|II.||Blood Samples and Instrumentation|
|IV.||TEG Variables and Correlation with Routine Coagulation Assays|
|V.||Validation Studies and Preanalytical Factors|
|VI.||Clinical Applications in Veterinary Medicine|
|A. Hypercoagulable states|
|B. Hypocoagulable states|
|C. Monitoring anticoagulant therapy|
|D. Platelet function|
Abstract: Thrombelastograph analyzers are point-of-care hemostatic analyzers that provide global assessment of the hemostatic process. Thrombelastography (TEG) detects and provides a continuous recording of the changes in the viscoelastic properties of whole blood from initial clot formation through fibinolysis. TEG has been validated for use in dogs, horses, and cats. Hemostasis research using TEG has focused on test validation, alterations of TEG tracings in animals with naturally occurring diseases, and the use of TEG for monitoring various therapeutic modalities. This article reviews TEG methodology and terminology, including potential sources of preanalytical and analytical errors, the correlation between TEG and other routine hemostatic assays, and current clinical applications of TEG, with emphasis on veterinary medical practice. Data suggest that TEG may be a sensitive and useful adjunctive tool for evaluating an animal with an underlying coagulopathy, including hypercoagulability and hypocoagulability. Additional prospective studies are needed to (1) correlate TEG tracing patterns with a clinical predisposition for bleeding or thrombosis in various disease states and (2) determine whether monitoring and treating hemostatic disorders based on TEG tracings improve clinical outcome.
Hemostasis is a complex physiologic process in which blood vessels, blood cells, and soluble factors in plasma interact in order to prevent hemorrhage and thrombosis. In 1856, the physician Rudolf Virchow suggested that altered blood flow, endothelial injury, and hypercoagulability contribute to the development of venous thrombosis (VT). These factors are known as Virchow's triad and are still considered the basic underlying mechanisms of thrombosis today. Thrombelastography (TEG) is an in vitro diagnostic technique initially introduced in Germany in the late 1940s by Hartert.1,2 TEG integrates the cellular and soluble components of the hemostatic process to yield a global assessment of hemostasis. The technique is based on a continuous detection and recording of changes in the viscoelastic properties of whole blood while it clots. TEG tracings may better reflect the cell-based model of hemostasis3 and thus better predict the kinetics of coagulation compared with routine plasma-based assays. Routine coagulation assays are limited in their capacity to predict hemorrhage or thrombosis, especially in patients who undergo invasive diagnostic and therapeutic procedures. The increased use and interest in TEG in veterinary medicine is based on the potential for TEG results to better predict thrombotic as well as hemorrhagic events in the clinical setting.4,5
The terms thrombelastography, thrombelastograph, and TEG were used generically in the scientific literature until 1996. At that time, Haemoscope Corporation (Niles, IL, USA) named its thrombelastograph analyzer “TEG,” which became a registered trademark of the company. Later, a second analyzer, rotational thromboelastometry (ROTEM), became available (Pentapharm GmbH, Munich, Germany). The ROTEM analyzer is currently used primarily in Europe and has recently been approved by the US Food and Drug Administration for clinical use in the United States.6 Comparison of TEG and ROTEM analyzers is beyond the scope of this review and for a more in-depth comparison the reader is referred elsewhere.7,8 For the purpose of this article, “TEG” will be used generically, and the terms “Haemoscope TEG” or “ROTEM” will be used when referring to specific analyzers.
TEG was originally designed to be a bed-side point-of-care analyzer that utilized whole nonanticoagulated blood. However, recent advances, including the use of anticoagulated blood and a variety of coagulation activators and specific inhibitors, have allowed TEG instrumentation to be utilized by diagnostic laboratories as well.8 Since the 1980s, TEG has been widely used in clinical settings in human medicine, especially during liver transplantation and cardiac surgery. TEG results guide administration of blood products and anticoagulants and predict hemorrhage during surgery and postoperatively.9,10 In veterinary medicine, TEG and ROTEM have been validated for use in dogs, horses, and cats.11–17
The objectives of this review are to introduce TEG technique, including its strengths and limitations, highlight important preanalytical factors and quality control checkpoints that affect TEG results, review published data regarding clinical applications of TEG in human and veterinary medicine, and discuss potential future applications of this technique.
Blood Samples and Instrumentation
Blood should be collected atraumatically into sodium citrate (3.2% or 3.8%) anticoagulant. TEG protocols using anticoagulated citrated blood to which various activators are added have been validated for animals. The use of whole nonanticoagulated blood or anticoagulated blood without activation (recalcified method) is impractical in the veterinary setting and likely leads to an unacceptably high degree of intraassay variation.12,13,17 Various activators and specific inhibitors are currently in use for TEG analysis. Standard reagents and products manufactured by Haemoscope Corporation for use with this company's instrument are kaolin reagent, heparinase-coated cups for monitoring heparin therapy, and a rapid TEG (r-TEG) reagent that includes a mixture of kaolin and tissue factor (TF). Additional nonstandardized activators, including celite and recombinant human TF (rhTF), are also in use.17–20 Reagents for the ROTEM include a TF activator, a contact activator, a reagent that includes TF and a platelet inhibitor for qualitative assessment of fibrinogen, a TF and aprotonin mixture for detection of increased fibrinolysis, and a contact activator with heparinase for monitoring heparin therapy.21 Recalcification serves as the set point from which coagulation is initiated and the tracing begins.
The TEG instruments consist of a pin attached to a torsion wire that is introduced into a cup, preheated to 37°C, containing a 360 μL aliquot of the blood sample (Figure 1). Movement is initiated by the cup (Haemoscope TEG) or the pin (ROTEM). As the blood clots, tension exerted on the wire is translated to an electrical signal that yields a graphic display, or tracing, in which the amplitude of the wire oscillation in millimeters is on the y-axis and the time in seconds is on the x-axis (Figure 2A). Amplitude is directly converted to physical units that represent the strength of the formed clot.
The TEG and ROTEM measure the same variables but utilize different terminology (Table 1). R or CT (clotting time) is the time in minutes from clot initiation until the first fibrin polymers are produced and the amplitude reaches 2 mm. K or CFT (clot formation time) is the time in minutes from the end of R until an amplitude of 20 mm is reached and represents the speed of clot formation. Alpha (α) is the angle in degrees tangent to the curve as K is reached and represents the acceleration/kinetics of fibrin formation and cross-linking. MA or MCF (maximal clot strength) is the maximum amplitude in millimeter and reflects maximal clot strength. G is the modification of MA to physical units. LY30/60 or CLI30/60 are the percent of clot lysis detected at 30 and 60 minutes, respectively, after MA is reached. A normal canine TEG tracing (Figure 2A) illustrates each variable, and examples of tracings that depict hypocoagulable, hypercoagulable, and secondary fibrinolytic states (Figure 2B) are provided.
|Initiation time (time 0–2 mm amplitude)||Minutes||R||CT (clot time)|
|Clot kinetics (time 2–20 mm amplitude)||Minutes||K||CFT (clot formation time)|
|Clot kinetics (the slope between 2 and 20 mm amplitude)||Degrees||α angle||α angle|
|Maximal clot strength and stability||Millimeters||MA (maximum amplitude)||MCF (maximal clot strength)|
|Maximal clot strength and stability||Dynes/cm2||G||—|
|Clot lysis||Percent||LY30, LY60||CLI30, CLI60|
TEG Variables and Correlation with Routine Coagulation Assays
Multiple studies have examined how the results of TEG analysis correlate with standard coagulation assays.20,22–25 Traditionally, the R time (Figure 2A) has been associated with soluble clotting factor activity.20K time and α angle (Figure 2A) are nonspecific and have been associated with platelet concentration and function, fibrinogen concentration and function, and clotting factors activity.20 The MA and G, a value calculated from MA and discussed together with MA, have been associated with platelet concentration and function and, to a lesser degree, with fibrinogen concentration.20 As TEG usage has increased, these previously ascribed correlations between TEG results and other hemostatic assay results (such as clotting factor activity and platelet concentration) have been modified. Although some studies have demonstrated a significant positive correlation between R time and prothrombin time (PT) or activated partial thromboplastin time (aPTT), these results could not be reproduced in other studies.9,13,20,24–26 PT correlated with R time using rhTF-Haemoscope TEG in dogs admitted to an intensive care unit (ICU) and in healthy horses13,26; however, there was no correlation between R time and PT or aPTT in healthy people or in people with malignancies when using native or celite-activated Haemoscope TEG.20
Fibrinogen concentration has been shown to be highly correlated with G/MA and K time. This has been demonstrated utilizing various activators and methodologies, both in people and in various animals.13,20,26–29 In a group of human patients with solid tumors, hyperfibrinogenemia had a sensitivity and specificity of 86.5% and 83.3%, respectively, for a ROTEM tracing indicating hypercoagulability.27 Hyperfibrinogenemia is likely an independent risk factor for both venous and arterial thrombosis.30,31
Platelet concentration also correlates with G/MA and K time in dogs and in people.20,22,26,29 The correlation between platelet count and G/MA was stronger after application of a logarithmic transformation to the platelet concentration.22,24 We and others have noted that platelet concentration correlates with G and MA when platelet concentration is within the reference interval or decreased; however, thrombocytosis (>400,000 platelets/μL) does not correlate with an increase in G and MA,29 an observation consistent with the concept that whereas thrombocytopenia may contribute to a hemorrhagic phenotype, reactive thrombocytosis is not associated with hypercoagulability.32,33
In human medicine, there is an association between increased D-dimer concentration, fibrinogen/fibrin degradation products (FDPs), and increased TEG fibrinolytic variables (LY30 or CLI30; Table 1 and Figure 2B).9,25,34 The TEG markers of hyperfibrinolysis may be superior to other laboratory measurements in human trauma and liver transplantation settings.9,25,34,35 To date, establishing similar correlations in veterinary medicine has had little success.29 In dogs with experimentally induced thrombus formation, there was good agreement between TEG fibrinolysis variables, FDPs, thrombus size, and local blood flow at the site of the thrombus after fibrinolytic therapy.36 However, in a recent case series of dogs with disseminated intravascular coagulation (DIC) and subsequent secondary fibrinolysis, there was no association between D-dimer concentration and increased rhTF-Haemoscope TEG fibrinolytic variables.29 Similar results were reported in a study in which human patients with severe sepsis, with or without overt DIC, had increased D-dimer concentration with no concurrent increase in ROTEM fibrinolytic markers using TF activation.37 The latter findings are compatible with our experience, in which D-dimer concentration correlated poorly with kaolin-activated Haemoscope TEG variable, LY30 (unpublished data).
Validation Studies and Preanalytical Factors
In veterinary and human medicine, validation studies have been conducted to better characterize and define preanalytical and analytic factors that affect TEG variables.12–14,17,38–41 As with most hemostatic assays, preanalytical factors, including the delay between blood collection and analysis and the sample temperature, are important in TEG analysis. Blood for TEG analysis should be collected using a standardized protocol to minimize preanalytical variation. In several studies, a clinically acceptable intraassay coefficient of variation (CV) for the various Haemoscope TEG variables in dogs has been reported and varies from 3% to 18% depending on the variable.12,17 Storage of blood for 120 minutes may result in a mild but significant trend toward a tracing that indicates hypercoagulability.17,41 In horses, acceptable intraassay CVs of 1–12%, depending on the variable, were found when using both the Haemoscope TEG and ROTEM analyzers,13,14,16 although effects on results were noted as soon as 60 minutes after blood collection in one study.14 Blood samples refrigerated at 4°C had decreased α angle and prolonged CT but also increased MCF, changes that are consistent both with hypocoagulability and hypercoagulability, respectively.16 These findings are consistent with those noted in human medicine. In human medical practice, 120 minutes is an acceptable period of time between blood collection and TEG analysis with all variables having clinically acceptable CVs using various activators.38–40 Variability in TEG results has also been attributed to differences between analyzers and operators.13,38 These data indicate that TEG is valid for clinical use in dogs, horses, and cats although special attention should be given to minimize preanalytical factors, such as traumatic venipuncture, increased storage time, aberrant temperature, and in vitro hemolysis, as possible sources of error.
In addition to preanalytical factors, red cell mass correlates with various TEG variables. Evidence suggests there is an inverse linear correlation between hematocrit and G/MA and α angle in people and various animal species.16,20,42–46 Whether decreased red cell mass truly reflects in vivo hypercoagulability is subject to debate.43,44,46,47 In people and dogs, decreased and increased hematocrit has been associated with hypercoagulability and hypocoagulability, respectively, in vivo.16,20,42–46 Isolated reduction of hematocrit, in people, with no change in platelet concentration, coagulation factor concentration, or anticoagulant concentration accelerated blood coagulation, as demonstrated with celite-activated samples using the Haemoscope TEG and ROTEM.44,46 Moreover, people with iron deficiency anemia and splenectomized thalassemic individuals had ROTEM tracings that were consistent with hypercoagulability.43,44 However, a separate measure of hypercoagulability, the endogenous thrombin potential test, failed to provide supporting results.43,44 Some authors have interpreted these data to suggest that decreased red cell density will result in an artifact in tracings that suggests hypercoagulability. However, others have argued that given the increased risk for thrombosis in thalassemic patients and the lack of increased potential for thrombin generation, the ROTEM may be superior to the plasma-based endogenous thrombin potential assay in detecting hypercoagulability that precedes thrombosis. In animals, increased hematocrit was associated with TEG variables indicative of hypocoagulability in Greyhound dogs and in transgenic mice with marked erythrocytosis.42,45 In both studies, TEG tracings that indicated hypocoagulability were associated with hypocoagulable phenotypes, ie, an increased bleeding tendency. However, when the murine blood was diluted in vitro with maintenance of the platelet count, the TEG tracing normalized.45 Induction of moderate-to-severe in vitro hemolysis in canine and equine citrated blood samples resulted in significantly decreased G and MA values, consistent with hypocoagulability.16,48 This was attributed to in vitro platelet activation with resultant hyporeactive platelets contributing to decreased clot strength.48
Clinical Applications in Veterinary Medicine
TEG has been studied in various clinical settings, mostly in dogs. Although recently validated for use in horses and cats, clinical studies have yet to be published outlining the clinical utility of TEG in these species. However, published abstracts suggest that the clinical use of TEG is being focused on critically ill foals and horses with colic.49–51 There are sporadic reports of evaluation of coagulation in cattle, pigs, sheep, guinea pigs, rats, and various species of fish using TEG,52–59 but, to date there are no widely accepted guidelines to direct appropriate clinical use or interpretation of TEG tracings in these species.
VT is a serious and life-threatening complication of many underlying disease processes, including neoplasia, immune-mediated hemolytic anemia (IMHA), sepsis, heartworm disease, hyperadrenocorticism, and others; associated risk factors for the development of VT in animals are well described.60–62 Routine hemostatic assays have poor sensitivity for the detection of hypercoagulability,5,26,29,63,64 and TEG has been evaluated for its capacity to provide early, sensitive detection of hypercoagulability.26,29,50,51,63–68 Discrepancies among studies, including variability of TEG methodology and the occasional absence of concurrent hematologic data, including platelet count, fibrinogen concentration, and hematocrit, limit our ability to fully interpret and compare TEG results among studies. Interpretation is further hindered by the absence of long-term prospective studies that confirm a positive correlation between TEG tracings that indicate hypercoagulability and true risk of thrombosis in the animal. To date, hypercoagulability, as evidenced by alterations in the TEG components G and MA, has been demonstrated in dogs with parvoviral enteritis, neoplasia, DIC, and IMHA and in dogs admitted to ICU.26,29,63–65
In an early case–controlled study, 9 puppies with naturally occurring parvoviral enteritis were compared with 9 age-matched control dogs.63 Hypercoagulability was detected in all of the puppies with parvoviral entiritis, as evidenced by an increased MA using recalcified citrated blood. These puppies also had significantly increased fibrinogen concentrations compared with control puppies. Four of the dogs with parvoviral enteritis developed clinical evidence of catheter-associated VT or phlebitis.63
Underlying malignancy is considered a risk factor for thrombosis in animals and people.60–62,69–73 The proposed mechanisms of thrombus formation are not fully characterized but may include altered plasma- or cell-based procoagulant or fibrinolytic activity, increased production of cytokines, including tumor necrosis factor, interleukin-1β, and vascular endothelial growth factor, by tumor cells,72 aberrant TF expression, procoagulant factor production by neoplastic cells, and antiphospholipid antibody production.72,74 In one study using rhTF-Haemoscope TEG, 67% of dogs with malignant neoplasia had hemostatic dysfunction, defined as an altered G value. Of these dogs, 75% were hypercoagulable and 25% were hypocoagulable.64 Dogs with malignant neoplasia were significantly more likely to be hypercoagulable compared with dogs that had benign neoplasms, although 31% of the dogs with benign neoplasms were also hypercoagulable.64 Hypercoagulability was not attributable to fibrinogen concentration as there was no significant difference in the fibrinogen concentrations of dogs with malignant compared with those with benign neoplasia. No information was provided on the overall correlation between fibrinogen concentration and G/MA or the number of dogs with TEG tracings indicative of hypercoagulability that developed VT. These findings are consistent with a similar study in which people with malignant neoplasms had increased G/MA values.27
TEG tracings that indicated hypercoagulable, normocoagulable, and hypocoagulable states were also found in dogs with DIC. In 50 dogs diagnosed with DIC based on a wide battery of hemostatic assays, tracings obtained using rhTF-TEG indicated that 22% were hypocoagulable, 34% were normocoagulable, and 44% were hypercoagulable.29 Interestingly, hypocoagulable dogs had a 2.4 times greater relative risk of death within 28 days than hypercoagulable dogs. The G value was concluded to best reflect the hemostatic status of dogs with DIC, and a TEG tracing indicating hypercoagulability was considered a favorable prognostic indicator.29 In a similar study in human patients with severe sepsis with or without overt DIC, the ROTEM MCF value (equivalent to the MA value) in patients with severe sepsis did not differ significantly from the value in the healthy control group; however, patients with overt DIC had significantly decreased MCF, indicative of hypocoagulability.37
Hypercoagulability was also a common hemostatic abnormality in dogs admitted to an ICU with a variety of underlying disorders and in dogs with IMHA.26,65 Eleven of 27 dogs in the ICU were classified as hypercoagulable by rhTF-TEG with supportive evidence that included decreased antithrombin activity and increased D-dimer concentration.26 The majority of dogs with IMHA had TEG tracings, obtained using recalcified citrated blood, indicating hypercoagulability with only 6 of 39 dogs classified as normocoagulable.65 One potential confounding factor in the dogs with IMHA was that all dogs in the study were pretreated with corticosteroids, which are suspected to induce a hypercoagulable state75 and alter TEG variables.68 The investigators claimed that, in this specific clinical setting, a TEG tracing indicating normocoagulability was a poor prognostic marker.65
Evidence in human clinical settings supports the hypothesis that a hypercoagulable state indicated by a TEG tracing is predictive of thromboembolic events, especially postoperatively.18,76–78 Nonetheless, the data are not consistent. In a recent review paper, the predictive accuracy of TEG results for postoperative thromboembolic events was judged to be “highly variable,” and the authors recommended further prospective studies.76 Certainly in veterinary medicine, prospective studies are warranted to better characterize the effects of fibrinogen concentration, platelet concentration, and red cell mass on TEG tracings and to determine the sensitivity, specificity, positive predictive value, negative predictive value, and accuracy of TEG tracings that indicate hypercoagulability in predicting thromboembolic events in various clinical settings.
Accurate detection of hypocoagulability with resultant increased risk of hemorrhage could guide transfusion and hemostatic therapy during and after surgery or other invasive procedures, such as liver biopsy. Coagulation assays such as PT and aPTT, fibrinogen concentration, and platelet concentration are commonly used in both veterinary and human medicine to assess the risk of bleeding as a result of a diagnostic or surgical procedure. However, a poor correlation was found in people between prolonged PT and hemorrhage after invasive procedures,4 leading to the conclusion that there was insufficient evidence to support the use of transfusion before procedures based on results of plasma-based hemostatic tests.4 Similarly, in veterinary medicine, 2 retrospective studies have demonstrated variable correlation between coagulation test results and bleeding after fine-needle aspiration and tissue-core biopsy.79,80 Given the poor capacity of these assays to predict bleeding, the use of TEG to accurately detect hypocoagulability and predict bleeding events has been evaluated.
TEG may be superior to plasma-based assays in its capacity to correctly predict hemorrhagic episodes in dogs.23,81,82 In a prospective study that compared the hemostatic phenotype in 27 dogs in hypocoagulable states, 27 in normocoagulable states, and 27 in hypercoagulable states based on results from rhTF-TEG, it was found that G value alone had a positive and a negative predictive value for bleeding of 89% and 98%, respectively. Moreover, G value more accurately predicted bleeding than the combination of platelet concentration, PT and aPTT results, D-dimer concentration, and fibrinogen concentration.23 TEG results may also predict bleeding in dogs with severe factor VIII deficiency (hemophilia A).81 In one study, bleeding was induced in anesthetized dogs with severe factor VIII deficiency and bleeding time was documented. The Haemoscope TEG component G, obtained using recalcified citrated blood, was superior to aPTT in predicting bleeding in vivo.81 TEG tracings showed incremental improvement associated with a dose-dependent response to therapy, whereas standard plasma-based assays failed to do so.81 TEG tracings may also be useful in monitoring hemostatic patterns and developing exercise regimens in dogs with severe hemophilia A; however, in this study, correlation of TEG results with hemostatic phenotype or bleeding tendency was not attempted.82 Finally, a tracing indicating hypocoagulability, obtained by rhTF-TEG, was associated with a poorer prognosis and increased mortality risk in dogs admitted to ICU with a clinical suspicion of DIC.29 These findings are supported by a new study in which a hypocoagulable state, based on kaolin-activated Haemoscope TEG tracings, in people admitted to ICU was an independent risk factor for death within 30 days.83
Monitoring anticoagulant therapy
Low-molecular-weight heparin (LMWH) is increasingly being used in veterinary medicine for treatment of thromboembolic diseases and as thrombophylaxis in animal patients at increased risk for thrombosis.84–88 Anti-factor Xa activity is considered to be the “gold standard” for monitoring the effect of heparin on coagulability; however, this assay is expensive and not readily available. TEG has been assessed for its efficacy in monitoring heparin therapy in animal and human patients.85,87–89 In recent prospective studies, healthy dogs21 and cats19 were given therapeutic doses of different heparins, including LMWH, subcutaneously. Only treatment with unfractionated heparin reached therapeutic levels as evidenced by anti-Xa activity and marked prolongation of Haemoscope TEG R time (recalcified method) with no clot formation up to 12 hours after a single subcutaneous administration. The in vitro effects of LMWH have also been tested on citrated canine blood.87 TEG tracings showed dose-dependent prolongation of R and K times and decreased clot strength using rhTF-TEG and heparinase-coated cups, whereas TEG tracings using kaolin-activated TEG and heparinase-coated cups were unremarkable, except for a prolonged R time.87 Similar results have been found in a recent study that explored the use of native Haemoscope TEG and heparinase-coated cups for monitoring LMWH for thrombophylaxis therapy in people with increased risk for deep vein thrombosis (DVT).89 The authors suggested that concurrent measurement of Haemoscope TEG using plain and heparinase-coated cups may detect anticoagulated patients at increased risk of developing DVT; however, sensitivity, specificity, positive predictive value, and negative predictive value were not calculated.89 To date, it appears that TEG may have some clinical utility for the monitoring heparin therapy in animals; however, expected TEG results will depend on the choice of activator, the choice of plain or heparinase-coated cups, and the dose and type of heparin used.
Congenital and acquired disorders of platelet dysfunction are well characterized and relatively common in animals and have been reviewed.90–92 Acquired platelet dysfunction can occur secondary to uremia, infection with various agents, such as Ehrlichia canis and FeLV, snake envenomation, neoplasia, liver disease, and drug administration, especially nonsteroidal antiinflammatory drugs (NSAID).92–98 The use of specific platelet inhibitor drugs, such as clopidogrel (platelet ADP chemoreceptor [P2Y12] inhibitor), and glycoprotein (GP) IIb/IIIa inhibitors may predispose animals to bleeding. As such, there is interest in platelet function assays that are readily accessible, reliable, and cost-effective for monitoring antiplatelet therapy or diagnosing congenital or acquired platelet disorders.99,100
Platelets have multiple physiologic activators and are a major contributor to the TEG G/MA value. Thrombin is a major and powerful platelet activator, and its presence masks the detection of platelet dysfunction that results from alteration of pathways of weaker activators, such as ADP or collagen.101 Sporadic studies have shown that TEG tracings did not differ between normal dogs and dogs with either a specific platelet dysfunction, such as Scott syndrome, or dogs that were treated with NSAID.102,103 Haemoscope TEG PlateletMapping is a modified TEG assay specifically designed to asses platelet function.101 The concept behind PlateletMapping is to eliminate the potent effect of thrombin on blood platelets during clot formation. Blood is drawn into heparin and further activated with reptilase and factor XIIIa to allow a small, polymerized fibrin clot to form in the absence of thrombin. The clot serves as a scaffold for platelet activation. Two other TEG cups are used and platelet agonists, ADP and arachadonic acids, are added to induce platelet activation. A fourth heparinase-coated TEG cup and kaolin activation are also used. Equations are derived to assess the percent MA aggregation response to an agonist. Haemoscope TEG PlateletMapping showed good correlation with optic platelet aggregometry in detecting platelet dysfunction after in vitro inhibition by antagonists of various platelet receptors, such as GP IIb/IIIa, P2Y12, and thromboxane A2 receptor.101 Preliminary data suggest that Haemoscope TEG PlateletMapping may be used to assess platelet inhibition in dogs that are treated with clopidogrel.104 The primary uses to date for TEG PlateletMapping include monitoring platelet inhibition therapy and diagnosing naturally occurring thrombocytopathies; however, this novel technique may also be able to identify platelet hyperreactivity.105 Recently, PlateletMapping was also validated for use in ROTEM analyzers.106
TEG is an old technique being used in new ways to advance our understanding of hemostasis. TEG has been validated in dogs, horses, and cats. TEG analysis should be performed on citrated blood by experienced personnel using standard protocols in which preanalytical factors, including storage temperature and time to analysis, are controlled. Reference intervals should be generated locally using specific activators. Hematologic measurements that may confound TEG analyses, including fibrinogen concentration, platelet concentration, and RBC mass, should be considered when interpreting TEG results. We believe that TEG analysis provides complementary information to routine coagulation assays. Additional long-term prospective studies are needed to determine whether TEG can be used to (1) guide and monitor blood product transfusion and anticoagulant therapy or (2) accurately predict severe clinical complications associated with disruptions in hemostasis, including thrombosis and hemorrhage.
Disclosure: The authors have indicated they have no affiliations or financial involvement with any organization or entity with a financial interest in, or in financial competition with, the subject matter or materials discussed in this paper.