency. The worst-case scenario is the complete failure of one of the PCR amplicon targets to be amplified. This would result in the loss of one-half of the fluorescent signal. The PCR of that specimen would then need one additional thermal cycle to achieve the threshold cycle (CT) for quantification. Hence, with a CT + 1, the specimen HIV-1 RNA titer would be quantified by half, equivalent to 0.3-log difference, from the “true” value. Guidelines state that a difference of <0.5 log copies/ml (cp/ml) is clinically not significant (5), and this variation has no clinical consequence.
The aim of the present study was to evaluate if the underquantification issue of the CAP/CTM HIV-1 test is solved with the introduction of the CAP/CTM v2.0 test. A comparison was performed with the routine Cobas Amplicor HIV-1 monitor version 1.5, ultrasensitive mode (CAP/CA PHS), the first version of CAP/CTM and CAP/CTM v2.0.
(Part of this research was presented at the 7th European HIV Drug Resistance Workshop, Stockholm, Sweden, 25 to 27 March 2009, poster 86.)
From May to September 2008, 375 consecutive EDTA anticoagulated plasma samples (collected in Sarstedt EDTA “lavender” tubes) stored at below −70°C after acquisition with HIV-1 viral load ≥4,000 cp/ml in the CAP/CA PHS (or with manual RNA extraction at Université Libre de Bruxelles [ULB]) routine test were selected. Apart from this condition and the availability of a sufficient sample volume, no selection for the inclusion of samples was done. At UZB, 5 of 83 (6.0%) samples were not included due to insufficient sample volume. Only one sample per patient was included.
After routine testing of the original specimen the remaining EDTA anticoagulated plasma was diluted 1:5 or 1:10 in HIV-1 RNA negative EDTA plasma pool (Roche Diagnostics GmbH, Germany) and divided into aliquots. Aliquots were stored frozen below −70°C until tested. All aliquots were subjected to equal numbers of freeze-thaw cycles.
Samples were selected and divided into aliquots at 3 laboratories in Brussels: the AIDS Reference Laboratory of the VUB, subunit UZ Brussel (UZB; n = 78) and subunit University Medical Center Saint-Pierre (STP, n = 149) and the AIDS Reference laboratory of the ULB, Erasme University Hospital (EHB, n = 148).
All samples were analyzed in the AIDS reference laboratory of the VUB. The subunit UZ Brussel analyzed the samples collected at EHB and UZB, while the subunit STP tested their own samples.
Each specimen was tested in singlet in the three study tests. The assays were performed in accordance to the package inserts of the manufacturer. For quantification of the target nucleic acid and control of PCR inhibition an internal quantification standard was used. The results were expressed in copy number of HIV-1 RNA copies per milliliter. The prevention of back-contamination was ensured by the use of uracil-N-glycosylase, an enzyme that inactivates dUTP-containing amplicons.
After probe-specific extraction with the ultrasensitive protocol on the Cobas AmpliPrep (CAP), amplification was done with the Cobas Amplicor HIV-1 Monitor Test, v1.5, and endpoint detection was performed using the Cobas Amplicor (CA) instrument. The primers target the highly conserved gag region. If the result was described above the upper quantification limit (>100,000 cp/ml), samples were diluted 1:10 or 1:100 with HIV-1 RNA negative human EDTA plasma and retested to obtain a measurable concentration of HIV-1 RNA. Several lots of reagents were used to perform the assays.
After generic extraction with the CAP, amplification was performed using the Cobas TaqMan HIV-1 Test on the Cobas TaqMan 48 (CTM) instrument. A dually labeled hybridization probe targeting the gag region was used in the assay. Multiple kit numbers were used to perform the assays.
This assay simultaneously targets the gag and the ltr region with two dually labeled hybridization probes. For the present study, a prerelease version, CAP/CTM version 1.5 provided by Roche Diagnostics was used. Afterward, based on extended validation, a software modification was performed to extend the lower quantification limit from 40 to 20 cp/ml. Therefore, this test version was later renamed as version 2.0, but the reagents of both assay versions are the same. This change did not influence our results since no samples with low viral load were included in this evaluation. CAP/CTM v2.0 is not yet available in the United States.
After generic extraction with the CAP, amplification was performed using the Cobas TaqMan HIV-1 Test, v2.0 on the CTM instrument. All assays were performed using one lot of reagents.
Each run consisted of 20 samples and one high positive, one low positive and one negative control included in the kit. In addition, in order to have an independent control, one in-house produced internal run control was analyzed in each run.
The minimal change in viral load considered as clinically significant is a 0.5 log10 cp/ml (5). Thus, the maximum total error (TEMAX), defined as systematic error (SE) + 1.96 × the random error (RE), must be lower than 0.5 log10 cp/ml. Because in Belgium HIV patients are monitored at one AIDS reference center and we know from our validation file that the bias is very small, we can assume that the SE is negligible. Thus, the maximum allowable REMAX is 0.26 log10 cp/ml. Because we allow a TE of 0.5 log10 cp/ml for both methods in a comparison the total TEMAX is 1.96(0.262 + 0.262) or 0.71 log10 cp/ml. Thus, the absolute difference between CAP/CA PHS and CAP/CTM or CAP/CTM v2.0 should be <0.71 log10 cp/ml for at least 95% of all samples. Samples that deviate by more than 0.71 log10 cp/ml are referred to as moderately discrepant.
According to the analogous principles described above we can state that for at least 99% of all samples the absolute difference between CAP/CA PHS and CAP/CTM or CAP/CTM v2.0 should be less than 2.58(0.262 + 0.262) or 0.93 log10 cp/ml. Samples that deviate by more than 0.93 log10 cp/ml are referred to as severely discrepant.
Absolute bias plots were used to represent the degree of agreement between the assays. The x axis of the absolute bias plot shows the mean of the results, and the y axis represents the absolute difference between the values obtained by the two platforms. The dotted lines represent the above-defined criteria for considering the new assay equivalent to the routine assay.
To obtain a normal distribution of the values, results were transferred to log10 values. Normality was checked by using a modified Shapiro-Wilk test. Mean values were compared to the t test for paired data. Epidemiological data were compared to the chi-square test. Statistics were performed with Analyze-It for Microsoft Excel (version 2.12; Analyze-It Software, Ltd., United Kingdom).
Samples with severe discrepancies (n = 20) were sequenced by Roche Molecular Diagnostics (Pleasanton, CA) with unpublished primers to identify possible mismatches in all primer and probe regions.
Nucleic acids were extracted from 200 μl of the original plasma (n = 12) or, when the original plasma was no longer available, from 200 μl of the 1:5-diluted (n = 8) plasma using sample preparation reagents from the Cobas HIV-1 Monitor test. A 600-μl portion of lysis reagent was mixed with a 200-μl sample, followed by incubation for 10 min at room temperature. Nucleic acids were precipitated by the addition of 800 μl of isopropanol and then pelleted by centrifugation for 15 min in a microcentrifuge. The pellets were washed with 1 ml of 70% ethanol (EtOH) and centrifuged for 5 min. The alcohol was removed, the tubes were pulse spun for 30 s, and the residual EtOH was removed. The pellets were resuspended in 100 μl of sample diluent from the Cobas HIV-1 Monitor test and stored at −80°C until used.
HIV-1 sequences flanking the CAP/CTM HIV-1 version 2 target regions were amplified by reverse transcription-PCR. Murine leukemia virus reverse transcriptase (Applied Biosystems, Foster City, CA) was used for reverse transcription, and PCR was carried out with AmpliTaq DNA polymerase (Applied Biosystems). Heminested amplifications were then performed to increase the yield of amplicon. Amplification products were analyzed by agarose gel electrophoresis. The amplicons from reactions with one predominant product band were purified for sequencing by using either QiaQuick PCR purification (Qiagen, North Rhine-Westphalia, Germany) or ExoSAP-IT kit reagents (USB, Cleveland, OH). If multiple major amplicon populations were present, the products of appropriate size, separated by agarose gel electrophoresis, were excised from the gel and purified using reagents from a QiaQuick gel extraction kit (Qiagen). Sequencing was either performed on the ABI 3730xl analyzer using BigDye chemistry and Collection Software v3.0 (Applied Biosystems) or by Sequetech Corp. (Mountain View, CA).
To determine the subtype of the virus in each sample, the gag sequence from each sample was aligned to sequences from HIV-1 isolates of known genotypes from the GenBank sequence database (http://www.ncbi.nlm.nih.gov/GenBank/index.html) using the CLUSTAL V sequence alignment program, which also generates a phylogenetic tree, via the Lasergene software package (MegAlign version 5.07; DNASTAR, Inc., Madison, WI).
For each new HIV diagnosis in Belgium the attending physician is asked to fill in a questionnaire and send it to the AIDS Reference Laboratory that made the diagnosis. All coded data are processed once a year by the Institute of Public Health to map and characterize the Belgian HIV epidemic. These data, although not complete, were used to compare the populations consulting the three different centers and to track the probable origin of the HIV-1 strains yielding severely discrepant results.
The present study was approved by the biomedical ethical commission of the Universitair Ziekenhuis Brussel and the Medical Faculty of the Vrije Universiteit Brussel (reference number BUN B14320084122).
Because the initial result was outside the dynamic range of the CAP/CA PHS assay, 22 samples had to be diluted 1:10 for the CAP/CA PHS analysis and 1 sample was diluted 1:100 for additional analysis. No dilutions had to be made for CAP/CTM and CAP/CTM v2.0 due to the broader dynamic range of these assays.
One sample (EHB_141) demonstrated an extremely high quantitative result with both CAP/CTM tests (+1.29 and +1.84 log10 cp/ml) compared to CAP/CA PHS. This result was considered as an outlier and was not taken into account in our calculations. The sequence of this sample contains six mismatches to the upstream CA PHS primer. The subtype of this sample was determined as CRF18_cpx.
For one sample (EHB_084) the result for CAP/CTM was <40 cp/ml, although HIV-1 RNA was detected with CAP/CA PHS (3.70 log10 cp/ml) and CAP/CTM v2.0 (3.75 log10 cp/ml). For the calculations we made an approximation and used 40 cp/ml, the best-case scenario for EHB_084.
The results are represented in an absolute bias plot (Fig. (Fig.11).
The overall mean difference between CAP/CTM and CAP/CA PHS was −0.32 log10 cp/ml (P < 0.05). The median difference was −0.29 log10 cp/ml with a range between −2.64 and 0.78 log10 cp/ml.
A total of 36 samples (9.6%; EHB [n = 16], STP [n = 15], and UZB [n = 5]) were moderately and 20 samples (5.3%; EHB [n = 10], STP [n = 7], and UZB [n = 3]) were severely underquantified with CAP/CTM compared to CAP/CA PHS. The observed differences between the different sites are not statistically significant (P = 0.55 and 0.59, respectively).
The sequence results and the probable origin of the severely underquantified strains are represented in Table Table1.1. For one sample (EHB_039) the amplification of the gag region repeatedly failed. The other 19 samples were from patients infected with 9 different subtypes. Eight of these nineteen (42.1%) patients were infected with a subtype B strain. In 18 of the severely underquantified samples, there are mismatches to CAP/CTM gag primers or probe that may affect assay performance. No mismatches were found in one sample (UZB_001), and thus the observed difference of −1.06 log10 cp/ml is probably due to an anomalous error.
The amplification of the ltr region that was done for further sequence comparisons succeeded in 16 of these 20 samples. There were no mismatches found to the CAP/CTM ltr region that would be likely to affect CAP/CTM v2.0 assay performance. Two (0.5%; EHB [n = 1], STP [n = 1], and UZB [n = 0]) samples were moderately higher quantified with CAP/CTM. These two samples were also higher quantified with CAP/CTM v2.0. As for sample EHB_141, the sample defined as an outlier, two (EHB_090) or three (SPB_083) mismatches to the upstream CA PHS primer were detected.
The results are represented in an absolute bias plot (Fig. (Fig.2).2). The titer values obtained with CAP/CTM v2.0 were on average +0.08 log10 cp/ml higher compared to CAP/CA PHS (P < 0.05). The median difference was +0.05 log10 cp/ml, with a range from −0.68 to 1.02 log10 cp/ml. No sample was moderately or severely underquantified. Seven (1.9%; EHB [n = 3], STP [n = 3], and UZB [n = 1]) samples were moderately and three (0.8%; EHB [n = 1], STP [n = 1], and UZB [n = 1]) were severely higher quantified by CAP/CTM v2.0. Five out of seven moderately higher quantified samples were sequenced. Three of them had five mismatches to the CA PHS upstream primer. For two of them no mismatches were found.
Three parameters (nationality, risk category, and probable country of infection) were compared by using a chi-square test. A statistically significant difference (P < 0.05) was observed for all three parameters.
The patient population consulting at UZB contains a higher fraction of people with the Belgian nationality (44.9% versus 24.2% at STP and 25.2% at EHB). The associated risk category is more often men having sex with men (MSM; 44.9% versus 22.8% at STP and 20.9% at EHB), and the probable country of infection is more often Belgium (39.4% versus 16.8% at STP and 10.8% at EHB).
The patients seeking medical advice at STP are more often European (not Belgian), North African, American, and Asian (11.4% versus 5.1% at UZB and 4.0% at EHB). Moreover, the risk categories mother-to-child transmission (MTCT) and intravenous drug use (IVD) are more prevalent at the STP patient population (7.4% versus 0.0% at UZB and 2.0% at EHB).
Both American (5) and European (7) guidelines for the treatment of HIV-1-infected adults make use of two markers—the viral load and the CD4+ T-cell count—to assess the level of HIV-1 viremia and the immune function of infected patients. These tests are used as predictors for deciding when to begin antiviral therapy and to assess virologic and immunologic efficacy of treatment. Therefore, accurate measurement of HIV-1 viral load is essential to provide clinicians with valuable information to determine treatment decisions. Although the new quantitative HIV-1 assays are designed to cope with increasing molecular diversity of the virus, there were several reports about serious underquantification issues with the first version of the CAP/CTM test (3, 9, 20). To overcome this problem, additional primers and a probe, located in the highly conserved ltr region of HIV-1, were included in the second version of the kit in addition to the gag primers and probe. We report here the results of a three-site multicenter evaluation study of the CAP/CTM and CAP/CTM v2.0 tests in comparison to the routine CAP/CA PHS test.
In Belgium, 58% of newly diagnosed HIV patients were of foreign origin in 2007 (17). Among HIV-1-infected patients attending UZB, subtype B virus is only present in 41% of the patients (6). Therefore, the sample collection used was expected to contain a wide variety of subtypes, including a lot of non-B subtypes, and consequently could represent a great challenge for the new generation of viral load tests.
The patient population attending the three centers is significantly different. The most prominent differences were that the patient population consulting at UZB contains a higher fraction of MSM with the Belgian nationality and the probable country of infection is more often Belgium. The patients seeking medical care at STP are more often from European (non-Belgian), North African, American, and Asian nationalities. Moreover, the risk categories MTCT and IVD are more prevalent at the STP patient population.
The results of the comparison of CAP/CTM and CAP/CA PHS are represented in an absolute bias plot (Fig. (Fig.1).1). A total of 36 of 375 (9.6%) samples were moderately and 20 of 375 (5.3%) samples were severely underquantified. Although not statistically significant (P = 0.59), the prevalence of severe underquantification was highest at EHB: 6.7% of samples compared to 4.6% at STP and 3.8% at UZB. No particular subtype is affected by the underquantification problem. The 19 subtyped, severely discrepant samples were from patients infected with nine different subtypes. Eight of these nineteen (42.1%) patients were infected with a subtype B strain. Thus, in contrast to earlier-generation HIV-1 VL assay problems (1, 2, 4, 19), even subtype B strains could be affected by the underquantification issue of the first version of the CAP/CTM test, indicating that the mismatches were not associated with the HIV-1 subtype.
As illustrated in Table Table1,1, both the primer and the probe binding region polymorphisms were identified as the root cause in the rare cases of significant underquantification. Due to patented primers and probes, Roche did not wish to disclose the exact location of the mismatches. Therefore, we cannot support or refute the theory of Korn et al. (12) that mutations at nucleotide position 1488 of the HXB2 reference sequence, the −3 position of the suggested downstream primer, are an important cause of the underestimation of HIV-1 RNA levels. Nevertheless, there should be other critical mutations because 8 of the 19 sequenced underquantified samples had no mismatches to the downstream primer.
We found that 2 of 375 (0.5%) samples were moderately higher quantified by CAP/CTM. These two samples are also moderately higher quantified (+0.79 and +0.91 log10 cp/ml, respectively) with CAP/CTM v2.0. Although not well documented since we only had access to the number of mismatches and not to the sequences as such, this finding is probably due to a better primer-probe match than with the CAP/CA PHS assay, the method used as the “reference” or comparison method. This is also the case with sample EHB_141, which we considered an outlier.
The overall mean difference between CAP/CTM and CAP/CA PHS was −0.32 log10 cp/ml (P < 0.05). This difference is statistically significant. Although this difference is <0.5 log10 cp/ml, which is generally accepted as clinically relevant (5), it clearly illustrates the underquantification issue of the first version of CAP/CTM. Moreover, the high number of moderately and severely underquantified samples is clinically problematic. The criteria used above to consider the new method equivalent to the routine method are not fulfilled for CAP/CTM.
The results of the comparison of CAP/CTM v2.0 and CAP/CA PHS are represented in an absolute bias plot (Fig. (Fig.2).2). No sample was moderately or severely underquantified. Eight (2.1%) samples were moderately and three (0.8%) of 375 samples were severely higher quantified with CAP/CTM v2.0. In all, four samples (three + EHB_141) had several mismatches to the upstream CA PHS primer and were probably underquantified with the older CAP/CA method. Thus, it appears that the high genetic diversity of HIV-1 also affects the performance of the “reference” or comparison method.
A mean difference of +0.08 log10 cp/ml was found with the CAP/CTM v2.0 compared to CAP/CA PHS (P < 0.05). This difference is statistically significant but not clinically relevant (<0.5 log10 cp/ml). The above-described criteria to consider the new method equivalent to the routine method are fulfilled for CAP/CTM v2.0.
In addition, one sample (EHB_141) quantified extremely high with both Cobas TaqMan tests (+1.29 and +1.84 log10 cp/ml; CAP/CTM and CAP/CTM v2.0, respectively) compared to the routine CAP/CA PHS test and was considered to be an outlier. The patient's CD4 count and clinical presentation correlated better with the higher VL (this patient stopped taking all antiretroviral therapy at sampling time). The sequence of the HIV-1 strain from this sample revealed six mismatches to the upstream CA PHS primer. Therefore, we consider that it was underquantified by the CAP/CA PHS test.
The present study has three limitations. (i) Lot-to-lot variability for the CAP/CTM v2.0 kit was not evaluated during our study. Only one lot of reagents was used during the evaluation. (ii) A bias was possibly generated by selecting only samples with a VL of ≥4,000 copies/ml, which were diluted in order to obtain enough sample volume to run the three tests in parallel. Indeed, most of the included samples were from untreated patients, harboring viral strains without mutation pressure from antiretroviral drugs. However, gag and ltr are not current drug targets and, although gag cleavage site mutations have been described to be involved in protease inhibitor resistance (21), all known gag mutations associated with drug resistance lie outside the gag PCR amplicon target region used by the CAP/CTM assays (S. Rose, Roche Molecular Diagnostics, personal communication). Hence, mutations associated with drug resistance should not affect the test performance. Moreover, dilution with HIV-1 RNA negative EDTA plasma might have influenced the efficiency of the extraction and amplification. Nevertheless, even with a possibly biased sample pool, we were able to clearly demonstrate the underquantification problem with CAP/CTM. (iii) The present study did not evaluate the performance of the tests in the low quantification range. In order to find significant differences in quantification, specimens with higher nominal viral loads were used. However, as seen with the first version of this kit (16, 18), the second version will probably also show an increased sensitivity at low viral loads in comparison with the CAP/CA PHS assay, as this fact is linked to the real-time technology. The important practical impact of this issue was recently discussed (8, 13).
In conclusion, the criteria needed to consider the new method equivalent to the routine method are fulfilled only for CAP/CTM v2.0. We clearly demonstrated the underquantification issue with the first version of the CAP/CTM test, not only in 13 samples of patients infected with a non-B subtype but also in 7 samples from patients infected with subtype B. Although the samples tested are all of Belgian origin (or at least collected in Belgium), the CAP/CTM test is not adequate for HIV-1 viral load testing on a general, worldwide basis. With the CAP/CTM v2.0 test no underquantified sample was identified compared to the routine test CAP/CA PHS. Thus, only the second version of this kit can be used for routine HIV-1 viral load testing in a routine clinical laboratory.
In spite of the good correlation between CAP/CA PHS and CAP/CTM v2.0, clinical biologists and clinicians should remain aware of the high degree of genetic variability of HIV-1 and the difficulties that this entails in the design of appropriate primer and probe sets for a real-time PCR that should be able to quantify all known HIV-1 variants. When viral load status does not develop as expected from clinical picture and/or CD4+ T-cell counts, other plasma HIV-1 RNA assays should be used in order to detect possible variation of the virus, whatever its subtype or origin.
We thank Roche Diagnostics, Ltd., Rotkreuz, Switzerland for supplying the reagents used in this study, for sequencing the severely discrepant samples and for lending us the Cobas TaqMan 48 analyzer.
We gratefully acknowledge A. Wyns, C. Vanneste, M.-H. Jurion, K. Miller, and N. Gijbels for their excellent technical assistance. We thank A. Sasse from the Belgian Institute of Public Health for his support.
Published ahead of print on 17 February 2010.
†The authors have paid a fee to allow immediate free access to this article.