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Use of a Cytokine-Release Assay to Demonstrate Loss of Platelet Secretory Capacity During Blood Bank Processing and Storage.

A major challenge in transfusion medicine is the development of techniques for the preparation and storage of blood components that allows them to retain as much functionality as possible. Platelets (PLTs) are particularly problematic, because they are labile and short lived even in vivo. (1,2) Efforts to date at optimizing PLT preparation and storage have largely focused on in vitro measures such as PLT count and pH. Preservation of PLT function, in particular secretory capacity, has been largely overlooked. Secretion, nevertheless, is a key component of overall PLT function. For example, PLT alpha granules, whose contents are released into the vascular space following PLT activation, are rich in many important mediators of hemostatic and nonhemostatic processes, such as procoagulants factor V and von Willebrand factor, mitogens like PLT-derived growth factor, other cytokines such as RANTES (chemokine ligand 5; regulated on activation, normal T cell expressed and secreted), immunoglobulins, and antimicrobial proteins. (1,3) Platelet secretion is important in the setting of an acute injury, because the discharged alpha-granule contents mediate the recruitment of leukocytes and PLTs, promote clotting, provide immunity against infection, and contribute to wound healing. (3-5)

During storage, PLTs undergo significant deleterious changes in morphology and function as part of a process known as the PLT storage lesion. (6,7) Optimization of storage conditions to overcome this lesion requires sensitive assays that accurately reflect aspects of PLT function that one wishes to preserve. Storage optimization studies to date have relied primarily on relatively basic tests of PLT quality. As noted in the Food and Drug Administration8 Guidance for Industry for Platelet Testing and Evaluation of Platelet Substitute Products, many of the current tests have limitations and few actually measure PLT function. Biochemical tests like pH, Po2, Pco2, lactate dehydrogenase, glucose, and adenosine triphosphate "do not correlate well with PLT performance in vivo." 8 In fact, many of these tests reflect the storage milieu rather than directly measuring PLT functional capabilities. Platelet morphology and surface markers like P-selectin are measures of activation, but again do not measure function.

Because the goal of blood banks and transfusion services should be to devise storage conditions that maximally preserve the functionality of stored PLTs for transfusion, in vitro assays of PLT quality should actually measure important aspects of PLT function. One existing functional assay is the measurement of PLT aggregation in response to stimulants such as adenosine diphosphate (ADP), collagen, and epinephrine. (9,10) However, few studies have examined the integrity of other cellular functions of PLT preparations, such as secretory capacity and the granule release reaction. (11-13) Because maintenance of PLT secretory capacity may be important, as noted above, particularly for immunosuppressed or anticoagulated patients, it is potentially useful to examine the impact of current blood bank preparation and storage conditions on this aspect of PLT function.

To address this issue, we report the use of a cytokine-release assay to measure the secretory capacity of stored PLTs. This assay measures the in vitro release of the alpha-granule cytokine RANTES in response to the agonist ADP. RANTES was chosen because it is present in PLT alpha granules in relatively high concentration, and therefore should yield a sensitive assessment of the capability of PLTs to degranulate in response to agonist stimulation. (14,15) We believe that the retention of this alpha-granule secretory capacity should better reflect overall PLT function than many of the current indirect measures of PLT integrity. As such, a cytokine-release assay might be used to design improved methods of PLT preparation and storage.


Preparation of Fresh Whole Blood and Mini-Preparations of PLT-Rich Plasma and PLT Concentrate Using a Small-Scale Adaptation of Blood Bank Techniques

This study was approved by the human investigations committee of the hospital institutional review board. About 20 mL of whole blood (WB) was drawn into 3.2% citrate from volunteer donors (N = 4). An aliquot of WB (approximately 5 mL) was saved for later analysis. The remaining WB was subsequently centrifuged at 2000g for 3 minutes at 22[degrees]C to yield PLT-rich plasma (PRP). About 5 mL of PRP was set aside for analysis. The remaining PRP was centrifuged at 5000g for 5 minutes at 22[degrees]C to yield a PLT pellet and supernatant plasma. This pellet was allowed to rest (ie, left undisturbed) for 60 minutes. Following this rest period, the PLT pellet was resuspended in the entire volume of the supernatant on a rotating 3-rpm PLT agitator (Platelet Mixer Model 348, Fisher Scientific, Waltham, Massachusetts). (16) The resulting "PLT concentrate" (PC) was not a concentrate per se, because it had the same volume as PRP; however, it was otherwise subjected to standard PC preparation steps.

Studies on Blood Center PCs

Leukocyte-reduced, WB-derived PCs, collected into tri-2ethylhexyl trimellitate plastic bags and prepared according to American Association of Blood Banks and Food and Drug Administration specifications, (16) were obtained from the clinical inventory of a regional blood center on day 2 of storage. These blood center PCs were subsequently stored with continuous rotational agitation at 3 rpm, as described above, in an incubator maintained at 20[degrees]C to 24[degrees]C. The PCs were sampled using sterile techniques at various storage time points for analysis (see below). Prior to analysis, the PLT counts of the aliquots were adjusted to approximately 250 000/[micro]L using PLT-poor plasma prepared from the same PLT bag from which the aliquot was obtained. Platelet poor plasma was the supernatant that resulted from centrifuging a small volume of the PC at 5000g for 7 minutes.

PLT Activation

Unless otherwise noted, PLTs were activated by pipetting 50 [micro]L of concentrated agonist solution into 950 [micro]L of the relevant PLT preparation: WB, PRP, or PC. The final concentration of agonists in the reaction mixture was 20 [micro]M ADP, 0.19 ng/mL collagen, and/or 4 X [10.sup.-5] M epinephrine (Bio/Data Corp, Horsham, Pennsylvania). Lyophilized agonists were reconstituted with 0.9% normal saline prior to their addition to the PLT preparations. Controls were run in parallel for each activation experiment, in which 0.9% normal saline was substituted for the agonist solution.

Cytokine Release Measurement

After addition of either PLT agonist or normal saline as noted above to aliquots of WB, PRP, or PC, the specimen was gently agitated by hand to ensure even distribution of agonist. The specimens were then centrifuged at 10 000g for 10 minutes at 4[degrees]C. The supernatant was pipetted off the PLT pellet and frozen at approximately -20[degrees]C. For RANTES analysis by enzyme immunoassay (R&D Systems, Minneapolis, Minnesota), thawed supernatant was diluted at 1:200 using kit assay buffer. Results represent means of duplicate determinations on each sample tested. For the calculation of molecules of RANTES released per PLT, we used a RANTES molecular weight of 10 075 Da. (17)

PLT Aggregation by Single-PLT Counting

Baseline PLT counts were obtained by an impedance method (Beckman Coulter, Brea, California) on PLT aliquots prior to activation. Following addition of the agonist, the aliquot was gently agitated by hand; then, the PLT count was immediately repeated. The percentage aggregation was calculated from the reduction in countable single PLTs as follows (18):

% aggregation = [(baseline PLT count - post-agonist PLT count) / baseline PLT count] X 100%

Statistical Analysis

For comparison of results obtained before and after agonist stimulation of PLT preparations, the 1-sample sign test was performed. (19) Other data comparisons were done with the Kruskal-Wallis test; pair-wise comparisons were made with the Mann-Whitney U test when the Kruskal-Wallis test indicated significant differences. (20) P values less than .05 were considered significant. Calculations were performed using computer software (Excel 2010, Microsoft Corp, Redmond, Washington).


A cytokine-release assay was developed in order to test the secretory capacity of PLTs in vitro in response to PLT agonists. We evaluated several known PLT agonists (ie, ADP, collagen, and epinephrine) for their ability to stimulate RANTES release from unfractionated PLTs in fresh WB (Figure 1). These 3 agonists stimulated RANTES secretion about 4- to 5-fold above baseline levels when added to PLTs at concentrations recommended for PLT aggregation studies. (18,21,22) In addition, we examined the effects of combinations of 2 of the 3 agonists on RANTES release and observed that the effect was approximately additive (Figure 1). This indicated that the RANTES release obtained with single agonists at the concentrations used was not maximal.

In subsequent experiments, 20 [micro]M ADP was used to examine secretory capacity and aggregation in WB and mini-preparations of PRP and PC. Mean PLT counts showed a reduction of about 30% in processing from WB to PC, although this change was not statistically significant (Figure 2, A; P = .06; Kruskal-Wallis test). Figure 2, B, shows the comparison in RANTES release after each preparation step. ADP stimulated release of RANTES from PLTs in fresh WB on average by 4.1-fold (19.9 [+ or -] 0.62 ng/mL versus 4.84 [+ or -] 0.51 ng/mL [control]; P < .001; 1-sample sign test; N = 4) and in fresh PRP by an average of 4.7-fold (26.0 [+ or -] 5.49 ng/mL versus 5.50 [+ or -] 1.37 ng/mL [control]; P = .002; 1-sample sign test; N = 4). Following an additional centrifugation step, freshly prepared PC still demonstrated RANTES release in response to ADP (40.1 [+ or -] 18.4 ng/mL versus 30.3 [+ or -] 19.6 ng/mL [control]; P < .001; 1-sample sign test; N = 4), although to an apparently lesser degree (1.3-fold over control) than WB or PRP because of an increase in the baseline/control. No difference was found in the amount of mean RANTES secreted per activated PLT in processing from WB (63.1 attograms [ie, 3772 molecules] RANTES secreted/activated PLT) to PC (62.5 attograms [ie, 3736 molecules] RANTES secreted/activated PLT; P = .14; Kruskal-Wallis test).

Comparisons of baseline, nonstimulated RANTES levels showed significant changes during processing from fresh WB to PRP to PC (P = .02; Kruskal-Wallis test). Pair-wise comparisons showed a 6.3-fold higher level of baseline plasma/supernatant RANTES levels in PC compared with WB (P = .02; Mann-Whitney U test) and a 5.5-fold higher level in PC compared with PRP (P = .02; Mann-Whitney U test). No difference was found between WB and PRP (P = .56; Mann-Whitney U test). As shown in Figure 2, C, the amount of RANTES secreted following activation with ADP and the baseline level of RANTES changed in a reciprocal fashion.

Platelet aggregation studies by single-PLT counting (SPC) were run in parallel for each step of PLT processing (N = 3; Figure 2, D). The results revealed that 20 [micro]M ADP stimulated 95.6% [+ or -] 1.70% aggregation in fresh WB, 82.5% [+ or -] 7.00% aggregation in PRP, and 60.5% [+ or -] 8.60% aggregation in PC. Overall aggregability was decreased in processing from WB to PC (P = .04; Kruskal-Wallis test). However, pair-wise comparisons showed changes of marginal significance in aggregation between WB and PRP (P = .05; Mann-Whitney U test) and between WB and PC (P = .05; Mann-Whitney U test); no significant difference was found between PRP and PC aggregability (P = .13; Mann-Whitney U test).

Cytokine-release experiments were subsequently performed on PCs (N = 10 units) obtained from a regional blood center on day 2 of storage. As shown in Figure 3, the mean PLT count of stored PCs showed a slight downward trend of about 16% from day 2 to day 7 of storage, although this change was not significant (P = .23; Kruskal-Wallis test). Figure 3 also shows the adjusted PLT counts in aliquots used for subsequent analysis. ADP failed to stimulate RANTES release at all storage times examined (P = .31 for day 2; P = .53 for day 5; P = .35 for day 7; 1-sample sign test; Figure 4, A). A progressive and significant increase in baseline (ie, nonstimulated) levels of RANTES was observed in PCs during storage (P < .001; Kruskal-Wallis test). It should be noted that RANTES levels were substantially higher in blood bank PCs compared with the mini-preparations from Figure 2, B. This was expected because of the higher unadjusted PLT counts of the blood center PCs and the fact that the PLTs had been stored for 2 additional days. Pair-wise comparisons showed significant differences for baseline RANTES levels on day 2 versus day 5 (P < .001; Mann Whitney U test) and day 2 versus day 7 (P = .002; Mann-Whitney U test). No significant change was found between day 5 and day 7 (P = .11; Mann-Whitney U test).

For the same stored blood center PCs, PLT aggregation in response to ADP showed progressive decreases over time: 41.0% [+ or -] 15.8% on day 2, 10.5% [+ or -] 8.10% on day 5, and 2.30% [+ or -] 2.70% on day 7 (P < .001; Kruskal-Wallis test; Figure 4, B). Pair-wise comparisons were significant for all time points (day 2 versus day 5, P < .001; day 2 versus day 7, P < .001; day 5 versus day 7, P = .01; Mann-Whitney U test). The decrease in PLT aggregability was noted to occur more slowly than the decrease in agonist-stimulated RANTES release; significant PLT aggregability was still demonstrable on day 2, whereas cytokine release was completely lost.


Traditional endpoints for measuring stored PLT quality mostly reflect changes in the PLT milieu rather than PLT function. Therefore, we sought to develop an assay that directly measures an important but often overlooked aspect of PLT quality, that is, PLT alpha-granule secretory capacity. Because secretion is dependent on complex extracellular and intracellular pathways, measurement of secretory capacity may be a better assessment of the totality of PLT function during preparation and storage. We assessed secretory capacity with a cytokine-release assay, which consisted of measuring the ability of ADP to induce RANTES release from PLTs in vitro. In addition to being a measure of PLT alpha degranulation, RANTES is significant in its own right and may be important in overall PLT function. Among a variety of biological activities, RANTES promotes leukocyte chemotaxis, activates certain natural killer cells, enhances monocyte adherence to endothelial cells, and appears to have microbicidal activity. (2-5,23-27)

Our study revealed several findings indicating that certain steps in PC preparation and storage are deleterious to PLT secretory capacity. Notably, the "hard spin" used to pellet PLTs in making PC appeared to be the single most damaging step in PLT processing. This was demonstrated by the reduced release reaction in PLTs in PC compared with those in WB or PRP. Our results indicated that this reduction was likely due to a smaller number of PLTs responding to the agonist rather than less RANTES release per PLT, because the amount of RANTES secreted per PLT was not different between WB and PC. Loss of secretory capacity progressed further during early PLT storage, with complete elimination of the release reaction by day 2.

We observed that RANTES levels increased in the plasma portion of PCs during blood bank storage in the absence of agonist stimulation, as has been previously reported. (14,15,28) A finding that has not been previously reported was the cytokine release observed during the initial processing of PC from WB in the absence of agonist stimulation. The processing step that caused the greatest increase in baseline plasma/supernatant RANTES was the hard spin used to make PC from PRP. This increase in baseline cytokine levels is either a consequence of inadvertent PLT activation during processing and storage (eg, during centrifugation or following exposure to artificial surfaces), or PLT damage that results in cytokine leakage. Additional studies are necessary to distinguish between these possibilities.

We found that during the early stages of PC processing an inverse relationship existed between plasma/supernatant RANTES levels and the amount of RANTES actively secreted following ADP stimulation. However, this reciprocal relationship was no longer evident at day 2 of storage or later. The reciprocal relationship was no longer observed at later time points because plasma/supernatant levels of RANTES continued to increase after responsiveness to ADP was completely lost. For this reason, measurement of plasma/supernatant RANTES levels is not a reliable surrogate marker for residual secretory function during Pc storage, particularly at later storage points.

Few previous studies have addressed PLT secretory capacity. Scott et al (11) examined ADP-induced release of beta-thromboglobulin, another alpha-granule component, at day 1 of storage. However, they did not examine processing steps or later storage times. In addition, Wenzel and colleagues (12,13) examined the release of soluble CD40 ligand (sCD40L) in stored PLTs, but also did not investigate PC processing steps. Their studies showed a progressive reduction over time, but not a complete loss, of the ability of sCD40L to be released from PLTs in response to calcium-induced clot formation. Because the authors did not use direct agonist stimulation, it is unclear whether their approach measured secretory capacity or cytokine leakage from damaged PLTs trapped in a clot. As such, our work clarifies and extends these earlier studies.

Efforts to better preserve alpha-granule secretory capacity of stored PLTs may be warranted. Platelet alpha-granule contents play wide-ranging and important roles at the site of acute injury, such as provision of coagulation factors, promotion of leukocyte chemotaxis and wound healing, enhancement of alloimmune responses, and suppression of bacterial growth. (3-5,24-27) However, the role of alpha-granule contents and their contribution to clinical outcomes have not been sufficiently studied in the transfusion setting; further research is needed.

In addition to examining cytokine release, we assessed PLT aggregation by the technique of SPC. We found that aggregability decreased in parallel to the loss of secretory capacity, but did so more slowly. Although cytokine release capacity was essentially eliminated by day 2 of storage, PLTs demonstrated aggregation, although substantially reduced, through day 7 in response to agonist stimulation. Therefore, PLT aggregability is not a marker for all PLT functions. Aggregation and cytokine secretion also differed in that 20 [micro]M ADP gave near-complete aggregation in fresh WB, but, at best, 50% maximal RANTES release (in comparison with the response obtained with 2 agonists).

To our knowledge, SPC has not been previously used to assess aggregation in stored PLTs. However, the successful use of SPC has been reported in several other in vitro settings, such as the assessment of inherited PLT disorders, the monitoring of anti-PLT agents, and the measurement of PLT function in thrombocytopenic patients. (18,22,28-30) Moreover, SPC results correlate well with more traditional PLT function platforms such as light transmission aggregometry. (18,22,29) Single-PLT counting offers two advantages over other PLT function tests: (1) it can be performed with basic hematology cell counters (18) and (2) it does not require the reconstitution of WB to measure aggregation, unlike some other PLT function assays. (9,10)

The possibilities for extending the shelf life of PLT products have been examined. (31) However, our findings indicate that a substantial reduction in PLT secretory capacity and aggregability have already occurred within 5 days of storage. As such, the benefits of further extending the shelf life of current PLT products are questionable. Any efforts to extend PLT shelf life should be combined with the better preservation of PLT function. Functional assays, such as cytokine release and PLT aggregation, might be useful endpoints in such efforts.

It is important to note that our studies of fresh, day 0 WB, PRP, and PC were performed using small volumes of nonleukoreduced blood in test tubes rather than leukoreduced blood bank products in storage bags, as were used for later storage experiments. It is unclear whether, or to what extent, the effects of centrifugation on PLT function are dependent on the volume, shape, and composition of the PLT storage container. Nevertheless, the results were consistent with the intuitive expectation that activation might occur with a higher-speed rather than with a lower-speed centrifugation step. With regard to leukoreduction, the differences observed in RANTES release between fresh and stored products are unlikely to be due to passenger leukocytes in the freshly prepared, nonleukoreduced specimens. Leukoreduction has not previously been shown to affect the accumulation of RANTES in the plasma portion of stored PLT preparations. (32-34) This indicates that leukocytes are not a significant source of RANTES in PLT products. It should also be emphasized that our study examined only WB-derived PCs stored with rotational agitation. It is possible that different PLT preparations (eg, apheresis or buffy coat PLTs) or modes of agitation (eg, flatbed) could have affected the results. (35)

We also wish to point out that our studies focused entirely on alpha-granule secretory capacity and may not have any implications for dense granule release. We do not know what degree of correlation there may be between alpha and dense degranulation. Further studies in this regard may be warranted. Moreover, RANTES secretion is not necessarily representative of alpha degranulation overall. It is possible that inducible alpha-granule secretion might have been obtained in stored PCs at day 2 or later if we had examined other alpha-granule contents or tried higher concentrations of ADP or multiple agonists. Nevertheless, we clearly saw a difference in responsiveness in freshly prepared PLTs as compared with stored PLTs. Therefore, at a minimum, our data indicate that one aspect of alpha-granule secretory capacity is compromised as a result of PLT storage.

In summary, we have developed a cytokine-release assay for the evaluation of blood bank-stored PLTs and demonstrated a substantial reduction in PLT secretory capacity during PLT preparation leading to complete loss by day 2 of storage. Secretory capacity deteriorated more rapidly than PLT aggregability, which was not eliminated until day 5. We believe that the cytokine-release assay could serve as a useful endpoint in the development of improved methods of PLT storage that optimize PLT function.

This project was supported by a College of American Pathologists Foundation John H. Rippey Grant for Laboratory Quality Assurance. This material is the result of work supported with resources and the use of facilities at the VA Connecticut Healthcare System, West Haven, Connecticut.

Please Note: Illustration(s) are not available due to copyright restrictions.


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Christopher A. Tormey MD; Gary Stack, MD, PhD

Accepted for publication January 28, 2014.

From Pathology and Laboratory Medicine Service, VA Connecticut Healthcare System, West Haven; and the Department of Laboratory Medicine, Yale University School of Medicine, New Haven, Connecticut.

The authors have no relevant financial interest in the products or companies described in this article.

The views expressed in this article are those of the authors and do not necessarily reflect the position or policy of the Department of Veterans Affairs or the United States government.

Reprints: Christopher A. Tormey, MD, Department of Laboratory Medicine, Yale School of Medicine, 330 Cedar St, PO Box 208035, New Haven, CT 06520 (e-mail:

Caption: Figure 1. RANTES (chemokine ligand 5; regulated on activation, normal T-cell expressed and secreted) release from unfractionated platelets in fresh whole blood after the addition of agonists, singly or in combination. Following the addition of 100 [micro]L of agonist or control solution (0.9% saline; Con), the final concentration of agonists in the reaction mixture was 20 [micro]M for adenosine diphosphate (ADP), 0.19 ng/mL for collagen (Coll), and/or 4 X [10.sup.-5] M for epinephrine (Epi).

Caption: Figure 2. Cytokine release and platelet (PLT) aggregation during PLT processing steps on day 0. A, Bar heights represent mean PLT counts associated with each PLT preparation step (N = 4). Platelet counts trended downward with processing from whole blood (WB) to platelet concentrate (PC), although these changes were not significant (P =.06). Error bars represent standard deviation. B, RANTES (chemokine ligand 5; regulated on activation, normal T-cell expressed and secreted) levels in the plasma/supernatant of fresh WB, platelet-rich plasma (PRP), and PC before and after the addition of 20 [micro]M adenosine diphosphate (ADP) or 0.9% normal saline (N =4). Gray bars represent RANTES levels in nonstimulated controls to which normal saline was added (Control), and black bars represent RANTES levels in identical preparations to which ADP was added (Post-ADP). Bar heights represent mean RANTES from 4 independent preparations. Significant differences were found between controls and ADP-stimulated samples for WB (P < .001), PRP (P =.002), and PC (P < .001); the asterisks indicate comparisons that were significantly different. Error bars represent standard deviation. C, The dashed line with closed circles (-*-) represents the ratio of ADP-stimulated RANTES levels to nonstimulated RANTES levels (control) for the same preparation step from Figure 2b; the higher the ratio, the greater was the magnitude of stimulated RANTES release observed. The solid line with closed squares (-?-) represents the ratio of baseline, nonstimulated RANTES levels in PRP and PC to baseline, nonstimulated RANTES levels in WB; the higher the ratio, the greater was the magnitude of spontaneous RANTES release associated with each processing step. D, Percentage PLT aggregation in fresh WB, PRP, and PC after the addition of 20 [micro]M ADP. Bar heights represent mean percentage aggregation from 3 independent preparations. As indicated by the asterisk, aggregation in PC was significantly less than that in WB (P = .04); other pair-wise comparisons revealed no significant differences (P >.05 in all other cases). Error bars represent standard deviation. Note that aggregation was not measured in 1 of the 4 specimens that had been tested for RANTES release.

Caption: Figure 3. Platelet (PLT) counts in stored platelet concentrates (PCs) from a blood donor center and in aliquots used for subsequent testing. The mean PLT count of stored PCs from a blood donor center (gray bars; Original bag) did not change significantly from day 2 to day 7 (P =.23). PLT counts in aliquots from donor center PCs used in subsequent assays (see Figure 4) were adjusted to an approximate PLT count of 250 000/[micro]L (black bars; Assayed aliquot) by diluting the specimen with PLT-poor plasma from the same PC unit (N = 10). Error bars represent standard deviation.

Caption: Figure 4. Cytokine release and platelet (PLT) aggregation in stored platelet concentrates (PCs) from a blood donor center. A, Bar heights represent mean RANTES (chemokine ligand 5; regulated on activation, normal T-cell expressed and secreted) levels in samples of whole blood (WB)-derived PCs (N = 10) obtained from a blood donor center before and after the addition of 20 [micro]M adenosine diphosphate (ADP; black bars; Post-ADP) or 0.9% normal saline (gray bars; Control). No significant differences were noted between ADP-stimulated and control samples on day 2 (P=.31), 5 (P = .53), or 7 (P =.35) of storage. Error bars represent standard deviation. B, Mean percentage PLT aggregation in PCs after the addition of 20 [micro]M ADP to aliquots on days 2, 5, and 7 of storage. As indicated by the asterisks, PLT aggregation showed a progressive, significant decrease over storage time (P <.001). All pair-wise comparisons (ie, day 2 versus day 5; day 2 versus day 7; day 5 versus day 7) showed significant differences (P [less than or equal to] .01 in all cases). Error bars represent standard deviation.
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Author:Tormey, Christopher A.; Stack, Gary
Publication:Archives of Pathology & Laboratory Medicine
Date:Nov 1, 2014
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