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Keywords:

  • Coagulopathy;
  • Equine;
  • Thromboelastometry

Abstract

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

Background

Hyperglycemia and endotoxemia have been associated with coagulation abnormalities in horses. Studies in humans suggest greater disturbances in coagulation with hyperglycemia and concurrent endotoxemia.

Objectives

To compare coagulation parameters in horses administered with lipopolysaccharide (LPS) with and without concurrent hyperglycemia.

Animals

Twelve healthy adult horses.

Methods

Hyperglycemia (180–240 mg/dL) was maintained for 6 hours in 6 horses (GLU-LPS) using 140 mg/kg IV bolus of dextrose followed by a 20% dextrose constant rate infusion. A similar volume of saline was administered to an additional 6 horses (SAL-LPS). LPS (20 ng/kg) was administered to each horse. Fibrogen concentration, prothrombin time (PT), activated partial thromboplastin time (aPTT), thrombin antithrombin concentration (TAT), and thromboelastometry were measured at baseline and after 1, 1.5, 2, 2.5, 3, 4, 6, and 22 hours. Repeated measures analysis of variance was used to examine temporal changes.

Results

Increases in PT (= .001) and TAT (= .027) were observed in the GLU-LPS group. Changes in thromboelastometry parameters including increased clot formation time (In-TEM, = .006; Ex-TEM, = .002) and decreased alpha angle (Ex-TEM, = .04) and maximal clot firmness (Ex-TEM, = .014) were observed in the SAL-LPS group. Differences between SAL-LPS and GLU-LPS groups were limited to increased maximal clot firmness (Ex-TEM) at 3, 6, and 22 hours (< .001) in the SAL-LPS group.

Conclusions and Clinical Importance

Minor alterations in coagulation parameters identified for each group are most likely not clinically relevant. Observed differences between groups do not suggest that concurrent hyperglycemia and endotoxemia are associated with greater coagulation abnormalities in horses.

Abbreviations
aPTT

activated partial thromboplastin time

CBC

complete blood count

CFT

clot formation time

CRI

continuous rate infusion

CT

clotting time

HCT

hematocrit

LPS

lipopolysaccharide

MCF

maximal clot firmness

PT

prothrombin time

TAT

thrombin antithrombin concentration

TEM

thromboelastrometry

TNFα

tumor necrosis factor α

WBC

white blood cell

There are strong links between endotoxemia and sepsis with the development of coagulation abnormalities in multiple species.[1-3] Coagulopathy in hospitalized patients is associated with increased morbidity and mortality.[4, 5] Major procoagulant events at sites of inflammation include tissue factor expression, altered thrombogenicity of endothelial surfaces, and platelet activation.[6] Initiation of coagulation during severe infection is mediated primarily by the tissue factor VIIa pathway.[7] Tissue factor induces generation of thrombin, resulting in fibrin formation.[6] Furthermore, mediated by the actions of cytokines, the endothelium becomes a prothrombotic surface during sepsis or endotoxemia.[6]

Hyperglycemia has been associated with increased morbidity and mortality in humans[8] and horses.[9] Horses challenged with IV endotoxin have decreased insulin sensitivity and impaired glucose metabolism.[10] Increased intracellular concentrations of glucose can be cytotoxic to vascular endothelial cells.[11, 12] In studies of humans, hyperglycemia is prothrombotic and activates coagulation, particularly by the tissue factor pathway, regardless of insulin concentration.[13-16] Acute hyperglycemia, regardless of insulin concentration, activated coagulation as demonstrated by increased plasma soluble tissue factor and thrombin antithrombin (TAT) concentrations in humans.[13]

In healthy adult horses, coagulation parameters did not change in response to moderate hyperglycemia sustained for 6 hours.[17] Studies in humans and rodents demonstrated that the impact of hyperglycemia on coagulation abnormalities was significantly greater with concurrent induction of endotoxemia.[18, 19] Given the potential for endotoxemia to contribute to both hyperglycemia and coagulation abnormalities in other species, and given the sensitivity of horses to endotoxin, an investigation of the effect of hyperglycemia with concurrent endotoxemia on coagulation in horses is warranted.

This study investigated the effect of concurrent hyperglycemia and endotoxemia on coagulation parameters in healthy adult horses using a previously used model for sustained hyperglycemia.[17] It was hypothesized that experimental endotoxemia in conjunction with hyperglycemia would be associated with significant changes in markers of coagulation including platelet count, fibrinogen concentration, prothrombin time (PT), activated thromboplastin time (aPTT), TAT, and thromboelastometry parameters (TEM). In addition, we hypothesized that alterations in the coagulation profile induced by endotoxemia and hyperglycemia would be greater than those observed with endotoxemia alone.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

Animals

Healthy horses (12) were used (3 nonpregnant mares and 3 geldings for each group; age, 5–19 years; weight, 423–581 kg). Inclusion was determined based on normal results from physical examination, routine plasma biochemistry,1 and complete blood count (CBC) including platelet count,2 performed before study onset. Horses were allowed a 1-week acclimation period. Horses were housed in individual stalls and fed free choice grass hay. The study was approved by the University of Illinois Institutional Animal Care and Use Committee.

Study Design

Horses were randomized to either hyperglycemia and endotoxemia (GLU-LPS; n = 6) or normoglycemia and endotoxemia (SAL-LPS; n = 6). Feed was withheld for 12 hours before and during the study period. Free access to water was provided at all times. Heart rate, respiratory rate, and rectal temperature were evaluated every 0.5 hour for 6 hours after endotoxin administration, followed by every 2 hours until the 24 hour time point.

Three hours before study onset, catheters (14 gauge 5.25 inch)3 were placed aseptically in both jugular veins, 1 for glucose and insulin sample collection, and 1 for endotoxin and dextrose infusion. A 3-hour time lapse was allowed between catheter placement and study initiation because a previous study showed that placement of jugular catheters caused a transient increase in TAT.[17] Baseline blood samples were collected immediately before initiation of infusions. Blood was collected by direct venipuncture for all coagulation parameters.

Glucose Infusion and Hyperglycemic Clamp

Methods for initiation and maintenance of controlled hyperglycemia in the GLU-LPS group were identical to those previously described.[17] A 50% dextrose solution4 was diluted in saline5 to obtain a 20% solution. Horses in the GLU-LPS group received a priming dose of 20% dextrose at [1.4 × (180−baseline blood glucose concentration)] mg/kg as a 2-minute bolus, followed by an initial continuous rate infusion (CRI) of 20% dextrose at 300 mL/h.6 Blood glucose concentrations were measured every 5 minutes until the target glucose concentration (180–240 mg/dL) was reached (“time 0”). Blood glucose concentrations were determined every 10 minutes for 1 hour, then every 20 minutes for 5 hours. The dextrose infusion rate was titrated in 25–100 mL increments to maintain blood glucose concentration within the target range. The clamp was terminated after 6 hours of sustained hyperglycemia. Blood glucose concentration was measured hourly until glucose concentrations were within the reference range. When target blood glucose concentration (180–240 mg/dL) was reached, defined as time 0, endotoxin was administered as a CRI over 0.5 hour (see below).

In the SAL-LPS group, the 20% dextrose solution was replaced with 0.9% saline for all horses over the study duration. Volumes administered for the loading bolus and CRI were calculated similar to the GLU-LPS group. Horses of the SAL-LPS group received a volume of 0.9% saline (equivalent to the 20% dextrose loading volume for GLU-LPS group), before administration of endotoxin. Completion of the 0.9% saline bolus and initiation of the endotoxin infusion was time 0 for SAL-LPS group. All study details were identical for both groups, with the exception of the use of a stall-side glucometer7 for monitoring of blood glucose concentration in GLU-LPS horses.

Endotoxin Administration

One milliliter of endotoxin (E. coli O55:B5) 1 mg/mL solution8 was added to 1 L 0.9% saline5 to make a solution of 1 μg/mL. Endotoxin (20 ng/kg) was diluted to 10 mL and then added to 250 mL 0.9% saline (260 mL total).6 Endotoxin infusions were administered as IV CRI over 30 minutes in both groups starting at time 0. Each horse received endotoxin at a concentration of 20 ng/kg.

Blood Collection and Processing

Collection of blood for glucose and insulin measurement was done by aspiration from the indwelling catheter with the initial 5 mL discarded. Blood was transferred to lithium heparin tubes9 for glucose analysis and tubes without anticoagulant9 for insulin analysis. Blood in lithium heparin tubes was immediately centrifuged at 2,500 × g for 10 minutes and plasma supernatant collected. Blood in tubes without anticoagulant was allowed to clot and then centrifuged in the same manner. Serum was removed, aliquoted, and stored at −20°C.

Whole blood samples for cell counts and coagulation assays were collected directly into tubes using 21 ga × 1¼” vacutainer needles10 by atraumatic direct venipuncture of the left jugular vein proximal to the site of the indwelling catheter at baseline and 1, 1.5, 2, 2.5, 3, 4, 6, and 22 hours after hyperglycemia reached target range. The order of sample collection was Na EDTA9 sample and then the citrated sample. Citrated samples were collected at 9 : 1 ratio (3.2% citrate) into prewarmed (37°C) 4.5 mL buffered sodium citrate silicone-coated glass vacutainers11 and mixed by gentle inversion.

Complete blood counts were performed within 1 hour of collection. Prewarmed citrated whole blood was used for TEM immediately after collection. Additional citrated whole blood was centrifuged at 2,500 × g for 10 minutes at room temperature for collection of platelet-poor plasma. Plasma was aliquoted into 1.5 mL Eppendorf tubes, frozen on 100% ethanol/dry ice, and stored at −80ºC for analysis.

Assays

Whole blood glucose concentrations were assessed immediately after collection using a point-of-care glucometer7 for GLU-LPS horses and using a clinical laboratory chemistry analyzer1 on the hourly plasma samples for all horses. Serum samples were shipped frozen to an external laboratory12 for measurement of insulin concentration by radioimmunoassay.

Complete blood counts2 were performed in the clinical laboratory at the Veterinary Teaching Hospital. Platelet counts were manually confirmed. Tumor necrosis factor α (TNFα) concentrations in citrated plasma were evaluated by ELISA13 according to manufacturer's instructions.

Plasma coagulation tests (fibrinogen, PT, and aPTT) were performed on a Start 4 coagulometer using Diagnostica Stago14 reagents. TAT was measured by ELISA,15 the use of which was previously validated with equine plasma.[20]

Whole blood TEM was performed by a single individual using a ROTEM16 system according to the manufacturer's instructions. All TEM reactions were performed in duplicate either with manufacturer-supplied TF reagent (Ex-TEM) or ellagic acid (In-TEM) and followed for 1 hour.

Statistical Analysis

Shapiro–Wilk and Anderson–Darling tests of standard normal distribution and the Henze–Zirkler test of multinormality were used. The effect over time in each group was evaluated using repeated measures analysis of variance (ANOVA) with horse as a random effect. Posthoc comparisons were carried out where ANOVA indicated significant differences in the effect. For heart rate, respiratory rate, and temperature, differences between the GLU-LPS and SAL-LPS groups at each time point were assessed using Bonferroni-adjusted P-values. For all variables, the means for each group were compared with their baseline using Dunnett's test. Differences between GLU-LPS and SAL-LPS groups at each time point were assessed using Tukey's Honestly Significant Difference test. Analyses were performed using SAS 9.2.17 Significance was defined as < .05. Data are reported as mean (± standard deviation).

Results

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

All horses safely completed the study. Clinical signs of mild to severe lethargy and mild to moderate colic (eg, pawing, shaking, sweating, and attempting to lie down) were observed in all horses after lipopolysaccharide (LPS) infusion. The clinical response to LPS was variable among horses regardless of experimental group. Subjectively, the onset of clinical signs typically occurred toward the end of the endotoxin infusion (time 0.5 hour).

In both the GLU-LPS and SAL-LPS groups, results of the repeated measures ANOVA indicated significant time differences for heart (< .0001) and respiratory (< .0001) rates, temperature (< .0001), WBC and neutrophil counts (< .0001 both), TNFα (< .0001), and insulin (GLU-LPS, < .0001; SAL-LPS, = .01). Significant time differences for hematocrit (HCT) (= .011) and the TEM parameters clot formation time (CFT; In-TEM, = .006; Ex-TEM, = .002), alpha angle (Ex-TEM, = .04) and maximal clot firmness (MCF; Ex-TEM, = .014) were observed in the SAL-LPS group only. Significant time differences for PT (= .001) and TAT (= .027) were observed in the GLU-LPS group only. Changes in platelet count, fibrinogen, and aPTT were not observed. Significant group and time group differences in measured parameters were limited to the Ex-TEM parameter MCF (group, < .001; time group, = .04).

Heart rate, respiratory rate, and rectal temperature increased compared with baseline post LPS infusion. The maximal increase in heart rate was noted at 2 hours in both groups (GLU-LPS, = .0009; SAL-LPS, < .0001). Maximal increases in respiratory rate were identified at 6 hours (GLU-LPS, < .0001) and 5 hours (SAL-LPS, < .0001). Temperature was highest at 3.5 hours (GLU-LPS, < .0001) and 4 hours (SAL-LPS, < .0001). WBC and neutrophil counts decreased compared with baseline post LPS infusion. WBC counts were lowest at 1.5 hours for both groups (GLU-LPS, < .0001; SAL-LPS, = .005). Neutrophil counts in GLU-LPS horses were lowest at 1.5 hours post LPS infusion (< .0001) and in SAL-LPS horses at 1 hour (= .081). TNFα increased compared to baseline with the highest concentrations at 1.5 hours post LPS infusion (GLU-LPS, = .001; SAL-LPS, < .0001).

Hematocrit increased post LPS infusion in the SAL-LPS group with a maximal increase of 10.7% relative to baseline at 6 hours (= .039) (Fig 1). When compared with baseline values, PT was prolonged post LPS infusion in GLU-LPS horses with the maximal change at 6 hours post LPS infusion (= .016) (Fig 2). Because of sample processing errors, TAT measurements were excluded for 2 horses in each group. TAT analysis on the remaining horses (n = 4, each group) identified increases in TAT at 4 hours post LPS infusion (= .039) in GLU-LPS horses (Fig 3).

image

Figure 1. Change in mean (SD) hematocrit (HCT) over time. Significant changes compared with baseline are indicated by *.

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image

Figure 2. Change in mean (SD) prothrombin time (PT) over time. Significant changes compared with baseline are indicated by *.

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image

Figure 3. Change in mean (SD) thrombin antithrombin concentration (TAT) over time. Significant changes compared with baseline are indicated by *.

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Changes in TEM parameters were identified at the 3 hours time-point for the SAL-LPS group with values for all parameters returning toward baseline by 6 hours (Fig 4A–D). CFT increased (In-TEM, = .006; Ex-TEM, = .001), and alpha angle (Ex-TEM, = .04) and MCF (Ex-TEM, = .008) decreased compared with baseline. MCF was decreased in the SAL-LPS group relative to the GLU-LPS group at all time points post LPS infusion (all < .001).

image

Figure 4. Change in mean (SD) TEM parameters over time. Clot formation time (CFT) In-TEM (A), CFT Ex-TEM (B), alpha (α) angle Ex-TEM (C), and maximal clot firmness (MCF) Ex-TEM (D) are displayed. Significant changes compared with baseline are indicated by *.

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Baseline insulin concentrations for all horses fell within normal laboratory reference ranges and an increase in insulin from baseline was identified in both groups (Fig 5). Minor increases in insulin in the SAL-LPS group were limited to 2 hours post LPS infusion (= .022) whereas increases for the GLU-LPS group were identified at 3 hours (= .021) as well as at 4, 5, and 6 hours (all = .001).

image

Figure 5. Change in mean (SD) insulin over time. Significant changes compared with baseline are indicated by *.

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The volume of 20% dextrose in GLU-LPS horses required to maintain the target range in blood glucose concentration increased over the study duration (< .0001).

Discussion

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

Administration of LPS alone or with concurrent hyperglycemia was associated with changes in hemostatic function, as evidenced by a shift toward hypocoagulability on TEM and PT, and an increase in in vivo thrombin generation as measured by TAT. The majority of these changes occurred within study groups differences were not observed between the GLU-LPS and SAL-LPS groups. Furthermore, the magnitude of these changes was small in all cases and may not be clinically important.

For hemostatic assays, significant differences included increases in TAT and PT for GLU-LPS horses, and changes in Ex-TEM and In-TEM parameters, particularly CFT, for SAL-LPS horses.

Hyperglycemia and insulin resistance occur in critical illness (eg, sepsis, endotoxemia) in humans as a consequence of a hypermetabolic stress state and altered carbohydrate metabolism.[21] This stress hyperglycemia typically is associated with hyperinsulinemia and often is transient.[14] Similar metabolic derangements probably are responsible for hyperglycemia in critically ill horses, and a strong link between these 2 factors has been well established in adult horses.[9] In healthy human subjects, hyperglycemic clamp techniques, with and without insulin modification, have been utilized to evaluate the effects of acute hyperglycemia on markers of coagulation.[2, 8] In 2 separate studies involving healthy adult human subjects undergoing hyperglycemic clamps, acute hyperglycemia, regardless of insulin concentration, resulted in activation of coagulation as demonstrated by increases in TAT.[13, 18] We were previously unable to demonstrate such a change in healthy adult horses, using a similar model.[17] Experimental studies in humans and rodents demonstrate that the effect of hyperglycemia on coagulation is significantly greater with concurrent induction of endotoxemia.[18, 19] It therefore was plausible that changes in coagulation would be seen in a hyperglycemic, endotoxemic model in horses.

The endotoxemia model used in this study is similar to that previously described for horses and was based on previous reports of relevant clinical effects.[10, 23] Changes in vital parameters, hematology, and TNFα were similar to those described,[23] including significant increases in heart rate, respiratory rate, and temperature as well as leukopenia and neutropenia. In addition, the magnitude and time course of increases in plasma TNFα in our study were similar to those reported with a dose of 30 ng/kg.[23] A 20 ng/kg dosage of endotoxin resulted in clinical and hematologic effects similar to those reported here.[10] In that study, 19% of horses failed to respond to endotoxin, suggesting interindividual variability in the effective toxic dose.[10]

Glucose clamp techniques in humans[24] and horses[25, 26] are a reliable means to experimentally induce sustained and controlled hyperglycemia. Because glucose concentrations are held fairly constant, the glucose infusion rate becomes an index of glucose metabolism.[24] The priming dose and maintenance infusion rates we used maintained the target glucose concentration range (180–240 mg/dL) for the study duration. This range was selected because blood glucose concentrations >180 mg/dL in adult horses presenting with acute abdominal disease have been associated with increased mortality.[9] In addition, coagulation abnormalities occurred in humans in whom hyperglycemia was experimentally induced to a similar concentration.[13, 18, 22]

Endotoxemia, both experimentally induced and clinical, is associated with alterations in coagulation parameters in humans,[5] dogs,[27] and horses.[28, 29] In horses in a clinical setting, coagulation abnormalities are relatively common with systemic inflammation, where endotoxemia often is a contributing factor.[1, 4, 30, 31] Endotoxin administered to horses at 50 μg/kg caused significant increases in PT and aPTT and significant decreases in platelet count. In our study, prolongation of PT was only noted in the GLU-LPS horses, whereas changes in aPTT and platelet count were not identified in either group. The difference between that study and the present study may be a result of the lower endotoxin dose.

TEM parameters in the GLU-LPS group were expected to demonstrate a more profound shift toward hypercoagulability as compared with the SAL-LPS group. Prolonged hyperglycemia may cause upregulation of monocyte expression of tissue factor,[32] and previous studies in humans have demonstrated that hyperglycemia activates coagulation.[13, 22] Contrary to what was expected, the TEM-related changes observed in this study occurred in the SAL-LPS group and suggested a minor shift toward hypocoagulation with the associated MCF values representing the only statistically significant difference between groups in this study. Although experimentally induced hyperglycemia also resulted in clinically unimportant hypocoagulability in dogs,[33] in this study, the shift toward hypocoagulablity most likely was secondary to the increase in HCT observed in the SAL-LPS group. Increases and decreases in red cell mass, as measured by HCT, can have an ex vivo effect on TEM[34, 35] with increases in HCT causing TEM results to be consistent with hypocoagulability.[35]

This study found that TAT increased significantly in the GLU-LPS group, but not in the SAL-LPS group. Plasma thrombin concentration is not directly quantifiable because of an extremely short half-life, and it cannot be used to evaluate in vivo thrombin generation. Increases in TAT indicate increased in vivo thrombin generation. Acute hyperglycemia (210–220 mg/dL) over a 6-hour period resulted in significant increases in TAT by 3 hours in humans.[13] In a previous study, TAT values did not increase in response to acute hyperglycemia in horses but placement of an IV catheter caused a transient increase in TAT.[17] In the current study, IV catheters were placed 3 hours before the start of the study, and all samples for coagulation assays were obtained by direct venipuncture.

In contrast to experimental studies in humans[18] and other species,[19] endotoxemia alone or with concurrent hyperglycemia was not associated with marked alterations in coagulation in our study. Limitations to this study include its application to clinical disease and whether the magnitude or duration of hyperglycemia and endotoxemia were sufficient to cause the predicted alterations in coagulation. In clinical disease, both endotoxemia and hyperglycemia often are associated with concurrent comorbidities, and the dysregulation of coagulation is most probably complex involving the interaction of multiple factors. The hyperglycemia and endotoxemia model used in this study was based on similar experimental models in humans.[13, 18, 22] Horses may require a greater challenge with either endotoxin or hyperglycemia. The dose of endotoxin (LPS) administered in this study was based on its ability to humanely induce transient clinical signs of endotoxemia. Endotoxin has been identified in the blood and peritoneal fluid of horses with gastrointestinal disease and associated with increases in mortality.[36, 37] Unfortunately, it is unknown how endotoxin concentration changes over the course of clinical disease, and our knowledge of the optimal dose and method for administration (single or repeated bolus, CRI) is somewhat limited. A recent study comparing a low-dose CRI versus repeated bolus administration of LPS over 48 hours did not identify differences in the inflammatory response among horses.[38]

In this study, a higher rate of dextrose (approximately 2-fold) was necessary to maintain targeted hyperglycemia and insulin concentrations also were higher compared with those in a previous study[17] in which hyperglycemia alone was induced in healthy horses using an identical model. The marked increase in serum insulin concentrations in response to dextrose infusion is the most likely explanation for the increased dextrose infusion rate required to maintain hyperglycemia in the current study. Insulin resistance may be a factor explaining the increases in dextrose needed to maintain blood glucose in the target range. Insulin sensitivity in horses was decreased for 24 hours after administration of endotoxin at a dosage of 20 ng/kg.[10] In addition, horses treated with LPS (35 ng/kg) 24 hours before dextrose and insulin CRI were noted to have higher plasma insulin concentrations at every time-point over a 6-hour period compared with horses that received the same dextrose and insulin CRI without LPS.[39]

In summary, experimentally induced endotoxemia and concurrent moderate hyperglycemia were associated with subtle alterations in hemostatic parameters. As reported previously, experimentally induced endotoxemia alone caused alterations in the coagulation cascade. Additional studies are needed to establish the clinical relevance of hyperglycemia and hyperinsulinemia, and their relationship with coagulation in horses.

Acknowledgments

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References

The authors acknowledge Drs Patricia Flores, Ashley Houtsma, and Michael Miller for assistance with data collection and Dr James Morrissey for loan of the thromboelastometry equipment.16 The study was performed at and funded by the University of Illinois.

Conflict of Interest Declaration: Authors disclose no conflict of interest.

Footnotes
  1. 1

    Hitachi 917 Roche, Block Scientific Inc, Bohemia, NY

  2. 2

    Cell-dyne 3700 hematology analyzer, GMI, Ramsey, MN

  3. 3

    Mila International Inc, Erlanger, KY

  4. 4

    Clipper Distributing Company, St Joseph, MO

  5. 5

    Hospira Inc, Lake Forest, IL

  6. 6

    Baxter Flo-Gard 6301 Infusion Pump, Deerfield, IL

  7. 7

    GM100 glucometer, Bionime Corporation, Dali City, Taiwan

  8. 8

    Sigma, St Louis, MO

  9. 9

    Tyco Healthcare Group LP, Mansfield, MA

  10. 10

    BD Eclipse, Becton, Dickinson and Co, Franklin Lakes, NJ

  11. 11

    BD, Becton, Dickinson and Co

  12. 12

    Michigan State University Diagnostic Laboratory for Population and Animal Health, Lansing, MI

  13. 13

    ThermoFisher Scientific, Rockford, IL

  14. 14

    Diagnostica Stago, Asniernes, France

  15. 15

    Enzygnost TAT Micro, Siemens Healthcare Diagnostics, Marburg, Germany

  16. 16

    Pentapharm GmbH, Munich, Germany

  17. 17

    SAS Institute Inc, Cary, NY

References

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. References