Immune Thrombocytopenia (ITP) is a heterogeneous clinical entity for which no simple diagnostic test exists. The Immature Platelet Fraction (IPF), a marker of Reticulated Platelets (RP), is a good indicator of thrombopoiesis and can identify accelerated destruction of platelets by demonstrating compensatory increased platelet production. Measuring IPF is not routinely available in clinical practice; it has proven useful in studies of thrombocytopenia and the novel thrombopoietic agents. A multicenter clinical trial would be needed to explore IPF in management of patients with ITP; such a trial would require central testing. Therefore, we sought to assess the reliability of measured platelet counts and IPF in whole-blood filled EDTA tubes when the same tube is run freshly and again 24 hr after venipuncture. Based on analysis of our 103 samples, it is clear that EDTA tubes are stable at room temperature for 24 hr. Therefore, the results can be used to estimate thrombopoiesis when measured within 24 hr after phlebotomy.
Immune thrombocytopenia (ITP) is heterogeneous and affects both children and adults. It is characterized by autoantibody mediated increased platelet destruction, and reduced platelet production, leading to thrombocytopenia and its sequelae [1–3]. No standard diagnostic test, i.e., a platelet antibody test, exists to differentiate thrombocytopenia seen in ITP from other disorders. The diagnosis rests on applying clinical parameters, most of which are by exclusion, as outlined by various professional societies and recently updated [4–6].
In 1969, Ingram and Coopersmith identified certain platelets as having a coarse, punctuated condensation, i.e., reticulum, by using a methylene blue dye. The study suggested those platelets contained an increased amount of cytoplasmic RNA, reflecting active thrombopoiesis . Lee et al., in 1986, reported detection of platelet nucleic acid via flow cytometry using thiazole orange dye . Kienast and Schmitz showed that RNA-rich platelets measured with flow cytometry provided information on the thrombopoietic activity of patients with thrombocytopenic states . Ault et al. espoused that analysis of these reticulated platelets provided a good indication of the rate of platelet production . Furthermore, Rinder et al., in 1993, found that flow cytometric analysis of reticulated platelets was a better discriminant than platelet associated IgG in diagnosing ITP .
Subsequently, Sysmex corporation developed a method of measuring an entity called the IPF using a technology similar to thiazole orange staining of platelets; this technology used a proprietary dye and laser imaging to measure its association with platelets. Studies have shown that the IPF is virtually indistinguishable from the RP . Additionally, research has provided evidence on the utility of measuring IPF in the diagnosis of ITP [13–15]. The advantage of the IPF is that it can be routinely obtained as part of the complete blood count using a Sysmex autoanalyzer, such as the XE2100.
Despite the aforementioned studies and others that indicate the utility of measuring RNA-rich platelets (otherwise known as RP or IPF), the IPF has not been widely adopted into clinical practice. One important reason for this lack of utilization is that measurement of IPF requires special technology, which is not routinely available in most hospital laboratories.
Routine measurement of the IPF in patients with ITP would potentially be useful, including in clinical trials. To utilize measurement of reticulated platelets, there would need to be a sufficiently large study to have adequate power. Such a study would ideally be completed at a central location, with samples shipped from various centers to allow for the study itself to be multicentered. The IPF might be particularly useful in exploring the newly licensed thrombopoietic agents [16–18], determining if the IPF parameter discriminates between different causes of thrombocytopenia, and investigating whether it would predict response to various therapies.
The aim of this project was to assess the reliability of measuring platelet counts and IPF in a whole-blood filled EDTA tube when the sample is run on a Sysmex autoanalyzer 24 hr after it was drawn. To determine the stability of the IPF in patients with ITP, samples were run fresh within hours of venipuncture (TO) and compared to the same sample run 24 hr later (T24). No mock or overnight shipping was used.
A total of 103 blood samples were collected by a phlebotomist from 74 subjects with ITP, ranging in age from 4–83 years (mean 41 years) during regularly scheduled clinic visits to the Platelet Disorder Center of New York Presbyterian-Weill Cornell Medical Center between the months of January and March 2007. Each individual subject contributed one or two samples to the database. A total of 45 of the samples were from subjects represented only once in the database; 29 subjects were represented twice. A total of 22 of the 74 total subjects were children, ages 4–21; four children had repeat sampling. For subjects represented twice in the catalogue of samples, the time between obtaining the samples ranged from 1 to 63 days.
The mean IPF% value for all subjects at T0 was 17.64 +/− 15.26 (standard error 1.5); the mean IPF absolute for all subjects at T0 was 7.72 +/− 8.31 (standard error 0.82). The mean IPF% value for all subjects at T24 was 18.38 +/− 13.96 (standard error 1.3); the mean IPF absolute for all subjects at T24 was 8.62 +/− 8.61 (standard error 0.85). Considering all 103 samples, the Pearson correlation for platelet counts at T0 and T24 was 0.995 [r(103) = 0.995] (Fig. 1a). The Pearson correlation for absolute IPF at T0 and T24 was 0.930 [r(103) = 0.930] (Fig. 1b); for IPF%, the correlation at T0 and T24 is 0.941 [r(103) = 0.941] (Fig. 1c). These three correlations are all statistically significant at the level of P < 0.001. When considering the 74 samples that do not include repeats (i.e., 45 unique subjects, the first sample for the 29 subjects with repeated sampling), the values for Pearson correlation are essentially unchanged: r(74) = 0.995 for platelets T0 to T24, 0.922 for IPF absolute T0 to T24, 0.995 IPF% for T0 to T24.
For IPF absolute, there seems to be a discrepancy between T0 and T24 of greater than 10 (× 109/L) when the IPF values are larger than 30 (× 109/L) (Fig. 1b outlier values); for IPF%, the discrepancy between T0 and T24 values appears approximately 10% when the IPF% values are larger than 40% (Fig.1c, outlier values). Thus, the accuracy of repeated measurements seems to decrease at the very infrequent times when the values increase, with less correlation when A-IPF >30 (× 109/L) and IPF% >40%.
Watanabe et al. endorsed the reproducibility and stability of the IPF, but no specific data were provided . Briggs et al. reported that the IPF remained stable over two days when seven blood samples were stored at room temperature; there was no consistent increase or decrease in the IPF values 0.5–48 hr after sampling . The current study is from a larger number of samples (n = 103), patients with ITP, and complete data is provided. Furthermore, the estimates for IPF% and A-IPF are similar to that cited elsewhere in the literature, specifically Briggs et al. 2004 article (Table I).
|Mean IPF absolute [ × 109/L] (range)||Mean IPF% (range)|
|Briggs et al 2004 article for pts with plt count >50 × 109/La||8.4 (1.6–38.6)||16.8 (2.3–52.1)|
|Briggs et al 2004 article for pts with plt count <50 × 109/La||7.8 (1.6–34.3)||22.3 (9.2–33.1)|
|Current Study||7.7 (1–50.1)||17.6 (0.9–68.8)|
It is worth noting that an investigator can either use percent IPF or absolute values when translating laboratory parameters into clinical practice. Although we have presented data for both, our practice is to preferably utilize absolute values in clinical scenarios. Mathematically, as IPF% is defined at A-IPF/Platelet count, measures of IPF% may seem inordinately elevated in a patient with an extremely low platelet count. Complete considerations of the merits and clinical significance of IPF% versus IPF absolute in patients with ITP is outside the scope of this report since these blood samples were mostly only taken at one time point during a subject's treatment.
A potential confounder is that 29 subjects were represented twice; thus, trends of platelet and platelet precursor stability in certain individuals may be over represented. Furthermore, those patients who are represented twice may have worse disease, require more frequent visits, and may be more likely to be consented for inclusion in the study. The limit of only two samples per subject, however, makes this unlikely. Moreover, the analysis of the data with each of the 74 samples without repeats yielded essentially identical conclusions to that with all 103 samples. Another potential confounder is that a T24 sample may be run at slightly longer than 24 hr after phlebotomy [there is a 30 hr window if the specimen was drawn at 9 a.m. and then run at 3 p.m. the following day]. However, this potential confounder would likely give worse results for stability of measurement, which was not born out in the data.
Based on the aforementioned results and when performed under conditions used in this study, we conclude that platelet count and absolute immature platelet fraction are reliable when measured at one day later than collection. Power appears to be sufficient in this study, since the “n” is 103. These results should allow for further multicenter studies to proceed in which the IPF is thought to be relevant in patients who have ITP, or in discriminating between causes of decreased platelets, by measuring laboratory parameters at a standardized, centralized location. The sample can be shipped overnight at room temperature, and we propose that results would still be valid.
This report lends credence to the reliable determination of the A-IPF and IPF%, despite storage of a sample for 24 hr prior to obtaining the measurement. This finding could be of use in future studies in patients with ITP.