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Comparison of haematology, coagulation and clinical chemistry parameters in blood samples from the sublingual vein and vena cava in Sprague −Dawley rats J Seibel, K Bodié, S Weber, D Bury, M Kron and G Blaich Lab Anim 2010 44: 344 DOI: 10.1258/la.2010.009049 The online version of this article can be found at: http://lan.sagepub.com/content/44/4/344

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Original Article Comparison of haematology, coagulation and clinical chemistry parameters in blood samples from the sublingual vein and vena cava in Sprague – Dawley rats J Seibel1, K Bodie´1, S Weber1, D Bury1, M Kron2 and G Blaich1 1 Abbott GmbH & Co. KG, Preclinical Safety, D-67061 Ludwigshafen, Germany; 2Abbott GmbH & Co. KG, Global Data Management and Statistics, D-67061 Ludwigshafen, Germany Corresponding author: Karen Bodie´. Emails: [email protected]; [email protected]

Abstract The investigation of clinical pathology parameters (haematology, clinical chemistry and coagulation) is an important part of the preclinical evaluation of drug safety. However, the blood sampling method employed should avoid or minimize stress and injury in laboratory animals. In the present study, we compared the clinical pathology results from blood samples collected terminally from the vena cava (VC) immediately before necropsy with samples taken from the sublingual vein (VS) also prior to necropsy in order to determine whether the sampling method has an influence on clinical pathology parameters. Forty-six 12week-old male Sprague–Dawley rats were assigned to two groups (VC or VS; n ¼ 23 each). All rats were anaesthetized with isoflurane prior to sampling. In the VC group, blood was withdrawn from the inferior VC. For VS sampling, the tongue was gently pulled out and the VS was punctured. The haematology, coagulation and clinical chemistry parameters were compared. Equivalence was established for 13 parameters, such as mean corpuscular volume, white blood cells and calcium. No equivalence was found for the remaining 26 parameters, although they were considered to be similar when compared with the historical data and normal ranges. The most conspicuous finding was that activated prothrombin time was 30.3% less in blood taken from the VC (16.6 + 0.89 s) than that in the VS samples (23.8 + 1.58 s). Summing up, blood sampling from the inferior VC prior to necropsy appears to be a suitable and reliable method for terminal blood sampling that reduces stress and injury to laboratory rats in preclinical drug safety studies.

Keywords: Refinement, blood sampling, vena cava, sublingual vein, equivalence Laboratory Animals 2010; 44: 344– 351. DOI: 10.1258/la.2010.009049

The investigation of clinical pathology parameters (haematology, clinical chemistry and coagulation) is an integral part of the preclinical toxicological safety evaluation of pharmaceutical products. In the past, several methods have been recommended for blood sampling in rodents, including sampling from the retrobulbar plexus, jugular vein, sublingual vein (VS), vena cava (VC), lateral tail vein or cardiac puncture.1 – 5 Each of these methods has advantages and disadvantages. For instance, sampling from the retrobulbar plexus is a fast and efficient procedure but may cause severe tissue damage.6 Adverse effects caused by blood sampling may include bruising, haemorrhaging, thrombosis and handling-induced stress.5 Moreover, the need for anaesthesia during some blood sampling methods may adversely affect laboratory animals.3 Causing unnecessary stress to animals is not only unacceptable from an ethical point of view but may also affect the results of clinical pathology analysis and the quality of the blood samples.1 Laboratory Animals 2010; 44: 344 –351

Our standard method for blood withdrawal in rats has been via sublingual venous blood sampling. Due to the need for anaesthesia, blood sampling has to be performed one day prior to necropsy. To avoid causing additional stress in the rats, we investigated the suitability of blood sampling from the VC immediately before necropsy. To validate the method, we measured the routine clinical pathology parameters (haematology, coagulation and blood chemistry) in the VS blood samples of untreated male Sprague – Dawley rats and compared them with corresponding values from the VC samples.

Materials and methods Animals Forty-six male Sprague –Dawley (Crl:CD(SD)) rats were obtained from Charles River Laboratories (L’Arbresle,

Seibel et al. Comparison of blood sampling methods

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France). The rats were 11 weeks old upon arrival and were randomly allocated to two groups (Group 1 ¼ VC; Group 2 ¼ VS; n ¼ 23 each). Following physical examination by a veterinarian, the rats were allowed to acclimatize for seven days before the study started. All rats were individually housed in U-TempTM Type III cages (Tecniplast SpA Buguggiate, Italy), under barrier conditions with bedding (dust-free softwood shavings; Ssniff GmbH, Soest, Germany) containing chewing wood (Ssniff GmbH) as an enrichment under controlled conditions (temperature 22 + 28C, humidity 55 + 10%, light –dark (LD) 12:12). The rats had ad libitum access to a standard rodent diet (R/M-H, pelletsw, Ssniff GmbH) and acidified tap water ( pH 2.5– 2.8). Food was removed overnight for 18 h prior to blood withdrawal (2 –3 h prior to the dark phase). The rats were moved to clean cages once a week. All rats were examined at least once daily. Body weights were measured upon arrival and on the day of terminal blood sampling. Rats were housed, treated and sacrificed in accordance with the regulations of Germany and the European Union, including the German Animal Welfare Act that implements European Council Directive 86/609/EEC. Abbott’s laboratory animal programmes in general as well as the local facility conducting the study experiments are accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC). All the experiments were conducted according to the standards set forth in the National Research Council’s Guide for the Care and Use of Laboratory Animals and were approved by the government of Rhineland-Palatinate.

coagulation parameters, blood was collected in 1.5 mL citrated (0.11 mol/L, 0.1 þ 0.4 mL blood) micro-test tubes (Sarstedt AG & Co) and stored at RT until measurement on the day of sampling. Clinical chemistry samples were collected in 1.1 mL Z-Gel micro-tubes (Sarstedt AG & Co) and then kept at 2 –88C until they were centrifuged at 1850 g for 10 min and then stored at 2– 88C until evaluation. All rats were euthanized by exsanguination under deep Isofluranew anaesthesia (rats of the VC group immediately after blood sampling). Haematology and coagulation parameters Haematology parameters were evaluated on an Advia 120 (Siemens AG, Munich, Germany) analyser. The following parameters were evaluated: red blood cells (RBC), haemoglobin (HGB), haematocrit (HCT), mean corpuscular volume (MCV), mean corpuscular haemoglobin (MCH), mean corpuscular haemoglobin concentration (MCHC), red cell distribution width (RDW), absolute reticulocytes (ARETIC), percent low RNA reticulocyte ratio (LOWRET), percent middle RNA reticulocyte ratio (MEDRET), percent high RNA reticulocyte ratio (HIGHRET), white blood cells (WBC), platelet count (PLT), mean platelet volume (MPV), absolute basophils (ABSBASO), absolute esoinophils (ABSEOS), absolute lymphocytes (ABSLYMP), absolute monocytes (ABSMONO) and absolute neutrophils (ABSNEUT). For the evaluation of coagulation, activated partial thromboplastin time (APTT) and prothrombin time (PT) were measured using a KC 10A coagulometer (Amelung GmbH, Lemgo, Germany). Clinical chemistry parameters

Blood sampling methods and sample handling Blood samples (2 mL/rat) were collected by puncturing the VS or the VC. For sampling from the VS, the rats were lightly anaesthetized with Isofluranew (Abbott GmbH & Co KG, Wiesbaden, Germany) in an inhalation chamber and held in a supine position. The tongue was gently pulled out and one of the VSs was punctured with a 23G 100 hypodermic needle (Terumo Europe NV, Leuven, Belgium). The rat was then turned over to allow blood (2 mL/rat) to drip directly into the labelled test tubes (see below). After sampling, the rats were returned to their cages. For sampling from the VC, the abdominal cavity was opened following deep inhalation anaesthesia with Isofluranew, which was induced in an inhalation chamber and then maintained by supply via a respiratory mask, and the VC was exposed. Venous blood (approximately 2 mL/rat) was drawn using a 23G 100 hypodermic needle and a 2 mL syringe (B Braun Melsungen AG, Melsungen, Germany). The blood obtained by these methods was then transferred to the labelled test tubes. Blood samples for haematology evaluation were collected in 3 mL sample tubes (Sarstedt AG & Co, Nuembrecht, Germany) using K2 EDTA as the anticoagulant, stored at room temperature (RT) and evaluated on the same day. For the evaluation of

The following clinical chemistry parameters were determined using a Hitachi 917 E (Roche Diagnostics GmbH, Mannheim, Germany) analyser: alanine aminotransferase (ALT), aspartate aminotransferase (AST), g-glutamyltransferase (GGT), alkaline phosphatase (AP), total bilirubin (BIL), cholesterol (CHOL), triglycerides (TRIG), glucose (GLU), creatinine (CREA), inorganic phosphorus (PO), blood urea nitrogen (BUN), total protein (PR), albumin (ALB) sodium (Naþ), potassium (Kþ), calcium (Ca2þ), chloride (Cl2) and carbon dioxide (CO2). Statistical analysis The sample size calculation was based on a two-group t-test of equivalence in means. At a sample size of 23 per group, the study had 80% power to reject the hypothesis that the groups are not equivalent for a single parameter, based on the assumption that the expected difference in means is 0, that the equivalence limit is 1 common standard deviation (SD) (the variation within a group), and that each one-sided two-group t-test is made at the 2.5% level. All the parameters investigated were expressed as arithmetic means + SD and coefficient of variation. Additionally, percentage differences of means defined as (MeanVC 2 MeanVS)/MeanVS were calculated. Since the parameter differential blood count is not continuous, the median,

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first and third quartile, Hodges –Lehmann estimator7 of difference in medians and percentage difference in medians of VC compared with VS were calculated. For graphical comparison of the two groups, values are displayed as box-and-whisker plots with the whiskers extending to the adjacent values. Adjacent values were defined as the furthest observation within 1.5 times the interquartile range below or above the quartiles. More extreme values lying below or above the adjacent values were plotted as single values. To test for equivalence, the confidence interval (CI) inclusion rule, which is standard in clinical trials on bioequivalence or on equivalence of other clinical efficacy parameters, was used.8 For each of the investigated parameters (except for ALT and AST) 95% CIs for the difference in means between the two groups were calculated and compared with the equivalence limit of +1 SD ( pooled within group SD). The groups were considered to be equivalent if the upper limit of the 95% CI of the mean difference between groups was below the equivalence limit of one SD and the lower limit was above 21 SD. Calculations were based on the assumption that the parameters are normally distributed, verified by inspection of normal probability plots, and that the within-group variances are the same. The parameters ALT and AST display logarithmic normal distributions. Therefore, for these parameters, the values had to be logarithmically converted prior to calculating the 95% CIs for the difference in means. Thus, the calculated mean values were the logarithms of the geometric means of the original values. Since the parameter differential blood count is not normally distributed, a non-parametric 95% CI for the difference in medians (Hodges – Lehmann estimator) was calculated.9 Calculations were not corrected for multiple testing, which means multifold application of statistical tests leading to an inflation of the type I error.

Results Clinical observations and body weights All the rats appeared to be healthy throughout the study period. Their mean body weight on arrival was 387 + 16 g in Group 1 and 388 + 16 g in Group 2. On the day of blood sampling, rats of both groups had nearly identical body weight: the mean body weight was 409 + 20 g in Group 1 and 409 + 17 g in Group 2.

differences in haematology parameters. The smallest differences occurred in the RBC system. Comparison of the variation of the measured values revealed no major differences between the two sampling methods. Comparison of the differential WBC count revealed that, consistent with the equivalence of WBC count, the distribution of most of the VC values was also equivalent to that of the VS values. However, for basophils there was a remarkable difference in the medians for VC and VS blood, the VC median being 50% lower than the VS median. Therefore, no equivalence was established for this parameter. Variation between the two groups was similar. The results of the differential blood count measurements including historical data are presented in Table 2. In addition, box-and-whisker plots of the differential WBC count are depicted in Figure 1. Coagulation The results for coagulation including the historical data are presented in Table 3. Mean APTT values were 30.3% lower in venous blood from the VC than in VS blood. Mean PT values were also lower in blood from the VC compared with the respective VS value, although the difference was not as pronounced as with APTT. No equivalence was found for the clotting values and no major differences in variability between the blood samples were detected. Clinical chemistry The results for the clinical chemistry parameters including historical data are presented in Table 4. Equivalence between the results of the two methods was established for ALT, AST, BUN and Ca2þ. No equivalence was found for GGT, AP, BIL, CHOL, TRIG, GLU, CREA, PO, PR, ALB, Naþ, Kþ, Cl2 and CO2. The largest differences in mean values for these parameters were observed for GGT, BIL and GLU; the smallest differences were measured for Naþ and Cl2 as well as for ALB and PR. Coefficients of variation differences between the two methods greater than factor 1.5 were detected for ALT (1.9-fold), GLU (1.7-fold), Kþ (1.7-fold) and Ca2þ (2.3-fold). The coefficients of these parameters were higher in VC blood. For all other clinical chemistry parameters, the differences in the coefficients of variation were below factor 1.5.

Haematology All the haematology values (means, SD, percentage difference of means and 95% CI for difference of means) including historical data from our facility except for differential WBC count are presented in Table 1. Equivalence was established for the following parameters: MCV, RDW, MEDRET, WBC and MPV. No equivalence was found for RBC, HGB, HCT, MCH, MCHC, ARETIC, LOWRET, HIGHRET and PLT. The ARETIC and HIGHRET values displayed the largest

Discussion Comparison of blood samples from the inferior VC with those from the VS revealed equivalence for 13 of 39 clinical pathology parameters (MCV, RDW, MEDRET, WBC, MPV, ABSNEUT, ABSLYMP, ABSMONO, ABSEOS, ALT, AST, BUN and Ca2þ). Our results confirm previous reports which have shown that different methods of blood sampling in laboratory rats may produce relevant differences in the mean values

9.9 + 2.46 1184 + 197.8 8.5 + 0.52

109/L 109/L fL 24.9 16.7 6.1

37.3

8.6

3.9 18.0 14.3

2.1

3.7 3.7 3.3 2.9 3.3

CV

10.4 + 2.22 1022 + 184.0 8.6 + 0.63

13.3 + 5.59

34.8 + 3.25

12.4 + 0.55 231.2 + 42.31 51.9 + 6.55

22.7 + 0.29

8.35 + 0.331 9.9 + 0.35 0.437 + 0.0155 52.4 + 1.48 1.19 + 0.036

Mean + SD

VS (n 5 23)

21.3 18.0 7.3

42.2

9.3

4.2 18.3 12.6

1.3

4.0 3.5 3.6 2.8 3.0

CV D

20.5 162 20.1

2.8

0.8

20.2 230.7 23.6

20.3

20.23 20.5 20.015 20.4 20.02

25.0 15.9 21.2

20.5

2.3

21.1 213.3 26.8

21.1

22.9 24.4 23.4 20.7 21.7

D% 20.05 20.3 20.006 0.5 0.00

21.9, 0.9 49, 276 20.5, 0.2

20.7, 6.2

21.1, 2.7

20.5, 0.1 254.05, 27.26 27.6, 0.4

20.5, 20.0

20.42, 20.7, 20.024, 21.2, 20.04,

95% CI difference of means

+2.3 +191 +0.6

+5.8

+3.2

+0.5 +39.4 +6.7

+0.4

+0.32 +0.4 +0.014 +1.5 +0.04

Equiv. limit

Yes No Yes

No

Yes

Yes No No

No

No No No Yes No

Equiv.

11.5 + 2.80 1013 + 176.7 8.0 + 0.47

11.2 + 3.42

34.6 + 2.51

12.9 + 0.71 233.4 + 38.39 54.2 + 4.95

24.3 + 0.40

8.39 + 0.389 9.8 + 0.42 0.404 + 0.0164 53.0 + 2.20 1.29 + 0.059

Mean + SD

Historical data (VS, n 5 60)

SD ¼ standard deviation; CV ¼ coefficient of variation; CI ¼ confidence interval; D ¼ absolute mean value difference between the vena cava (VC) and vena sublingualis (VS); D% ¼ percentage mean value difference between VC and VS. Historical data are also shown for comparison

16.0 + 5.98

12.3 + 0.48 200.5 + 36.19 48.4 + 6.94

% 109/L %

%

22.4 + 0.47

mmol/L

35.6 + 3.08

8.11 + 0.300 9.5 + 0.35 0.422 + 0.0140 52.0 + 1.52 1.17 + 0.038

1012/L mmol/L L/L fL fmol

%

Mean + SD

Unit

Parameter

Red blood cell count (RBC) Haemoglobin (HGB) Haematocrit (HCT) Mean corpuscular volume (MCV) Mean corpuscular haemoglobin (MCH) Mean corpuscular haemoglobin concentration (MCHC) Red cell distribution width (RDW) Absolute reticulocytes (ARETIC) Percent low RNA reticulocyte ratio (LOWRET) Percent middle RNA reticulocyte ratio (MEDRET) Percent high RNA reticulocyte ratio (HIGHRET) White blood cell count (WBC) Platelet count (PLT) Mean platelet volume (MPV)

VC (n 5 23)

Table 1 Effect of sampling method on haematology parameters

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31.8 26.9 38.8 51.7 47.7 (1.19 – 1.68) (6.61 – 9.16) (0.13 – 0.22) (0.09 – 0.21) (0.01 – 0.03) neutrophils (ABSNEUT) lymphocytes (ABSLYMP) monocytes (ABSMONO) esoinophils (ABSEOS) basophils (ABSBASO)

CV ¼ coefficient of variation; CI ¼ confidence interval; D ¼ Hodges Lehmann estimator of difference in medians; D% ¼ percentage median value difference between the vena cava (VC) and vena sublingualis (VS); Q1 ¼ 25% quartile; Q3 ¼ 75% quartile. Historical data are also shown for comparison

(1.05 –1.65) (8.40 –10.58) (0.16 –0.27) (0.08 –0.13) (0.04 –0.07)

Median (Q1– Q3)

1.30 9.55 0.20 0.10 0.05 Yes Yes Yes Yes No

Equiv. Equiv. limit

+0.43 +2.03 +0.07 +0.1 +0.02 0.23 0.67 0 0.04 20.01 20.28, 21.67, 20.07, 20.06, 20.03, 24.7 27.3 223.8 6.7 250

D% D

20.04 20.54 20.03 20.01 20.02 109/L 109/L 109/L 109/L 109/L

1.48 8.27 0.21 0.15 0.04 1.41 7.67 0.16 0.16 0.02

Unit Parameter

Absolute Absolute Absolute Absolute Absolute

CV

Median (Q1– Q3) Median (Q1 –Q3)

CV

26.1 22.5 36.1 58.6 53.8

95% CI difference of medians VS (n 5 23) VC (n 5 23)

Table 2 Effect of sampling method on the differential white blood cell (WBC) count

(1.18 –1.69) (6.95 –9.56) (0.16 –0.24) (0.1 –0.27) (0.02 –0.05)


................................................................................................................................................

of several clinical pathology parameters such as APTT, PLT, RBC, HGB, GLU, ALB and CREA.10 – 14 However, whereas these studies specifically investigated the differences resulting from various blood sampling techniques, our aim was to evaluate the equivalence of blood sampled from the VC immediately before necropsy and that from the VS in order to determine whether VS sampling could be replaced by VC sampling. The concept of testing for equivalence was originally developed for clinical trials to investigate the bioequivalence of generic pharmaceutical products. In analogy to these studies, this concept was chosen for the present study in order to evaluate the comparability of the two blood sampling techniques. VC sampling immediately before necropsy is less stressful to the rats as it eliminates the need to collect blood one day before necropsy while avoiding additional animal handling and anaesthesia. Despite finding that equivalence was not demonstrated for several parameters in this study, there was no clear trend towards higher or lower mean and median values in the VC samples compared with the mean values obtained by VS sampling, although the VC data did tend to yield slightly lower mean values. The most extreme differences in mean values were measured for GGT (275.0%) and HIGHRET (þ20.5%) in the VC group versus the VS group. Comparing the coefficients of variation revealed no general differences between the two blood sampling methods. Only for the clinical chemistry parameters ALT, GLU, Kþ and Ca2þ was variation approximately 1.7– 2.3 times higher in VC blood than in VS blood (Table 4). The reason is not known but these differences are not considered to be of biological relevance. Although the mean values obtained by both methods cannot be considered equivalent for most clinical pathology parameters as defined by the selected test criteria, careful evaluation of the data from the present study as well as their comparison with our historical laboratory data (n ¼ 60) led to the conclusion that, even though equivalence was not demonstrated, there was considerable similarity between the two groups with the exception of APTT. The notable difference in APTT mean values (230.3% lower in VC blood; see Table 3) was considered to be biologically relevant. An explanation for the prolonged APTT in samples taken from the VS may represent a local decrease in plasma coagulation subsequent to activation in traumatized tongue tissue, sample activation, coagulation factor consumption or a release of anticoagulant factors as proposed by Dameron et al. 12 and Salemink et al. 15 Alternatively, incision of the abdominal cavity might have activated the coagulation cascade prior to blood withdrawal, resulting in a decrease in APTT values. The VC APTT values obtained in the present study are in line with the values published by Salemink et al. 15 The VS values differ notably from our historical data. This difference may lie in a change in the procedure for handling the blood used for blood clotting analysis in our laboratory. Summing up, comparison of VS blood sampled prior to necropsy with blood sampled from the VC immediately before necropsy in male Sprague – Dawley rats revealed similarity in the clinical pathology parameters


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Figure 1 Comparison of the differential white blood cell (WBC) count of blood from the vena cava (VC) and vena sublingualis (VS): (a) absolute neutrophils, (b) absolute lymphocytes, (c) absolute monocytes, (d) absolute esoinophils, (e) absolute basophils and (f ) WBC. Numbers given on y axis are multiples of 109/L cells. The box-and-whisker-plots show median values (white line), 25% and 75% quartiles (box range), adjacent values (whiskers), and extreme values (dots)

higher APTT values in the VS samples. However, in a standard toxicological safety evaluation this effect would be irrelevant due to the presence of control data obtained by the same blood sampling method. In view of the results of this study, terminal blood sampling from VC during necropsy appears to be a feasible,

(haematology, clinical chemistry and coagulation). The differences between the two methods were within the normal range for Sprague –Dawley rats of this strain and age. This view is supported by a similar variation for most parameters and their consistency with our own historical data. The only relevant exception was the consistently

Table 3 Effect of sampling method on coagulation parameters VS (n 5 23)

VC (n 5 23)


Equiv. limit


Parameter

Unit

Mean + SD

CV

Mean + SD

CV

D

D%

Equiv.

Mean + SD

Prothrombin time (PT) Activated partial thromboplastin time (APTT)

s

39.98 + 3.421

8.56

42.74 + 2.376

5.56

22.76

26.5

24.5, 21.0

+2.95

No

33.18 + 2.218

s

16.6 + 0.89

5.4

23.8 + 1.58

6.6

27.2

230.3

27.97, 6.45

+1.3

No

15.0 + 0.83


Alanine aminotransferase (ALT) logALT Aspartate aminotransferase (AST) logAST g-Glutamyltransferase (GGT) Alkaline phosphatase (AP) Total bilirubin (BIL) Cholesterol (CHOL) Triglycerides (TRIG) Glucose (GLU) Creatinine (CREA) Inorganic phosphorus (PO) Blood urea nitrogen (BUN) Total protein (PR) Albumin (ALB) Sodium (Naþ) Potassium (Kþ) Calcium (Ca2þ) Chloride (Cl2) Carbon dioxide (CO2)

47.6 + 13.73 3.8 + 0.22 100.9 + 19.12 4.6 + 0.17 0.1 + 0.17 140 + 23.3 1.69 + 0.514 1.62 + 0.388 0.49 + 0.157 6.95 + 1.191 24.5 + 4.25 2.84 + 0.302 7.44 + 1.211 62 + 3.1 36.1 + 1.96 145 + 2.2 5.75 + 0.790 2.48 + 0.172 103 + 1.7 30.1 + 2.62

Mean + SD 28.8 5.7 19.0 3.8 189.5 16.7 30.4 24.0 32.3 17.1 17.3 10.6 16.3 4.9 5.4 1.5 13.7 7.0 1.6 8.7

CV 46.5 + 7.1 3.8 + 0.15 99.8 + 21.51 4.6 + 0.19 0.4 + 0.49 157 + 31.9 2.29 + 0.487 1.93 + 0.313 0.55 + 0.135 5.96 + 0.602 28.6 + 4.44 2.51 + 0.199 7.23 + 1.019 65.4 + 2.53 38.1 + 1.30 144 + 2.0 5.2 + 0.42 2.53 + 0.075 102 + 1.6 28.1 + 1.96

Mean + SD 15.2 3.8 21.6 4.2 124.9 20.3 21.3 16.2 24.7 10.1 15.5 7.6 14.1 3.9 3.4 1.4 8.1 3.0 1.6 7.0

CV D 1.1 0 1.1 0 20.3 218 20.6 20.31 20.06 1.0 24.1 0.33 0.21 23.4 22.0 1 0.55 20.05 1 1.9

2.3 0 1.1 0 275.0 210.8 226.2 216.1 210.9 16.6 214.3 13.1 2.9 25.2 25.2 0.7 10.6 22.0 1.0 7.1

D% N/A 2 0.1, N/A 2 0.1, 20.5, 234, 20.89, 20.52, 20.149, 0.44, 26.7, 0.18, 20.46, 25, 22.9, 20.1, 0.17, 20.13, 0.4, 0.5, 0.1 20.1 21 20.3 20.11 0.03 1.56 21.5 0.48 0.88 22 21.0 2.4 0.93 0.03 2.3 3.3

0.1


N/A +0.2 N/A +0.2 +0.4 +28 +0.5 +0.35 +0.147 +0.94 +4.3 +0.25 +1.12 +3 +1.7 +2.1 +0.66 +0.13 +1.7 +2.3

Equiv. limit

No No No No No No No No Yes No No No No Yes No No

Yes

Yes

Equiv.

48.3 + 8.20 3.9 + 0.17 108.8 + 23.85 4.7 + 0.20 0.6 + 0.61 172.9 + 36.61 2.47 + 0.673 1.86 + 0.336 0.85 + 0.311 5.16 + 0.968 28.8 + 4.53 2.88 + 0.207 5.65 + 0.939 62.0 + 2.67 36.1 + 1.37 140.9 + 2.10 5.56 + 0.662 2.57 + 0.063 99.3 + 1.43 24.76 + 2.129

Mean + SD



U/L U/L mmol/L mmol/L mmol/L mmol/L mmol/L mmol/L mmol/L g/L g/L mmol/L mmol/L mmol/L mmol/L mmol/L

U/L

Unit

U/L

Parameter

VS (n 5 23)

Volume 44

VC (n 5 23)

................................................................................................................................................

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Table 4 Effect of sampling method on blood chemistry parameters

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reliable, less painful and less stressful method of collecting terminal blood samples in preclinical toxicity studies conducted in rats.

ACKNOWLEDGEMENTS

The authors would like to thank Gerhard Marsch, Maren Fischer, Ute Dressler and Markus Woerner for the technical study conduct. They would also like to thank Dr Lisa Rehm (DVM) for her support in veterinary matters, Peter Gruener for handling the graphics and Bridget Grout for proofreading the manuscript.

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(Accepted 10 May 2010)

Laboratory Animals

Laboratory Animals

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