Reticulated Platelets: Definition and Measurements
Definition and Functional Properties
Heterogeneity of platelet size and function has been recognized since the 1930s. Serious
attention turned to differential platelet characteristics following the studies of
Karpatkin.[8] The author separated platelets in the human circulation according to the size and
weight and described ‘large heavy’ and ‘small light’ platelet populations. Similar
platelet populations were then identified in rabbits; survival studies of radiolabelled
platelets suggested that the large heavy platelets represented a younger population
than the small light platelets. Metabolic studies pointed towards increased thrombotic
activity in large heavy platelets. Adenosine diphosphate (ADP) and platelet factor
4 release following stimulation with ADP, thrombin or epinephrine were enhanced several
fold in this group, compared with small light platelets.[9] Subsequently, Blajchman et al studied newly released platelets in rabbits that had
undergone bone marrow irradiation. Platelets harvested before the nadir platelet count
occurred were considered to be old platelets, while those harvested during recovery
of the counts were considered to be newly released. Newly released platelets were
about 50% larger, had a measured survival time that was about threefold greater and
contained greater amounts of membrane glycoproteins (GPs) than the older platelets.
Bleeding times in the rabbits were also shorter when the platelet population consisted
of younger platelets.[10] Since the late 1960s, a preponderance of evidence has indicated that larger platelets
are more likely to be younger than smaller platelets, contain higher concentrations
of thrombotic mediators and are more likely to participate in thrombosis. However,
the literature does not universally support specifically identifying larger platelets
as younger platelets, as some investigators have reported that quantitative differences
between platelet populations (i.e. increasing size imparting increasing quantities
of functional GPs) rather than qualitative differences (i.e. age alone) are responsible
for functional differences.[11]
[12]
The term ‘RPs’ was first used to describe newly synthesized platelets by Ingram and
Coopersmith. These investigators subjected dogs to acute blood loss and then sampled
circulating platelets 5 to 7 days later. Using light microscopic techniques, they
stained platelets with methylene blue, which was known to bind to ribonucleic acid
(RNA) and had previously been used as a supravital stain to identify nucleated red
blood cells that indicated accelerated erythropoiesis. They then studied platelets
before and after phlebotomy. During the recovery phase after phlebotomy, reticular
patterns were up to fivefold more common than prior to bleeding. Reasoning that these
same patterns were analogous to those seen in juvenile red blood cells, they proposed
that the RPs were recently released from the bone marrow.[13] More sophisticated identification of newly released platelets began in the early
1990s. Reticulated red blood cells were identified with flow cytometry using the fluorophore
thiazole orange (TO), which binds to nucleic acids with particular affinity for RNA
and fluoresces at 488 nm, increasing the fluorescence of RNA approximately 3,000-fold.[14] Interestingly, ‘platelet contamination’ was originally listed among the sources
of erroneous reticulocyte determinations. In 1990, Kienast and Schmitz reported this
technique after TO staining in thrombocytopenic patients who were found to have normal
or increased numbers of megakaryocytes in the bone marrow. TO-positive staining was
present in 26.9% of platelets in this population compared with 8.6% of platelets in
control subjects. In contrast, patients with impaired platelet production did not
have increased TO staining.[15] The implication was that TO-labelled platelets indicated increased platelet production
in the bone marrow. This hypothesis was later confirmed in biotinylation experiments.
Biotin binds covalently to free amino acids in the cell membrane of the circulating
cells, but not to the membranes of cells sequestered in the bone marrow. Biotin injection
can therefore be used to label cells that are already present in the circulation and
this distinguishes them from cells that enter the circulation after the injection
has been performed. In a series of experiments, circulating (biotin-positive but TO-negative)
platelets were distinguished from newly synthesized (biotin-negative but TO-positive
platelets), establishing that the nucleic acid-rich platelets were less than 24 hours
old.[16]
[17] Accordingly, RP counts have been used clinically to distinguish between thrombocytopaenia
due to platelet consumption from that to decreased platelet production,[18] and to monitor response to exogenous thrombocytopaenia.[19] It should be noted, though, that TO also stains dense granule contents. When platelets
are incubated with high concentrations of TO or when the duration of incubation is
prolonged, the finding of TO-positivity loses specificity and therefore platelets
categorized as being ‘reticulated’ may not necessarily be newly synthesized.[20]
Much of our understanding of RPs is derived from the observation of platelet size
heterogeneity with the assumption that the pool of larger platelets consists predominantly
of RPs. Under conditions of stimulated thrombopoiesis, the bone marrow tends to produce
larger platelets.[11] As platelets increase in size, they tend to produce more rapid and complete aggregation,[21] contain higher concentration of adenosine triphosphate, adenosine monophosphate
and glycogen,[9] have greater lactate dehydrogenase activity and serotonin uptake and release,[11] produce more thromboxane B2[9]
[12]
[21] and express more GPIb and integrin αIIbβ3 GPIIb/IIIa.[22] These properties make larger platelets functionally more active.[11] Several studies have shown a significant correlation between mean platelet volume
(MPV) and RP in patients with stable ischaemic heart disease (SIHD) and ACS.[23]
[24]
[25]
[26] Lakkis et al showed that in a group of patients with ACS, compared to healthy controls,
MPV increased across the spectrum of ACS and was highest in patients with ST-elevation
myocardial infarction (STEMI). This was accompanied with a significant increase in
RPs.[24] The findings were later confirmed in a group of healthy volunteers stratified into
tertiles according to RP counts. Individuals in the upper tertile had the highest
MPV value.[26]
Until recently, the evidence for increased metabolic activity among RPs has been by
association rather than by direct observation. Recently, important observations provided
direct evidence that these findings in larger platelets, in fact, reflected the physiology
of RPs which appear to play an extensive role in all the steps involved in thrombus
formation. In 2002, Saving et al reported that RPs have higher expression of two important
adhesion receptors GP-IV and α6β1 (VLA-6).[27] GP-IV binds to thrombospondin, the major alpha-granule protein of platelets which
in turns facilitates their adherence to the vascular endothelium. On the other hand,
the VLA-6 facilitates the adhesion of activated platelets to laminin, one of three
major components of sub-endothelial matrix (along with fibronectin and collagen).
RPs were later found to have higher expression of P-selectin and procaspase activating
compound-1 (an epitope on activated integrin αIIbβ3 ),[26]
[28] as well as GPIbα, the receptor for von Willebrand factor.[29] Additionally, other components of thrombotic signalling system cyclooxygenase-1
and -2 (COX-1 and COX-2) were shown to be up-regulated in RPs.[26]
[30] While the contribution of COX-2 to the production of prostanoids may be minimal
in normal individuals, this effect appears to be accentuated in patients with increased
platelet turnover. In fact, NS-398, a selective COX-2 selective inhibitor, has a profound
inhibitory effect on prostanoid synthesis (PGE2 and TxB2) in patients with immune
thrombocytopaenia or peripheral blood stem cell transplantation.[31] Lador et al also examined the ability of RPs to respond to activation. Using flow
cytometric triple staining, they identified RPs using TO, and studied expression of
P-selectin and annexin V. After stimulation with ADP (10 μM), the proportion of RPs
expressing P-selectin and annexin V rose fourfold and 1.5-fold, respectively, compared
with mature platelets, implying greater reactivity among RPs.[32] Apart from their role in the formation of the platelet plug, RPs also seem to play
an important role in the coagulation cascade through enhanced participation in pro-thrombinase
complex assembly as indicated by the higher expression of both surface-bound α-granule
factor V and factor X upon stimulation with thrombin.[29]
[33]
The mechanism underlying this enhanced ability may be related to the ability of messenger
RNA (mRNA) present within RPs to undergo translation, resulting in protein synthesis.
An early observation by Kieffer et al indicated that rough endoplasmic reticulum was
more prominent in platelets taken from patients with idiopathic thrombocytopenic purpura
than among platelets from healthy volunteers. These platelets were also able to incorporate
radiolabelled methionine and leucine into their GP pool, implying that the immature
platelet population had increased capacity for protein synthesis.[34] The literature is now replete with evidence that despite being anucleate, platelets
have a vestigial pool of mRNA that has the ability to be translated into proteins,[35] although the functional significance of this RNA is still debated. Angénieux et
al correlated the mRNA content in murine platelets with protein synthesis. Using diphtheria
toxin, they induced thrombocytopaenia in mice which was followed by thrombocytosis.
As expected, the proportion of circulating RPs increased dramatically as the platelet
count rose. This finding was accompanied by the synthesis of proteins containing radiolabelled
methionine and cysteine. Over a period of hours, RNA content decreased and, in parallel,
protein synthesis decreased. This was accompanied by loss of about half of their ribosomal
and beta actin RNA after 6 hours and > 90% after 24 hours.[36] Although these findings imply that the RNA observed in RPs is functional, the short
duration of elevated RNA content raises important questions concerning the degree
to which increased protein synthesis provides functional support of thrombosis.[36]
At least two experiments have so far confirmed that RPs are indeed more active in
thrombosis compared to mature platelets. In the first experiment, plasma-rich thrombi
were generated by perfusion of the whole blood from normal individuals over porcine
carotid arterial segments under conditions of shear stress (shear rate: 3,350 s–1) similar to those seen in significant coronary artery stenoses. The thrombi were
then harvested by vortexing, and were then disaggregated and assessed with flow cytometry.
The intensity of TO staining was greater in platelets harvested from thrombi compared
to whole blood platelets. This finding was accompanied by a higher expression of the
integrin β3 chain in the platelets harvested from thrombi.[37] In the second experiment, plasma-rich platelets (PRPs) from healthy volunteers was
stimulated with arachidonic acid (AA) or ADP and the relative composition of the non-aggregated
platelet population was assessed. The investigators found that RPs contributed disproportionally
to the composition of the thrombus. Furthermore, confocal microscopic examination
showed that RPs are located in the core of aggregates.[38] These data suggest strongly that RPs play an essential role in platelet aggregation
by forming a nidus which facilitates recruitment of older platelets.[38] Whether this observation remains valid in vivo, where other blood elements participate
in thrombus formation, is unknown.
Measurement of Reticulated Platelets
Flow Cytometry
The standard for detecting the presence of RP is flow cytometry using a fluorescent
dye, most commonly TO, which binds to nucleic acids.[15] The results are generally expressed as the percentage of RP (RP%) in the total platelet
pool. This method has been criticized for its lack of standardization and for multiple
technical issues such as the non-specific binding of TO, varying types and concentrations
of fluorescent dye, varying incubation times and different gating and threshold settings
used in analysis.[39] Consequently, RP analysis using flow cytometry is time-consuming, operator-dependent
and subject to a large degree of variability in the reporting of a reference range,
which has varied between 1 and 15%.[40]
[41]
Immature Platelet Analysis
Although the terms ‘RP’ and “immature platelets” are used interchangeably in clinical
practice, it is important to distinguish between the two, as significant but incomplete
overlap exists in the populations measured. Sysmex (Kobe, Japan) developed a novel
method to measure RP in blood samples using automated analysers (XE-2100, XE-5000
and XN). The technique is based on a fluorescent dye, most commonly a mixture of polymethine
and oxazine, which penetrates cell membranes and stains platelet RNA. The stained
platelets are then passed through a semiconductor diode laser to measure the resulting
forward scattered light (cell volume) and fluorescence intensity (RNA content). A
computerized algorithm then separates the mature platelets (represented as blue dots)
from immature platelets (represented as green dots). RPs separated by this method
are expressed as (immature platelet fraction [IPF%], or as an absolute number, i.e.
immature platelet count [IPC, which is the product of IPF% and platelet count]). Using
the Sysmex XE-2100 haematology analyser, a reference interval for immature platelets
has been established for both IPF%, 0.5 to 3.3% (0.5–3.1% in men; 0.5–3.4% in women),
and IPC, 1.25 to 7.02 × 10(9)/L (1.30–6.80 × 10(9)/L in men; 1.21–7.15 × 10(9)/L in
women).[42] This method allows the measurement of immature platelets to be obtained along with
complete blood count measurement, saving time with a minimal added cost. Studies have
shown modest positive correlation between immature platelet analysis and the traditional
flow cytometry method. In patients with thrombocytopaenia of different aetiologies,
the overall correlation was moderate to strong (r = 0.57 and r = 0.65) and was highest in the group of patients with thrombocytopaenia due to peripheral
destruction.[43]
[44]
[45] We have demonstrated that this association tends to be highest (r = 0.63 vs. r ≤ 0.41) in patients with peripheral thrombocytopaenia who have higher IPF% (8.3%)
compared to patients with end-stage renal disease, stable coronary artery disease
(CAD) and post-coronary artery bypass surgery, who have lower IPF% (< 5% for all).
It is important to note that flow cytometry using TO generally reports a higher proportion
of circulating RP compared to IPF% measured using this technique, most likely as a
consequence of properties of non-specific binding of TO, to dense granules and deoxyribonucleic
acid.[46]
The Role of Reticulated Platelets in Thrombotic Diseases
Reticulated Platelets and Coronary Artery Disease
RPs and immature platelet analysis have not clearly distinguished patients with the
various acuity of CAD. The initial study by Lakkis et al evaluated 92 patients with
CAD and showed a significant increase in RP% in patients with ACS compared to stable
angina. Furthermore, RP% increased with the increasing acuity of ACS (10.53% in unstable
angina, 15.99% in non-STEMI, and 18.94% in STEMI).[24] Concordant findings were demonstrated in a larger group of healthy volunteers (n = 22) and patients presenting with CAD (n = 39 with SIHD and n = 359 with ACS) using the Sysmex XE-2100.[25] IPF% steadily and significantly increased from a mean of 2.51% in healthy volunteers
to 3.7% in patients with STEMI. These findings were independently confirmed by others
in a separate study.[23] On the other hand, in a group of 280 patients coming to the emergency room with
chest pain, immature platelets did not differentiate between patients with or without
ACS.[47] However, it is important to note that the latter study included all patients coming
to the emergency room including healthy subjects. It is known that RPs are increased
in many conditions beside CAD and are makers of inflammation, infection and enhanced
bone marrow activity. In a group of 190 patients with symptoms and signs suspicious
for infection, RP% were sensitive and specific for diagnosing infection and changed
dynamically with the progression and recovery of infection.[48] Thus, including all comers to the emergency room with chest pain of different aetiologies
such as pneumonia, costochondritis or cholecystitis to name a few may have contributed
to the elevated level of immature platelets and skewed these results.
Studies have also investigated the relationship between RPs and both short- and long-term
cardiovascular outcomes. In patients admitted with ACS, a high IPF% measured in the
first 24 hours of admission was an independent risk factor for hospital mortality
(odds ratio [OR] = 2.42, 95% confidence interval [CI]: 1.08–5.43; p = 0.032) after adjustment for clinical variables including ST-elevation and troponin
level.[23] In patients with STEMI, the proportion of RPs started to fall 4 hours after percutaneous
coronary intervention (PCI).[49] Moreover, while RPs remain elevated for 30 to 60 days following myocardial infarction
(MI), the level decreases after 1 year.[50] Thus, it is plausible that RPs may report long-term outcomes. Cesari et al measured
RP levels at the time of presentation with ACS and found that RP levels independently
predicted cardiovascular mortality at 1 year (OR = 4.15, 95% CI: 1.24–13.91; p = 0.02).[51] In another study of patients with both SIHD and ACS patients, we observed that RP
levels predicted the major adverse cardiovascular events (MACE) of death, MI, unplanned
re-vascularization and re-hospitalization for angina at a median of 31 months.[52] Patients with an IPC > 7,632 platelets/μL were more likely to experience a MACE
(hazard ratio, 4.65, 95% CI: 1.78–12.16; p < 0.002). Interestingly, RP reactivity as measured by light transmission aggregometry
(LTA) did not predict MACE. These findings suggest that RPs are better chronic predictors
of events than LTA which is more likely to reflect short-term status. In a concordant
observation, RPs also provided better prediction of MACE than either vasodilator-stimulated
phosphoprotein phosphorylation or multi-plate electrode aggregometry in patients following
PCI[53] ([Table 1]).
Table 1
Studies of reticulated platelets and outcomes in coronary artery disease
|
Cesari
|
Ibrahim
|
Freynhofer
|
|
RP method
|
Sysmex XE-2100
|
Sysmex XE-2100
|
Sysmex XE-2100
|
|
Population
|
SIHD
|
0
|
38
|
198
|
|
US/NSTEMI
|
104
|
47
|
164
|
|
STEMI
|
125
|
4
|
124
|
|
Cardiogenic shock
|
0
|
0
|
10
|
|
Total
|
229
|
89
|
486
|
|
Follow-up
|
1 y
|
Median 31 mo
|
Median 190 d
|
|
Outcomes
|
Death
|
22
|
10
|
20 (18 cardiovascular)
|
|
ACS
|
−
|
11
|
21
|
|
Re-vascularization
|
−
|
6
|
27
|
|
Angina
|
−
|
7
|
6
|
|
Stent thrombosis
|
−
|
−
|
10
|
|
TIA/Stroke
|
−
|
−
|
4
|
|
MACE composite
|
22
|
30
|
86
|
|
RP
|
Correlation with MACE
|
Yes
|
Yes
|
Yes
|
|
Cut-off
|
IPF > 3.3%
|
IPC > 7,632
|
IPF > 3.35%
|
|
Sensitivity
|
63.6%
|
70.7%
|
67%
|
|
Specificity
|
77.3%
|
82.1%
|
51%
|
Abbreviations: ACS, acute coronary syndrome; IPC, immature platelet count; IPF, immature
platelet fraction; MACE, major adverse cardiovascular events; NSTEMI, non-ST-elevation
myocardial infarction; RP, reticulated platelets; SIHD, stable ischaemic heart disease;
STEMI, ST-elevation myocardial infarction; TIA, transient ischaemic attach; UA, unstable
angina.
Reticulated Platelets in Acute Ischaemic Stroke
AIS results from embolic or thrombotic occlusion of a cerebral vessel that causes
a focal cerebral ischaemic injury. According to the Trial of Org 10172 in Acute Stroke
Treatment criteria, the causes of AIS have been classified in the following categories:
large vessel atherosclerosis, cardioembolism, acute lacunar and cryptogenic or stroke
of other determined aetiology.[54] Lipohyalinosis, resulting in cerebral small vessel occlusion, is thought to be the
main mechanism related to acute lacunar infarct. Platelets are believed to play a
small role in this syndrome.[55] Overall, initiation of anti-platelet therapy following AIS or transient ischaemic
attack (TIA) is associated with a reduction in the rate of recurrent ischaemic strokes.[56] However, adding single or dual anti-platelet therapy did not result in further risk
reduction compared to aspirin use alone.[57]
[58] More recently, in a randomized clinical trial performed in China, it was determined
that the initiation of dual anti-platelet therapy within 24 hours of TIA or minor
stroke resulted in a lower risk of stroke in the first 90 days.[59] These contrasting results may be due to the heterogeneous aetiologies of AIS and
the potential relationship of AIS to platelet non-responsiveness.[60] Elevated MPV is associated with increased platelet turnover, platelet reactivity
and resistance to anti-platelet therapy, as previously discussed in this review. MPV
was a stroke predictor in patients with atrial fibrillation, after adjustment for
other risk factors.[61] It was also found to be higher in non-lacunar stroke patients.[62] Moreover, elevated MPV was associated with worse acute stroke outcomes.[63]
Despite the relationship of MPV to AIS, studies investigating the role of immature
platelets in stroke incidence, pathophysiology and outcomes are lacking. In one study,
flow cytometric analysis of RP in patients with ischaemic stroke showed significantly
high proportions of circulating RP in patients with cardioembolism compared to control
patients.[64] In another study, the percentage of circulating RP was elevated in the early and
late stages following a stroke compared to control and a correlation with MPV was
observed. However, the MPV was similar between patients with AIS and control patients,
suggesting that RP may play a role beyond what can be measured by MPV alone.[65] Another recent report found that RPs are increased in both early (≤ 4 weeks) and
late (≥ 3 months) symptomatic (TIA or AIS) compared to asymptomatic moderate or severe
carotid stenosis, as measured by IPF% (5.78% vs. 3.48% and 5.11% vs. 3.48%, respectively).
While there was a significant correlation between RPs measured using flow cytometry
and by Sysmex XE-2100, RP% was not different between early or late symptomatic compared
to patients who were asymptomatic. These findings suggest that the automated measurement
of RPs may be more sensitive for RP quantification than is flow cytometry, possibly
as a result of the multiple sources of error that can occur when flow cytometry is
used as a clinical tool.[66] Further research is required to elucidate the role of RP in AIS and TIA.
Role of Reticulated Platelets in Response to Anti-Platelet Drugs
Reticulated Platelets and Response to Aspirin
Since its discovery more than 100 years ago, aspirin has remained one of the most
widely used medicines worldwide. Aspirin reduces the future risk of MI, stroke and
vascular death by approximately 25% in patients with acute MI or with a history of
MI, stroke, or TIA.[67] Aspirin is an irreversible inhibitor of prostaglandin GH synthase-2 aka: COX-1 that
inhibits prostaglandin GH synthase 2 production and subsequently the production of
thromboxane (TxA2).[68]
Although aspirin has a short half-life (30 minutes), its irreversible inhibition of
COX-1 renders platelets unable to form TxA2 and, therefore, unable to aggregate for
their entire lifespan (7--10 days). This irreversible inhibition may explain why an
aspirin dose as low as 40 mg per day can produce a cumulative and sustained inhibition
of platelet function.[69] Previous findings indicate that the platelet pool recovers the ability to produce
thromboxane B2 (TxB2) and aggregate as early as 4 hours after ingestion of 650 mg
of aspirin in normal individuals.[70] In fact when tested in vitro, platelet aggregation in response to a single aggregating
agent (either AA or collagen) occurred when 15 to 20% of platelets were aspirin-free.
On the other hand, only 5% of aspirin-free platelets were needed to achieve platelet
aggregation in response to the combination.[70]
About 100 billion new platelets are produced daily from megakaryocyte to maintain
a sufficient platelet count.[71] Therefore, one might speculate that in patients with increased level of RPs, a higher
turnover will be associated with enhanced early recovery of platelet function. DiMinno
et al[72] first described the relationship between enhanced platelet turnover and aspirin
resistance. These investigators found that the platelets of normal and diabetic individuals
taking aspirin once a day (regardless of the dose 100, 330 or 1,000 mg) were able
to aggregate and form measurable amounts of TxB2 after 12 to 15 hours. In vitro incubation
of PRP with aspirin abolished platelet aggregation and TxB2 production indicating
that recovery of platelet function is a result of aspirin-free new platelets entering
the circulation. In healthy individuals, platelet aggregation and TxB2 production
were diminished when aspirin dosing was increased to four times daily, but this was
not the case for patients with diabetes. However, in vitro incubation of PRP from
diabetic patients with aspirin abolished platelet aggregation and TxB2 production,
indicating the higher platelet turnover in diabetic patients compared to normal individuals.
These findings were again demonstrated in both normal individuals,[73] and in patients with CAD.[74]
The most convincing evidence that enhanced platelet turnover plays a role in decreased
response to aspirin comes from studies of patients with essential thrombocythaemia,
a disease characterized by immune-mediated platelet destruction and naturally increased
platelet turnover. In this patient population, serum TxB2 level and urinary TxM excretion
were significantly higher in patients on aspirin therapy (100 mg/day) compared to
aspirin-treated healthy volunteers.[75] However, the addition of aspirin (50 μM) in vitro completely suppressed production
of thromboxane to levels similar to aspirin-treated healthy volunteers.[75] One plausible mechanism is that platelet turnover results in the release of new
platelets unaffected by aspirin, which is especially important when the turnover rate
is high. However, although in a study of patients with diabetes, twice daily aspirin
dosing (75 mg) led to a reduction of AA-induced whole blood aggregation compared to
75 or 320 mg once daily, the proportion of RP did not discriminate between patients
who benefited from twice daily dosing.[76] Another explanation of how platelet turnover results in aspirin resistance is the
observation that circulating RP possess uninhibited COX-1 as well as COX-2 activity.
In a study of 60 healthy aspirin-treated volunteers stratified into tertiles according
to their RP levels, we observed that post-aspirin thromboxane levels were higher in
the upper tertile compared to the lower tertile and that that these levels decreased
when specific COX-1 or COX-2 inhibitors were added ex vivo.[26] In a different study, the effect of RPs on platelet aggregation appeared to be independent
of platelet turnover. In patients with SIHD receiving dual anti-platelet therapy (aspirin
with either clopidogrel or prasugrel) following PCI, a correlation was observed between
RP and platelet aggregation and was of similar magnitude during early (< 2 hours of
thienopyridine load) and late (24 hours later) phases of dosing.[77]
Reticulated Platelets and P2Y12 Antagonists
Clopidogrel, the most commonly used P2Y12 antagonist, is a pro-drug that must be metabolized
by hepatic CYP450 enzymes to inhibit the binding of ADP to platelet P2Y12. The platelet
response to clopidogrel has wide biological variability between individuals.[78] RPs have recently garnered interest as a plausible explanation for hypo-response
to clopidogrel. In a group of healthy individuals, hypo-responsiveness to clopidogrel
was found in 60% of subjects in the upper tertile of RP values compared to only 10%
of subjects in the lower tertile.[79] An IPF cut-off of 3.6% predicted clopidogrel hypo-responsiveness with a sensitivity
and specificity of 85.7 and 81.8%, respectively. These findings were further extended
to patients with diabetes,[80] SIHD on dual anti-platelet therapy[81] and post-PCI for SIHD[77]
[82] or for ACS.[82]
[83] Among a group of patients who underwent elective PCI, IPC was the strongest independent
predictor of anti-platelet response to P2Y12 inhibitors. In fact, 7% of on-clopidogrel
platelet reactivity was explained by IPC.[28] On the other hand, when platelet function tests were performed 30 to 90 days after
PCI for either SIHD or ACS, neither IPF nor IPC could be correlated with platelet
aggregation.[84] Moreover, while both platelet aggregation and turnover indices were significantly
elevated at time of STEMI and decreased with time over 3 months period, there was
no significant correlation between the two.[49] This difference in observations is likely due to the heterogeneity in these study
designs, including the differences in patient populations, platelet function test
used, agonist concentration, timing of blood collection and the definition of hypo-response
to anti-platelet therapy ([Table 2]).
Table 2
Studies of reticulated platelets implication on anti-platelet function testing
|
Author
|
Population
|
No. of patients
|
Anti-platelet
|
RP method
|
RP value
|
Platelet function
|
Agonist
|
↓Response to anti-platelet definition
|
Correlation with RP
|
Time[a]
|
|
Guthikonda et al[26]
|
Healthy
|
60
|
ASA
|
Flow cytometry
|
RP% (4, 8, 12) lower, middle, upper tertiles
|
LTA
|
Collagen 1.0 μg/mL, AA 1.5 mM, ADP (5, 20 μM)
|
≥70% by AA
|
+
|
24 h
|
|
Würtz et al[91]
|
SIHD
|
124
|
ASA
|
XE-2100
|
IPF 2.2%
IPC 5.2
|
VerifyNow
MEA
|
Collagen 1.0 μg/mL, AA 1 mM
|
Not studied
|
+ only
with IPC
(MEA)
|
1 h
|
|
Grove et al[92]
|
SIHD
|
177
|
ASA
|
XE-2100
|
IPC 6.8
|
VerifyNow
MEA
|
AA 1.0 mM, ADP 10 μM,
collagen 1.0 μg/mL
|
Upper tertile of platelet aggregation
|
+ only
with IPC
|
1 h
|
|
Ibrahim et al[79]
|
Healthy
|
29
|
Clopidogrel
|
XE-2100
|
IPF (2.1, 3.3, 4.9) % in the lower, middle and upper tertiles
|
LTA
VASP-P
|
ADP (2, 5, 20 μM)
|
Aggregation ≥50% in response to 5 μM ADP
|
+
|
NR
|
|
Freynhofer et al[82]
|
Post-PCI (SIHD, ACS)
|
102
|
Clopidogrel
|
NR
|
NR
|
VASP-P
|
NR
|
PRI > 50%
|
+
|
|
|
Guthikonda et al[81]
|
SIHD
|
90
|
ASA, clopidogrel
|
Flow cytometry
|
RP% (1.3, 3.1, 17.9) % in the lower, middle and upper tertiles
|
LTA
|
Collagen 1.0 μg/mL, AA 1.5 mM, ADP 5 μM
|
ASA: Aggregation ≥20% by AA. Clopidogrel: Aggregation ≥50% by ADP
|
+ for ASA and clopidogrel
|
12–24 h
|
|
Cesari et al[83]
|
ACS
|
372
|
ASA, Clopidogrel
|
XE-2100
|
IPF 3.9%
|
LTA
|
ADP 10 μM, AA 1 mM
|
Aggregation > 20% by AA and/or > 70% by clopidogrel
|
+
|
4–6 h
|
|
Mijovic et al[80]
|
Diabetics
|
79
|
ASA, clopidogrel
|
Cell-DYN Sapphire
|
RP%
Diabetic: 3.17%
Non-diabetic: 2.39%
|
ADP test
MEA
TRAP
|
ADP 20 μL,
ASPI 20 μL,
TRAP 20 μL
|
%DPAadp = 100 × (1–ADP/TRAP).
%DPAaspi = 100 × 1–ASPI/TRAP)
|
+ with clopidogrel
none with ASA
|
NR
|
|
Funck-Jensen et al[49]
|
STEMI
|
48
|
ASA, clopidogrel
|
XE-2100
|
Not reported
|
VerifyNow
MEA
|
Collagen 1.0 μg/mL,
AA 1 mM, ADP (6.4, 20) μM
|
MEA (AA > 300, ADP > 416) AUC,
VerifyNow ASA > 550 ARU
P2Y12 > 230 PRU
|
No correlation
|
≤ 1 h
|
|
Perl et al[86]
|
STEMI
|
62
|
Prasugrel
|
Flow cytometry
|
Not reported
|
VerifyNow
MEA
|
ADP
|
MEA: ≥ 47 AU*min,
VerifyNow: ≥208 PRU
|
+
|
NR
|
|
Vaduganathan et al[88]
|
NSTEMI
|
53
|
Ticagrelor
|
Flow cytometry
|
RP% (21.5, 23.8) % at 2–4, and 30 d
|
VerifyNow
MEA
|
ADP
|
MEA: ≥47 AU*min,
VerifyNow: ≥208 PRU
|
No correlation
|
NR
|
|
Eisen et al[50]
|
AMI
|
35
|
Ticagrelor, prasugrel
|
Flow cytometry
|
RP% (17.5, 14.9, 10.5) % at
2–4 d,
30–60 d and
1 y
|
VerifyNow
|
NA
|
≥208 PRU
|
No correlation
|
NR
|
|
Bernlochner et al[87]
|
ACS
|
124
|
Ticagrelor, prasugrel
|
XE-5000
|
IPF 3.8 vs. 3.7%
IPC 8.6 vs. 9.2 in ticagrelor vs. prasugrel group
|
MEA
|
ADP
|
> 468 AU*min
|
+ with prasugrel, none with ticagrelor
|
6–48 h
|
|
Verdoia et al[84]
|
Post-PCI (SIHD)
|
386
|
Clopidogrel, ticagrelor
|
XE-2100
|
IPF (3.5 vs. 3.6) % in diabetics vs. not
IPC
(7 vs. 7.4) in diabetics vs. not
|
MEA
|
AA, Collagen, ADP, PE1, TRAP-6
|
ASA: > 862 AU*min
Clopidogrel: > 417 AU*min
|
No correlation
|
NR
|
|
Stratz et al[28]
|
Post-PCI (SIHD)
|
300
|
Clopidogrel, prasugrel
|
XE-2100
|
Not reported
|
MEA
|
ADP 6.4 μM
|
> 468 AU*min
|
+ with clopidogrel, none with prasugrel
|
Day 1[b]
|
Abbreviations: %DPA, percentage of decrease in overall platelet aggregability; AA,
arachidonic acid; ADP, adenosine diphosphate; AMI, acute myocardial infarction; ARU,
aspirin reaction units; ASA, aspirin; AU*min, aggregation units × minute; AUC, area
under the aggregation curve; ACS, acute coronary syndrome; IPC, immature platelet
count; IPF, immature platelet fraction; LTA, light transmission aggregometry; MEA,
multiple electrode aggregometry; NR, not reported; NSTEMI, non-ST-elevation myocardial
infarction; PCI, percutaneous coronary intervention; PRI, platelet reactivity index;
PRU, P2Y12 reaction units; RP%, percent reticulated platelets; RP, reticulated platelets;
SIHD, stable ischaemic heart disease; STEMI, ST-elevation myocardial infarction; VASP-P,
vasodilator stimulated phosphoretin phosphorylation.
a Time between drug administration and blood sample collection.
b On day 1 prior to intake of first maintenance dose.
In a sub-study of TRITON-TIMI 38 trial, prasugrel, a new P2Y12 inhibitor, resulted
in improved cardiovascular outcomes compared to clopidogrel. However, hypo-response
to prasugrel was reported to be as high as 27%.[85] When platelet function was evaluated 2 to 4 days after PCI in 62 patients with STEMI
who were started on prasugrel, high platelet reactivity was found in 11.3% of patients
and persisted at the 30-day follow-up.[86] Furthermore, RP were strongly correlated with platelet reactivity in patients with
ACS treated with prasugrel.[86]
[87] In contrast to these findings, RP could not be correlated with platelet aggregation
in another study of patients with ACS and treated with prasugrel.[50] However, the small number of patients, higher use of integrin αIIbβ3 antagonists and, most importantly, the very low platelet reactivity (median platelet
reactivity unit value of 25) may have accounted for this finding. As prasugrel irreversibly
binds to P2Y12 and has a half-life of about 7 hours, the RP newly introduced into
circulation will be unexposed to the active metabolite at the end of its once daily
administration. In fact, P-selectin, a marker of platelet activation, was found to
be higher in RP just before treatment with prasugrel compared to 2 hours after—the
daily maintenance dose.[87] These properties may be lessened when ticagrelor, a reversible active inhibitor
of P2Y12 with a longer half-life (11 hours) than the active metabolite of prasugrel
(7 hours), that is administered twice daily, is used.[87] Both ticagrelor and its active metabolites interact equipotently with the P2Y12,
which would be expected to result in more complete suppression of P2Y12 function.
Most studies of platelet reactivity in patients with ACS treated with ticagrelor have
very low residual platelet reactivity, which limits inter-individual variability,
and thus might explain the lack of significant correlation with RP levels.[50]
[87]
[88] This hypothesis was supported in a recent study by Armstrong et al demonstrating
that RPs play a role in hypo-response to clopidogrel but not ticagrelor. These investigators
stained PRP with TO and then stimulated aggregation with ADP and found that RPs disappeared
from the non-aggregated PRP in patients with CAD treated with aspirin and clopidogrel
but not ticagrelor[38] ([Table 2]). Similarly, RPs have been reported not to affect platelet reactivity in patients
receiving cangrelor, an intravenous P2Y12 antagonist. Stratz et al observed a significant
correlation between RPs and platelet aggregation in patients loaded with either clopidogrel
or prasugrel. However, the correlation was not observed but in patients who received
or ticagrelor or during cangrelor infusion in patients undergoing PCI to treat SIHD.[89]
