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Clinical Chemistry and Laboratory Medicine (CCLM)

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Volume 52, Issue 8


Reticulated platelets: analytical aspects and clinical utility

Johannes J.M.L. Hoffmann
  • Corresponding author
  • Abbott Diagnostics Division, Abbott GmbH and Co. KG, Max-Planck-Ring 2, 65205 Wiesbaden-Delkenheim, Germany
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Published Online: 2014-05-07 | DOI: https://doi.org/10.1515/cclm-2014-0165


Reticulated platelets are immature platelets circulating in blood; they reflect the activity of megakaryopoiesis in the bone marrow. Therefore, they can be used as a non-invasive test in patients with thrombocytopenia in various clinical conditions. The preferred method of analysis is by flow cytometry. However, there is an evident lack of analytical standardization, making it difficult to compare results obtained in different laboratories. Currently, two types of hematology analyzers are on the market offering fully automated measurement of reticulated or immature platelets: the high end analyzers manufactured by Sysmex (XE- and XN-series) and Abbott (CELL-DYN Sapphire). Although the methods are essentially different and cannot be used interchangeably, both have been proven to have clinical utility. Reticulated or immature platelet assays are useful for the differential diagnosis of thrombocytopenia and for monitoring bone marrow recovery after chemotherapy or stem cell transplantation. These assays may aid clinicians in platelet transfusion decisions when recovery from thrombocytopenia is imminent. In addition, preliminary findings indicate that there is a rationale for reticulated or immature platelets for risk stratification in acute coronary syndromes and for monitoring the effect of treatment with antiplatelet drugs in patients with coronary artery diseases. The aim of this paper is to present the present technology available for measuring reticulated platelets as well as an overview of the current status of clinical application. This overview also indicates that more research is needed before reticulated or immature platelet assays can be applied in other clinical conditions than thrombocytopenia and after transplantation.

Keywords: diagnostic use; immature platelets; megakaryopoiesis; reticulated platelets; thrombocytopenia

Formation and maturation of megakaryocytes

The study of blood platelets or thrombocytes started in the late 19th century, when Wright had developed his variant of the Romanovsky stain that allowed detailed observations of the smallest blood elements [1]. Platelets represent the terminal stage of megakaryopoiesis, a highly complex sequence of events that can be traced back to the generation of the pluripotent hematopoietic stem cell in bone marrow. This pluripotent stem cell proliferates and differentiates via several intermediates into a megakaryoblast and eventually into a megakaryocyte (Figure 1). Megakaryopoiesis is regulated by various growth factors and cytokines, of which thrombopoietin (TPO) is the most important. TPO stimulates the number, the size and the ploidy of megakaryocytic cells and is the key regulator of platelet concentration in the circulation [2]. Once the megakaryoblast stage is reached, the cell loses its proliferative capacity and starts the maturation process. Megakaryopoiesis has a unique way of maturation that does not occur in other cell lines: endomitosis. This is nuclear division without cell division; the cell is multiplying its nuclear material and increases its cytoplasm, all within the same cell. Finally, maturation results in a megakaryocyte that possesses multiple nuclear copies and abundant cytoplasm. The nuclear ploidy of a megakaryocyte is normally between 8N and 64N (median 16N), whereas higher and lower ploidy may occur in various pathological conditions. Depending on the organism’s need of new platelets, endomitosis stops and the formation of platelets commences (Figure 1). First, there is intracytoplasmic formation of channel-like structures composed of lipids, called the membrane demarcation system [3]. These lipids later assemble into membrane bilayers and eventually form the cell membranes of platelets when the megakaryocyte cytoplasm starts to disintegrate. In this phase there is also active protein synthesis and cytoskeletal assembly.

Schematic representation of megakaryopoiesis. After Kuter [2].
Figure 1

Schematic representation of megakaryopoiesis. After Kuter [2].

Platelet production by megakaryocytes

Eventually, megakaryocytes form pseudopodia-like extensions protruding into sinuses and release platelets into the extracellular space. How this exactly takes place is not fully elucidated [4–6]. A healthy human produces approximately 1–2 million platelets per second. In response to thrombocytopenia, platelet production can be accelerated by up to a 10-fold increase [7].

Megakaryocyte cytoplasmic volume expands synchronized with nuclear ploidy and this eventually determines the number of platelets that a megakaryocyte will produce. A single megakaryocyte can generate up to 5000 platelets [1, 2]. In steady-state conditions, the platelet production rate is aimed at keeping the total circulating platelet mass (platelet number×mean platelet volume, also called plateletcrit) constant [1]. In conditions of stress, platelets are released from megakaryocytes at an earlier stage, which results in platelets that are larger than normal.

Each individual has its own personal setpoint for platelet count and platelet volume; these are largely under genetic control [8]. As a consequence, intra-individual variations in platelet count are quite small in comparison with the population reference ranges. In the normal population, platelet count is inversely correlated with mean platelet volume (MPV), and consequently the total circulating platelet mass is less variable between individuals than platelet count.

Physiology of reticulated platelets

When platelets are released from the megakaryocyte cytoplasm, they still contain a small amount of RNA. For long it has been thought that this RNA was a vestigial remnant of megakaryocytic RNA, but there are recent indications that platelets are able using this RNA for protein synthesis [9]. Anyway, they represent the youngest platelets in the circulation and are named reticulated platelets (retPLT), in analogy with reticulocytes in erythropoiesis [10]. The concentration of retPLT in bone marrow is on the average 2–3 times higher than in peripheral blood, where they correlate with megakaryocyte numbers [11]. And whereas platelets persist in the circulation for 7–10 days, retPLT have a much shorter lifespan (<1 day). Therefore they can act as a marker of megakaryopoietic activity in the bone marrow [12, 13], which gives retPLT clinical and diagnostic utility.

Reticulated platelet methods – flow cytometry

The initial description of retPLT dates back to 1969, when Ingram and Coopersmith studied a canine model of acute blood loss and observed coarsely punctuated reticulum in platelets after supravital staining of blood with new methylene blue [14]. This technique was initially also used in human blood, but it is obviously not well suited for routine applications.

Kienast and Schmitz initiated a breakthrough in the field when they described a flow cytometric technique for analyzing retPLT, based on RNA staining by thiazole orange [15]. In subsequent years, several research groups published their findings in a wide variety of conditions like thrombocytopenia [16–21], thrombocytosis [22, 23], after stem cell transplantation [24–27], hereditary platelet diseases [28, 29], thrombo-embolic disorders [30, 31], kidney disease [32–34], preeclampsia [35], hyperthyroidism [36] and in healthy and thrombocytopenic neonates [37, 38]. The overall conclusion from these studies is that retPLT in blood represent a useful non-invasive marker of megakaryopoietic activity in the bone marrow. However, it also became evident that the flow cytometric assay was prone to methodological variation, which made it difficult to compare results obtained with different assays. For example, the normal reference range was reported to range between 1% and 15% [23, 39]. This wide range can be explained by lack of standardization of the methods. Many factors have been identified that contribute to this analytical issue: the type and concentration of fluorescent dye, incubation time and temperature, fixation, RNAse treatment and the flow cytometric data analysis, including gating and threshold settings [17, 18, 39–41]. One of the major problems is that platelets show non-RNA-specific binding of fluorescent dye, resulting in background staining, which is size-dependent [39, 42, 43]. This issue can be solved by optimizing the assay conditions and, in particular, by applying a two-dimensional gating process [39]. Nevertheless, standardization attempts for improving concordance between laboratories were unsuccessful, even when using the same protocol [9]. Recently, new initiatives were undertaken that are aimed at developing a method with the potential to become a future international reference method [44, 45]. However, even if a standardized method would be available, it would carry the disadvantage of not being well suited for routine clinical applications, since flow cytometry requires a great deal of expertise and is hardly available for patient care on a 24/7 basis. For the time being, a surrogate level of standardization is possible, since two manufacturers offer hematology analyzers that are capable of measuring reticulated or immature platelets (Table 1).

Table 1

Overview of some key characteristics of methods for measuring reticulated or immature platelets.

Immature platelet methods – Sysmex hematology analyzers

The first available fully automated method for measuring reticulated platelets was in the R-3000, a dedicated reticulocyte analyzer developed by Toa Medical (later Sysmex) [46, 47]. The method used auramine O as a fluorescent RNA dye and a 488 nm Argon laser. By plotting forward light scatter (representing cell size) against fluorescence (RNA content), reticulated platelets could be distinguished from mature platelets. In normal healthy individuals the mean retPLT count was 0.98%–1.27% and thrombocytopenic patients with a variety of diseases had increased concentrations, in accordance with the flow cytometric results mentioned above [18, 47, 48]. Notably, a strong positive correlation existed between percentage retPLT and the fraction of large platelets, except in patients with aplastic anemia [47]. Furthermore, the large platelet fraction highly correlated with MPV; data on the correlation between retPLT and MPV were not provided, but one can reasonably assume that such correlation was present.

Automated measurement was later integrated into the Sysmex XE-2100 and XE-5000 hematology analyzers, as a part of the reticulocyte determination. This enabled automated quantification of what from then on was called immature platelet fraction (IPF) [49]. As these instruments employ a 633 nm diode laser as the light source, other dyes were needed and a mixture of polymethine and oxazine was selected. As before, a forward scatter versus fluorescence scatterplot defined platelets with the highest fluorescence intensity as immature platelets (Figure 2A). The reference values in a healthy population were clearly higher than with the previous R-3000 method: mean 3.4% (range 1.1%–6.1%) [49]. These reference values were later confirmed by others [50–54]. In all studies, patients with idiopathic thrombocytopenia (ITP) had increased IPF; in some patients IPF was even strongly increased, up to 50%–60% [49, 54, 55]. Such extreme values were later recognized to be artifacts due to interference by white blood cell fragments [56].

Scatterplots of the automated assays of immature and reticulated platelets currently available. In the Sysmex XE-series, immature platelet fraction (IPF) is determined in the reticulocyte assay, using forward scatter against fluorescence (A). The Sysmex XN-series measure IPF as part of the fluorescent platelets assay, again using forward scatter against fluorescence (B). In the Abbott CELL-DYN Sapphire reticulocyte assay, platelets and red blood cells are first separated using a side scatter against intermediate angle scatterplot (not shown) and then reticulated platelets are determined using fluorescence (FL1) against intermediate angle scatter (7°), correcting for size-dependent background staining (C). IPF, immature platelet fraction; PLT, platelets; RBC, red blood cells; RETC, reticulocytes; retPLT, reticulated platelets; WBC, white blood cells.
Figure 2

Scatterplots of the automated assays of immature and reticulated platelets currently available.

In the Sysmex XE-series, immature platelet fraction (IPF) is determined in the reticulocyte assay, using forward scatter against fluorescence (A). The Sysmex XN-series measure IPF as part of the fluorescent platelets assay, again using forward scatter against fluorescence (B). In the Abbott CELL-DYN Sapphire reticulocyte assay, platelets and red blood cells are first separated using a side scatter against intermediate angle scatterplot (not shown) and then reticulated platelets are determined using fluorescence (FL1) against intermediate angle scatter (7°), correcting for size-dependent background staining (C). IPF, immature platelet fraction; PLT, platelets; RBC, red blood cells; RETC, reticulocytes; retPLT, reticulated platelets; WBC, white blood cells.

With the introduction of the XN-series hematology analyzers, Sysmex changed the IPF method. It is now part of the fluorescent platelet assay, which uses a proprietary oxazine-based dye for staining RNA. Still, IPF is derived from the forward scatter (cell size) versus sideward fluorescence (RNA content) scatterplot [56, 57], as shown in Figure 2B. The moderate correlation in IPF between XE-2100 and XN is explained, at least partially, by reduced interference in the XN system [56]. Preliminary data suggest that the XN reference values of IPF are comparable with the XE-2100 method [56].

Reticulated platelet methods – Abbott CELL-DYN Sapphire

Apart from the Sysmex method discussed above, the Abbott CELL-DYN Sapphire is currently the only hematology analyzer capable of measuring retPLT. The assay is an integral part of the reticulocyte assay, which is based on the fluorescent dye CD4K530 [58] that is excited by a 488 nm solid state laser. Three angles of scattered light plus fluorescence are recorded, which allows multi-dimensional separation of platelets and red blood cells. The embedded algorithm defines retPLT in an FL1 versus 7° scatterplot (Figure 2C). This approach enables correcting for the size-dependent background fluorescence of platelets [39]. In healthy individuals, mean retPLT are between 1.4% and 2.2%, and reported reference ranges are 0.4%–2.8% [59], 0.4% to 4.45%–6.0% [60–62] and 1.0%–3.8% [63].

In the CELL-DYN Sapphire, the correlation between retPLT and MPV was investigated in relatively small groups and was described as not significant [64] or only weak [63]. However, in a large group of subjects with normal platelet counts, we found a significant negative correlation between retPLT and MPV [62].

Comparison of reticulated and immature platelet methods

Due to the lack of a standardized reference method for retPLT, assessing the performance of the automated methods is difficult; only a few side-by-side comparisons are available.

Studies where Sysmex XE-2100 IPF was compared with reference flow cytometric retPLT indicated low or moderate correlations [65, 66]. Remarkably, the coefficients of correlation differed between patients groups, from no correlation in healthy individuals to relatively high correlations in patients with thrombocytopenia due to peripheral destruction [66, 67]. One study using the Sysmex XT-2000iV, which uses an identical method as XE-2100, demonstrated reasonable correlation between IPF and reference flow cytometry, albeit with a significant systematic bias [68].

Direct comparison between Sysmex XE-2100 or XE-5000 IPF and CELL-DYN Sapphire retPLT resulted in weak or modest correlations [63, 64]. Sapphire was found to have a distinctively narrower reference range than XE-5000, enabling higher sensitivity for separating normal and abnormal patients [63].

These weak correlations and the different relationships between IPF/retPLT and MPV mentioned previously, lead to the conclusion that although both parameters harbor information of platelet turnover, they do reflect different aspects of thrombopoiesis. As a consequence, the two parameters cannot be used interchangeably [63].

Pre-analytical aspects

Obviously, immature or reticulated platelets are normally measured in blood samples collected into K2-EDTA, as for other hematologic parameters. One single study advised that citrate-theophylline-adenosine-dipyridamole solution (CTAD) was to be preferred over EDTA, as IPF measured using a Sysmex XE-5000 remained more stable in blood from patients with chronic ITP [69].

Regarding storage temperature before analysis, most authors agree that XE-2100 IPF is stable for 24–48 h after blood collection, provided the samples are kept at ambient temperature [49, 70, 71]. Others, however, reported stability for 3–8 h only [50, 51]. When blood samples are kept at 4 °C, IPF is rather unstable [69, 72], but the increase in IPF is apparently so predictable that it can be corrected for by a simple algorithm [73].

Stability data on retPLT as measured with CELL-DYN Sapphire range between <6 h [63] and at least 26 h [62].

Post-analytical aspects

Traditionally, IPF is expressed in relation to the PLT count, as a percentage. The absolute IPF (IPF#), the concentration of immature platelets (in 109/L), might better reflect real-time platelet production in analogy with what absolute reticulocyte count does for erythropoietic activity [67]. There are indeed some reports indicating the usefulness of IPF# in neonatal infections [74], in chronic liver disease [51] and in differentiating acute ITP and from thrombocytopenia due to acute leukemia [75]. It has been reported that IPF#, but not relative IPF predicts imminent platelet recovery in chemotherapy-induced thrombocytopenia in children [76]. Importantly, IPF# seems not to be influenced by platelet transfusions, whereas IPF% decreases, most likely due to dilution [77]. Other authors found IPF# not helpful for assessing platelet turnover [63]. More studies are needed in order to fully appreciate whether the absolute IPF or absolute retPLT count have additional value to relative counts.

It is well accepted that accelerated megakaryopoiesis is associated with increased MPV. As also IPF is often increased in this condition, this has led to the widespread belief that immature platelets are synonymous with large platelets [47, 78]. This notion seems to be reinforced by high correlations between IPF and MPV [64, 66, 79–81]. This correlation is only present in Sysmex analyzers, most likely as a result of how IPF is derived: the scatterplots indeed suggest that immature platelets are the largest platelets [49, 54, 82]. In contrast, no significant correlation was found between retPLT and MPV in flow cytometry [17] and neither in CELL-DYN Sapphire [63, 83]. It is true that platelets produced in response to stress megakaryopoiesis are on the average larger, but there is no evidence that this can be extrapolated to normal megakaryopoiesis. In healthy individuals with steady-state platelet production, we found a significant negative association between retPLT and MPV [62]. This is more in keeping with the concept of a constant circulating platelet mass, which would require more small platelets or fewer larger platelets [1]. So, reticulated platelets are not necessarily large platelets.

Clinical utility of reticulated/immature platelets

In the earlier years, retPLT research focused on a possible differential diagnostic aid in patients with thrombocytopenia. Since megakaryopoietic activity is low in patients with bone marrow failure, the assumption was that consequently retPLT would be low, too. In contrast, conditions with peripheral platelet destruction like ITP are characterized by accelerated megakaryopoiesis and hence the retPLT count would be increased. Many studies have now confirmed that retPLT or IPF are valuable in establishing the cause of thrombocytopenia: decreased production can reliably be distinguished from peripheral destruction (see Table 2).

Table 2

Clinical conditions where retPLT or IPF can be regarded as established diagnostic or prognostic tools.

Another well-documented application of retPLT is monitoring the thrombocytopenic phase after chemotherapy and transplantation for hematological malignancies (Table 2). Generally, an increase in retPLT precedes the recovery of platelet count by 2–3 days. This creates the opportunity to defer platelet transfusions that would be given when transfusion decisions are based on platelet counts only. Until present, limited clinical evidence supporting this concept has been reported and there is an evident need for randomized, controlled clinical studies in this field [82, 97, 98]. A highly interesting observation that also needs to be confirmed is that prophylactic transfusions with high IPF platelet concentrates seem to be more effective than low IPF platelets [99].

Researchers in the field of cardiovascular diseases have recently gained interest in retPLT (Table 3). The current literature seems to provide preliminary support for the clinical utility of retPLT or IPF determinations, for risk assessment in acute coronary syndrome [100–104] as well as for monitoring drug treatment of coronary artery disease [78, 79, 105–107]. However, larger and randomized studies are necessary for proving that these concepts are effective and safe in clinical practice. There are also indications that it is preferentially immature platelets that are recruited into arterial thrombi [106] and if confirmed, this finding may have consequences for treatment with anti-platelet drugs [107].

Table 3

Clinical conditions where retPLT or IPF may have potential clinical utility, but further studies are required for confirmation.

Apart from the above disease states, there are several other conditions where retPLT are supposed to play a role (Table 3). Most of these associations are only described in a single paper or are based on small patient numbers, thus the scientific evidence is still weak. Therefore, additional research studies are absolutely required for obtaining independent confirmation on the utility retPLT or IPF in these settings.

Highlights and conclusions

Summarizing the literature reviewed above, it can be concluded that:

  • Reticulated platelets are the most immature platelets in the circulation and they reflect the megakaryopoietic activity in bone marrow

  • Reticulated platelets are not necessarily large platelets

  • The two commercially available methods (Sysmex IPF and Abbott reticulated platelets) are not interchangeable and each require their own reference values and clinical decision limits

  • There is an urgent need for a standardized method that can serve as an international reference for assessing the performance of reticulated/immature platelet methods

  • Measuring reticulated/immature platelets is of proven clinically usefulness in patients with thrombocytopenia of unknown etiology and for monitoring bone marrow recovery after chemotherapy and stem cell transplantation

  • Reticulated platelets seem to play a role in the etiology of coronary arterial diseases. If confirmed in large-scale trials, measurement of reticulated platelets or IPF may be useful for risk assessment as well as therapy monitoring

  • Some other indications look promising, but need further investigation before routine use of reticulated platelet assays is warranted


I thank my colleagues Ting Yu and Nigel Llewellyn-Smith for their valuable comments and advice.

Conflict of interest statement

Author’s conflict of interest disclosure: The author stated that there are no conflicts of interest regarding the publication of this article. Employment and leadership played no role in the study design; in the collection, analysis, and interpretation of data; in the writing of the report; or in the decision to submit the report for publication.

Research funding: None declared.

Employment or leadership: The author is a scientific employee of Abbott Diagnostics.

Honorarium: None declared.


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About the article

Johannes J.M.L. Hoffmann

Johannes (Hans) Hoffmann started his career in clinical chemistry in 1976 as a trainee. Once certified as a specialist he became the head of the hematology laboratory in a large tertiary-care teaching hospital in the Netherlands, where he later was also appointed director of the Department of Clinical Laboratories. In 1992 he obtained his PhD in medical sciences at Leiden University, the Netherlands, on a thesis in the field of fibrinolysis. Since 2008 he is responsible for scientific affairs in hematology with Abbott Diagnostics in Europe. His scientific work comprises over 100 papers in peer-reviewed journals, mainly focused on general hematology, flow cytometry, coagulation and fibrinolysis. He is also author and co-author of several books on laboratory medicine and hematology. He gave numerous oral and poster presentations in congresses and other scientific events. He acts as a reviewer for various journals, including Clinical Chemistry and Laboratory Medicine, where he currently serves his last term as an Editorial Board member. He is a member of several international committees and working groups on standardization in laboratory hematology.

Corresponding author: Dr. Johannes J.M.L. Hoffmann, Abbott Diagnostics Division, Abbott GmbH and Co. KG, Max-Planck-Ring 2, 65205 Wiesbaden-Delkenheim, Germany, E-mail:

Received: 2014-02-16

Accepted: 2014-04-08

Published Online: 2014-05-07

Published in Print: 2014-08-01

Citation Information: Clinical Chemistry and Laboratory Medicine (CCLM), Volume 52, Issue 8, Pages 1107–1117, ISSN (Online) 1437-4331, ISSN (Print) 1434-6621, DOI: https://doi.org/10.1515/cclm-2014-0165.

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Sabrina Buoro, Barbara Manenti, Michela Seghezzi, Paola Dominoni, Mauro Vavassori, Rosangela Trezzi, Monica Galli, and Guido Finazzi
Scandinavian Journal of Clinical and Laboratory Investigation, 2017, Volume 77, Number 1, Page 73
C. Stratz, T. Nührenberg, M. Amann, M. Cederqvist, P. Kleiner, C. M. Valina, D. Trenk, F.-J. Neumann, and W. Hochholzer
Thrombosis and Haemostasis, 2016, Volume 116, Number 5, Page 941
Christian Stratz, Timo Bömicke, Iris Younas, Anja Kittel, Michael Amann, Christian M. Valina, Thomas Nührenberg, Dietmar Trenk, Franz-Josef Neumann, and Willibald Hochholzer
Journal of the American College of Cardiology, 2016, Volume 68, Number 3, Page 286
A. Pecoraro, A. Troia, M. Sacco, A. Maggio, R. Di Marzo, and M. Gioia
International Journal of Laboratory Hematology, 2016, Volume 38, Number 5, Page e89
Monica Verdoia, Patrizia Pergolini, Matteo Nardin, Roberta Rolla, Lucia Barbieri, Alon Schaffer, Paolo Marino, Giorgio Bellomo, Harry Suryapranata, and Giuseppe De Luca
Journal of Thrombosis and Thrombolysis, 2016, Volume 42, Number 2, Page 245
Bianca Rocca and Steen Husted
Drugs & Aging, 2016, Volume 33, Number 4, Page 233
Mikala Klok Joergensen and Lise Bathum
Scandinavian Journal of Clinical and Laboratory Investigation, 2016, Volume 76, Number 2, Page 172
Thomas F. Lüscher
European Heart Journal, 2015, Volume 36, Number 45, Page 3125
Monica Verdoia, Chiara Sartori, Patrizia Pergolini, Matteo Nardin, Roberta Rolla, Lucia Barbieri, Alon Schaffer, Paolo Marino, Giorgio Bellomo, Harry Suryapranata, and Giuseppe De Luca
Journal of Thrombosis and Thrombolysis, 2016, Volume 41, Number 4, Page 663
Isabell Bernlochner, Alexander Goedel, Conny Plischke, Stefanie Schüpke, Bernhard Haller, Christoph Schulz, Katharina Mayer, Tanja Morath, Siegmund Braun, Heribert Schunkert, Wolfgang Siess, Adnan Kastrati, and Karl-Ludwig Laugwitz
European Heart Journal, 2015, Volume 36, Number 45, Page 3202
Leonardo Lorente, María M. Martín, Ester López-Gallardo, José Blanquer, Jordi Solé-Violán, Lorenzo Labarta, César Díaz, Alejandro Jiménez, Julio Montoya, and Eduardo Ruiz-Pesini
Journal of Critical Care, 2015, Volume 30, Number 5, Page 935
Qin Wu, Jianan Ren, Dong Hu, Pengjun Jiang, Guanwei Li, Nadeem Anjum, Gefei Wang, Guosheng Gu, Jun Chen, Xiuwen Wu, Song Liu, Yuan Li, Yunzhao Zhao, and Jieshou Li
Medicine, 2015, Volume 94, Number 19, Page e814
Romana Mijovic, Nada Kovacevic, Marija Zarkov, Zoran Stosic, Velibor Cabarkapa, and Gorana Mitic
Journal of Thrombosis and Thrombolysis, 2015, Volume 40, Number 2, Page 203
Francesco Zaccardi, Bianca Rocca, Dario Pitocco, Luigi Tanese, Alessandro Rizzi, and Giovanni Ghirlanda
Diabetes/Metabolism Research and Reviews, 2015, Volume 31, Number 4, Page 402
Thomas Pierre Lecompte and Michael Pierre Bernimoulin
Clinics in Laboratory Medicine, 2015, Volume 35, Number 1, Page 209
Luci Maria SantAna Dusse and Letícia Gonçalves Freitas
Clinica Chimica Acta, 2015, Volume 439, Page 143

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