Frozen or Chilled? Study Maps How Urine Storage Shapes a Key Schistosomiasis Test

Schistosomiasis remains one of the most stubborn parasitic diseases on the planet, infecting more than 200 million people worldwide and exacting its heaviest toll across sub-Saharan Africa. As elimination programs push into low-endemic and near-elimination settings, the diagnostic tools they rely on must become ever more sensitive, because the worms that remain after mass drug administration are few, scattered, and hard to catch. One of the most promising tools in this arsenal is the up-converting reporter particle lateral flow test for circulating anodic antigen, or UCP-LF CAA, a urine-based assay that detects a molecule shed by living Schistosoma worms. Now, a team of researchers working on Pemba Island in Tanzania has asked a deceptively simple question that could shape how elimination campaigns collect and ship samples for years to come: what happens to the test’s results when urine sits in a freezer, a refrigerator, or a cool box before it is analyzed?

The answer, published in Frontiers in Parasitology, is reassuring but nuanced. Freezing urine at minus 20 degrees Celsius for five to seven days caused a small but statistically significant drop in the proportion of samples testing positive for CAA, while storage under non-frozen conditions at temperatures between 4 and 25 degrees Celsius for roughly five weeks produced results that were generally comparable to those from freshly tested urine. The findings suggest that laboratories and control programs may have more flexibility in how they handle urine samples than the conventional freeze-first approach implies, though the edge of the assay’s detection range remains a fragile zone where storage choices can matter.

To understand why this matters, it helps to know what the CAA test actually measures. Circulating anodic antigen is a glycan-based molecule released by adult Schistosoma worms into the bloodstream of their host. Because it is only produced by living worms, its presence signals an active infection rather than a past one, making it a valuable marker for monitoring treatment efficacy and for surveillance in settings where egg-based microscopy misses low-intensity infections. Conventional parasitological methods such as urine filtration for Schistosoma haematobium eggs suffer from limited sensitivity precisely in the situations that matter most during elimination: after praziquantel treatment, when worm burdens are low and egg excretion is intermittent, and in low-endemic areas where most infections are light. The UCP-LF CAA test, which uses up-converting phosphor particles as a highly sensitive reporter label, has demonstrated strong performance across all known Schistosoma species, including hybrids and veterinary species, and can be run in laboratories with relatively basic equipment.

The study, led by Pytsje T. Hoekstra of the Leiden University Center for Infectious Diseases and colleagues from Leiden University Medical Center, the Public Health Laboratory-Ivo de Carneri on Pemba, the Swiss Tropical and Public Health Institute, and the University of Basel, was embedded in a larger prospective diagnostic evaluation among school-aged children in two schools on Pemba Island, conducted in February 2025. Pemba, part of the Zanzibar archipelago of the United Republic of Tanzania, is a near-elimination setting for S. haematobium, which makes it an ideal proving ground for diagnostics that must perform at the very limits of detection. The researchers collected urine from 272 children and split each sample into aliquots that followed different paths before testing.

The first aliquot was tested within four hours of collection, on the day itself, at the Public Health Laboratory-Ivo de Carneri on Pemba. The second was frozen within four hours, held at minus 20 degrees Celsius for five to seven days, thawed, and then tested at the same laboratory during the following week, reflecting the conventional approach used for CAA testing in many research and surveillance programs. A third aliquot, available for a subset of 153 samples with sufficient remaining urine volume, was stored at 4 degrees Celsius, transported to the Netherlands under cooled conditions with ice packs at the end of the second week, and tested at Leiden University Medical Center within two to three weeks of arrival, roughly five weeks after the sample was originally collected. This third condition was introduced partway through the study as a pragmatic test of a realistic field-to-laboratory workflow, mimicking what happens when freezing is not available in the field and samples must simply be refrigerated and shipped.

The assay itself follows a well-established protocol. Urine is pre-treated with trichloroacetic acid to release CAA from immune complexes and precipitate proteins, after which the clear supernatant is concentrated using centrifugal filtration devices with a 10-kilodalton cutoff. The concentrated sample is then mixed with assay buffer containing up-converting phosphor particles conjugated to anti-CAA antibodies, lyophilized into pellets for stability. Lateral flow strips bearing a test line of anti-CAA antibodies are added, and immunochromatography proceeds for at least three hours before the strips are scanned with a reader adapted for the up-converting label. Each run includes a serial dilution series of a trichloroacetic acid-treated crude adult worm antigen preparation spiked into CAA-negative urine, which serves both to validate the assay’s cut-off, set at 2 picograms per milliliter for the dry format used here, and to quantify the CAA concentration in unknown samples.

The headline numbers tell a clear story. In the full dataset of 272 samples, 44, or 16.2 percent, tested positive when urine was fresh, but only 35, or 12.9 percent, tested positive after freezing. Nine of the samples that flipped from positive to negative after freezing were low-positive samples with CAA concentrations between 2 and 5 picograms per milliliter, sitting close to the assay’s cut-off. Not a single fresh-negative sample turned positive after freezing, meaning the discordance ran entirely in one direction. A McNemar’s test on the paired proportions confirmed the difference was statistically significant, with a p-value of 0.004. Quantitative CAA concentrations showed a non-significant trend toward lower values after freezing, with a Wilcoxon signed-rank p-value of 0.051, and a Bland-Altman analysis found no statistically significant systematic bias, though fresh samples tended to have higher concentrations with a mean ratio of 1.18.

For the subset of 153 samples that also had a non-frozen stored aliquot, the picture was different in an interesting way. Positivity was 20 out of 153, or 13.1 percent, in fresh urine, 16 out of 153, or 10.5 percent, in frozen urine, and 24 out of 153, or 15.7 percent, in non-frozen stored urine, the highest of the three conditions. Compared with fresh samples, only 2 flipped from positive to negative after non-frozen storage while 6 flipped the other way, a difference that was not statistically significant. Against frozen samples, however, the non-frozen condition produced significantly more positives, with 1 sample going from positive to negative and 9 going from negative to positive, yielding a McNemar p-value of 0.02. Across all pairwise comparisons, Spearman rank correlations between quantitative CAA concentrations were strong, ranging from 0.86 to 0.93 with p-values below 0.0001, and Cohen’s kappa for qualitative agreement ranged from 0.71 to 0.87, indicating substantial to near-perfect concordance.

The authors interpret these findings with appropriate caution. The reduction in positivity after freezing was concentrated among samples with CAA concentrations close to the 2 picograms per milliliter cut-off, suggesting that analytical variability around the detection threshold, rather than wholesale degradation of the antigen, likely explains the lost positives. CAA is chemically a carbohydrate, which makes it remarkably resistant to degradation; its stability is famously illustrated by its detection in Egyptian mummies, where it has persisted for thousands of years. The comparable results between fresh and non-frozen stored samples indicate that CAA detection was maintained across a realistic field-to-laboratory workflow involving refrigerated storage, cooled transport, and international shipping, though the authors note that this comparison cannot fully separate the effect of storage from the effects of transport, timing, and laboratory conditions, since the non-frozen samples were analyzed in Leiden while the fresh and frozen ones were tested on Pemba.

Several limitations shape the conclusions. The non-frozen condition was introduced halfway through the study and assessed only in a pragmatic subset defined by available urine volume, which could have introduced bias even though selection was not based on participant characteristics or diagnostic outcomes. More fundamentally, the study was designed to compare storage conditions rather than storage durations: fresh samples were tested within four hours, frozen samples after five to seven days, and non-frozen samples after roughly five weeks, with intervals dictated by logistics rather than prospective assignment. The effects of storage condition and storage duration therefore cannot be disentangled, and no conclusions can be drawn about the impact of longer-term non-frozen storage, prolonged freezing, or repeated freeze-thaw cycles. The authors call for further studies addressing these gaps, noting that bacterial or fungal growth during prolonged non-frozen storage could potentially affect CAA detection and that standardized storage and transport procedures remain important for preserving sample integrity.

Even with those caveats, the practical implications are significant for elimination programs. The results suggest that urine storage strategies for delayed centralized CAA testing can be adapted to local circumstances: where freezers are available, samples can be frozen with the understanding that a small fraction of very low-positive results may be missed; where they are not, refrigerated storage and cooled transport over several weeks appear to preserve detection well. As schistosomiasis programs move from control toward elimination, and as the UCP-LF CAA test finds growing use in surveillance, sample pooling, post-treatment follow-up, pharmacokinetic studies, migrant health, and even controlled human infection models, knowing that the antigen in a child’s urine sample from Pemba can survive a five-week journey to a Dutch laboratory without meaningful loss of signal is the kind of unglamorous but essential knowledge on which elimination campaigns are built.

Subject of Research: Effect of urine storage conditions on circulating anodic antigen detection with the UCP-LF CAA test for schistosomiasis diagnosis

Article Title: Effect of urine storage conditions on circulating anodic antigen detection using the UCP-LF CAA test

Article References: Hoekstra, P. T., van Dam, G. J., Blokland-Flink, M., de Jong, D., Ali, S. M., Ali, M. N., Ndum, N. C., Knopp, S., Corstjens, P. L., & van Lieshout, L. (2026). Effect of urine storage conditions on circulating anodic antigen detection using the UCP-LF CAA test. Frontiers in Parasitology, 5, Article 1894516. https://doi.org/10.3389/fpara.2026.1894516

Image Credits: AI Generated

DOI: 10.3389/fpara.2026.1894516

Keywords: schistosomiasis, circulating anodic antigen, UCP-LF CAA, urine storage, diagnostics, Schistosoma haematobium, Pemba, Tanzania, elimination, freezing, surveillance, neglected tropical diseases