ABSTRACT
Congenital cytomegalovirus (cCMV) infection is a major cause of childhood hearing loss and neurodevelopmental delay. Identification of newborns with cCMV infection allows provision of beneficial interventions. However, most infants with cCMV infection have subclinical infection and go undiagnosed. Thus, expanded neonatal CMV testing is increasingly recommended. Saliva is an attractive sample type for CMV testing of newborns, because it is easier to collect than urine and more sensitive for CMV detection than dried blood spots. We evaluated the Alethia CMV assay, a rapid, easy-to-use loop-mediated isothermal amplification method for qualitative detection of CMV DNA in neonatal saliva samples. Saliva swabs were collected prospectively from newborns <21 days old and tested by the Alethia assay according to the manufacturer’s instructions. Archived saliva swabs from newborns with cCMV infection were also tested retrospectively. A composite reference method (CRM; two validated PCR assays followed by bidirectional sequencing of amplicons) was performed on all samples as the reference standard comparator. Of 1,480 prospectively collected saliva swabs, 1,472 (99.5%) were negative by both the Alethia assay and CRM, 5 (0.34%) were positive by both the Alethia assay and CRM, and 3 (0.20%) were positive only by the Alethia assay. All 34 (100%) archived swabs from newborns with cCMV infection were positive by both the CRM and the Alethia assay. Overall, the Alethia assay showed 100% and 99.8% positive and negative agreement with the CRM, respectively. The Alethia CMV assay is an accurate method for identifying neonates with cCMV infection and, given its simplicity, appears suitable for CMV testing using neonatal saliva outside a reference laboratory, including remote and resource-limited settings.
INTRODUCTION
Congenital cytomegalovirus (cCMV) infection occurs in an estimated 0.6 to 0.7% of live births in high-income countries and 1% to 5% of live births in developing countries (1, 2) and is a major cause of childhood hearing loss, intellectual disability, and other permanent neurologic sequelae (3). Identification of newborns with cCMV infection is desirable because early interventions, including antiviral therapy for selected infants and identification of hearing loss that develops during early childhood, have been shown to result in better hearing and developmental outcomes (4–7). However, most newborns with cCMV infection are asymptomatic or have mild, nonspecific findings (8), such that only a small fraction of infections are identified in the absence of coordinated neonatal CMV testing programs (9, 10).
Definitive diagnosis of cCMV infection requires the detection of the virus in a specimen collected before 3 weeks of age, because a positive test result from samples collected later may reflect postpartum infection, which does not incur the neurodevelopmental risks (11). The recommended clinical specimens for the diagnosis of cCMV infection are urine and saliva (5, 6, 11), because these contain high quantities of virus, resulting in high sensitivity compared to dried blood spots (DBS) (12, 13). Saliva swab collection is more convenient than urine collection from newborns, making saliva the preferred specimen type for routine CMV testing (14–17).
The Alethia CMV DNA amplification assay uses loop-mediated isothermal amplification (LAMP) technology with the ability to provide results in less than an hour. The Alethia CMV assay targets a 194-bp region of the UL33 gene. The amplicon sequence aligns with ∼260 strains of human herpesvirus 5 (CMV) at 99 to 100% homology for which a complete genomic sequence was reported. Because of its ease of use and short turnaround time, the Alethia assay allows CMV testing of saliva swabs from newborns outside reference or other subspecialized laboratories, which may be advantageous for efforts to diagnose a larger proportion of infected children. Here, we report the results of a recent clinical study that resulted in the Alethia CMV assay being the first assay to receive FDA clearance for detection of CMV in neonatal saliva swabs.
MATERIALS AND METHODS
Study design.Approval from all of the relevant institutional review boards and ethics committees was obtained for all parts of this study. After written informed consent was received from their parents or guardians, neonates less than 21 days of age were enrolled at seven investigational sites—University of Utah, BC Children’s Hospital, Prince of Wales Hospital, University of Alabama at Birmingham, University of Bologna, Med Fusion, and Sacred Heart Hospital—for collection of deidentified saliva swab specimens. Specimens were excluded if they were collected later than 21 days of age or less than 1 h after breastfeeding or if the package insert instructions (18) were otherwise not followed. In addition, to increase the number of positive samples, deidentified, archived samples from newborns with cCMV infection were eligible for testing if they were collected and stored in a manner consistent with the package insert instructions. The CMV-positive archived specimens used in the clinical trial were obtained from one of the clinical trial sites that enrolled prospectively collected and tested specimens. These archived specimens were distributed among all the sites and tested using the Alethia CMV assay.
Specimen collection and storage.Nylon flocked swabs (Copan FLOQSwab; catalog no. 553C) were used to collect saliva from each participant. After collection, the swabs were either transported dry or placed in 1 ml of transport medium (Copan UTM 1-ml minitube; catalog no. 360C; BD UVT, catalog no. 220526; or Puritan UniTanz-RT UTM, catalog no. UT116). Specimens were tested as soon as possible after collection, prior to which they were stored for up to 48 h at room temperature or 7 days at 2 to 8°C; specimens that could not be tested within 7 days were stored at or below −20°C.
Assay procedures.Testing was performed using the Alethia CMV DNA amplification assay at each site according to package insert instructions provided by the manufacturer (18). The saliva swab samples were added to the tube labeled buffer 1, vortexed for 10 s, and incubated for 2 min. Fifty microliters of buffer 1 was then added to the tube labeled buffer 2 and vortexed for 10 s. Fifty microliters of buffer 2 was transferred to both the test and control chambers of the Alethia test device, which was then placed in the Alethia instrument to initiate the run. Each batch of specimens was run with external controls according to the manufacturer’s instructions. Specimens without amplification of the external controls or without amplification of the mitochondrial DNA internal controls were reported as invalid. Testing of specimens with invalid results was repeated. The composite reference method (CRM) used as the comparator consisted of two in-house-validated qualitative PCR assays followed by bidirectional sequencing on the positive PCR samples. The comparator assays were validated following FDA guidance for reference assays, which included an evaluation of the limit of detection and demonstration of analytical reactivity. Sequences generated from bidirectional sequencing met stringent acceptance criteria as per the FDA guidelines.
After Alethia CMV testing at the sites, the remaining sample was sent to the study sponsor, where the CRM was then performed. The target sequences of the CRM PCR assays have no DNA sequence overlap with the Alethia CMV assay region and instead target the UL31 and UL87 genes of the CMV genome. The sizes of the amplification products from the UL31 and UL87 sequences were 364 and 419 bp, respectively. For the UL31 PCR assay, the forward (5′-TGTCCATCACCGAGCAGTA-3′) and reverse (5′-ACGGCGTAATGCGGGCA-3′) primers were each added at a final concentration of 300 nM along with a fluorescein (6-carboxyfluorescein [FAM])- and Iowa Black (quencher)-labeled probe (5′-FAM-TGCGGCTACAAGTACGAGTGGTCC-black hole quencher [BHQ]-3′) at a final concentration of 100 nM. For the UL87 PCR assay, the forward (5′-GGCTGGTTCAAAGCGGCT-3′) and reverse (5′-CGGCCCAGGTGATCCAAT-3′) primers were each added at a final concentration of 300 nM along with a fluorescein (FAM)- and Iowa Black (quencher)-labeled probe (5′-FAM-CAGGCAAGCACGCAGAAACAGCAC-BHQ-3′) at a final concentration of 100 nM. For both PCR assays, SensiFAST (Meridian Biosciences, Inc.; Bioline catalog no. BIO-86020) containing hot-start Taq polymerase along with other PCR buffer components was added at a final concentration of 1×. Thermocycling was performed on a Rotor-Gene Q (Qiagen, Inc.) with the following program: initial hold at 95C for 7 min for enzyme activation followed by 50 cycles of denaturation at 95°C for 30 s (UL31 PCR) or 20 s (UL87 PCR) and 60°C for 1 min. Specimens that generated a positive amplification signal from either of these two PCR assays were further analyzed by performing double-stranded-DNA sequencing followed by BLAST analysis, whereas specimens negative by both PCR assays were not further analyzed. Sequencing primers specific to the UL31 364-bp (forward, 5′-TCAAACGTGACGGAGAAAGC-3′, and reverse, 5′-CCATGACCAGTTGTGTGTG-3′) and UL87 41- bp (forward, 5′-ATTGTGCCGCGCGTCTC-3′, and reverse, 5′-GACTTTTTTGTTCCGGGCC-3′) amplicons were used for double-stranded sequencing (Neogenomics, Inc.). Obtained nucleotide sequences at least 200 bp in length with a quality score (PHRED) of at least 20 were compared to a reference CMV sequence (19, 20). Sequences with an E value of 10−30 or less were considered positive (21) and were submitted to GenBank.
Sample size estimates and statistical analysis.Endpoints and sample sizes were determined based on requirements for FDA clearance, which was obtained through the de novo premarket review pathway. A minimum of 1,000 negative prospective CMV specimens were required to be tested to achieve the prespecified negative percent agreement (NPA) of 95%, with a lower 95% confidence interval (CI) bound of 85%. A minimum of five positive prospectively collected specimens was prespecified, which could be combined with an additional number of archived positive specimens to achieve 95% positive percent agreement (PPA), with a lower 95% CI bound of 85%. Data management and analyses were performed by the study sponsor.
RESULTS
A total of 1,616 specimens were collected for study. Of these, one specimen was excluded due to revocation of study consent and another 101 due to package insert violations, including inappropriate collection or storage. Of the 1,514 eligible specimens, 1,480 were collected prospectively and 34 were archived. These were from 715 females, 775 males, and 24 newborns of unknown sex. All eligible specimens were tested by the Alethia CMV assay and the CRM.
The overall performance comparison of the Alethia CMV assay to the CRM is shown in Table 1. The NPA compared to the CRM was 99.8% (1,472 of 1,475; 95% CI, 99.4% to 100%), and the PPA was 100% (39 of 39; 95% CI, 91.0% to 100%). There were 27 negative samples that were classified as invalid based on internal control amplification and detection; 26 (96%) of these invalid specimens were resolved as being negative upon repeat testing, for a total of one invalid result (0.06%) out of 1,514 specimens. There were three specimens that produced an amplicon by only one of the two comparator PCR assays that was not confirmed with sequencing and definition of the CRM algorithm, and these were called negative. These specimens were also negative by the Alethia CMV assay.
Overall performance of the Alethia CMV DNA amplification assay
Table 2 shows the stratified results of prospectively collected and archived specimens. Of 1,480 prospectively collected saliva swabs eligible for testing, 1,472 (99.5%) were negative by both the Alethia assay and the CRM, five (0.34%) were positive by both Alethia assay and CRM, and three (0.20%) were positive by Alethia assay but negative by the CRM. All 34 (100%) archived swabs from newborns with cCMV infection were positive by both the CRM and the Alethia assay.
Assay performance as a function of prospective and archived specimens
There was no statistically significant difference in the performance of the Alethia CMV assay with respect to sex; two of the discordant results were from females and the third was from males. Of the eligible specimens, 1,323 were collected as dry swabs and 292 were collected and placed in 1 ml of viral transport medium. No significant differences were observed between specimens collected dry and those in medium or between study sites (data not shown). Additional performance characteristics are available for review in the product insert (18).
DISCUSSION
In this study, we found that the Alethia CMV assay performs similarly to the reference standard of PCR for qualitative detection of viral DNA in neonatal saliva swabs. In-house PCR and commercial assays are the most common methods used for CMV testing of saliva and urine samples from newborns, having replaced antigen detection and viral culture due to their high sensitivity and wider availability (12, 22). However, the complexity of CMV PCR testing typically requires it to be performed in reference laboratories. Furthermore, because there were previously no FDA-cleared assays for detection of CMV in neonatal saliva, laboratories were forced to conduct their own validation of PCR assays developed in-house or modified commercial assays for other specimen types. In contrast, the Alethia CMV assay is a simpler, more rapid test, and it was found to be accurate and robust compared with PCR, with an invalid-test-result rate of 0.06%. As such, the Alethia CMV assay may be attractive in order to avoid the need for internal assay validation, when rapid results are desirable or where testing at reference laboratories is not feasible.
Expanded testing of newborns for cCMV infection is increasingly recommended, whether via a universal approach or by targeting neonates with risk factors such as a failed hearing screen (5, 6, 23). The benefits of diagnosing cCMV infection include antiviral therapy for symptomatic newborns and early directed care for all infected children to improve developmental outcomes (10, 24), as well as favorable estimates of cost-effectiveness in various settings (15, 25, 26). A saliva swab is often the preferred sample type for neonatal CMV testing (5, 6, 11), since it is more practical to collect from neonates and more sensitive than DBS (12, 13). The results of the Alethia CMV assay are used to aid in the diagnosis of cCMV infection and are not intended to be used as the sole basis of patient management, which should incorporate confirmatory testing and other clinical data (5, 6, 11). In some cases, given the possibility of false-positive saliva tests due to CMV in breast milk (27, 28), confirmatory testing using urine is recommended for diagnosing cCMV infection (5, 6, 23). The need for this confirmatory testing may be minimized by waiting at least 1 h after nursing prior to saliva swab collection, which is recommended when using the Alethia CMV assay. However, our findings indicate that the Alethia CMV assay performs similarly to PCR methods and therefore has the same limitations of the current standard-of-care saliva swab tests for the diagnosis of cCMV infection.
The strengths of this study include the large number of prospectively collected neonatal saliva specimens from multiple centers, supplementation with additional CMV-positive specimens, and the use of a reliable comparator method. Limitations include the lack of viral load data, the absence of confirmatory urine testing, and other clinical information about the infants, such as symptoms of cCMV infection. Thus, we are unable to comment on the rate of false positives or negatives or to calculate positive and negative predictive values of the assay.
In summary, the Alethia CMV assay reliably detects CMV in saliva of neonates <21 days of age and is the first FDA-cleared assay for this indication. This assay represents an option for rapid, accurate CMV testing of newborns outside a reference laboratory and may therefore assist in the development of efforts to more effectively identify children with cCMV infection and provide them with the best available care.
ACKNOWLEDGMENTS
This study was funded by Meridian Bioscience, Inc. S.G. reports consulting fees from Merck, GSK, Moderna, and Curevo and research support from Merck and GSK. A.P. reports consulting fees from Merck and Roche. S.B.B. reports consulting fees from Merck, Sanofi, and Moderna and research support from Merck. W.R. reports being an unpaid member of advisory boards for Genetic Signatures, Moderna and Roche. T.L. reports consulting fees from Merck.
We thank Wendy van Zuylen, Heather F. Thiesset, Gabrielle Turello, Silvia Felici Maria, Grazia Capretti, Sunil Pati, Misty Purser Latting, Neelam Dhiman, and Donna Mayne for their assistance with the study.
FOOTNOTES
- Received 10 December 2019.
- Returned for modification 30 December 2019.
- Accepted 17 January 2020.
- Accepted manuscript posted online 22 January 2020.
- Copyright © 2020 American Society for Microbiology.
