2020 年 57 巻 3 号 p. 241-245
Glucose is a major circulating carbohydrate in birds and its level in the blood is often used as a biometric indicator in clinical diagnosis and various studies. Notably, hypoglycemia is often associated with Spiking Mortality Syndrome in broilers; therefore, blood glucose levels need to be correctly evaluated in clinical diagnosis. In the present study, we investigated the effect of different blood treatment methods after blood collection on chicken blood glucose measurements. The blood glucose level of plasma separated from blood cell components immediately after blood collection was used as a reference and compared with glucose levels in serum and stored plasma. The mean glucose level in plasma separated from blood cell components immediately after blood collection was 236.1±15.9 mg/dL and remained stable for at least one week in refrigerated storage (between 2°C and 5°C). However, glucose levels decreased slowly in plasma unseparated from blood cell components in storage with ice water. Mean glucose level in serum separated from blood cell components 1 h after blood collection was 206.4±9.2 mg/dL and fell to 108.3±30.0 mg/dL after 24 h. Therefore, the chicken blood serum glucose level was significantly lower than the level in plasma immediately after blood collection, regardless of elapsed time after blood collection. For the measurement of glucose in chicken blood, it is necessary to use refrigeration, use plasma from which blood cell components have been removed, and take measurements within at least 30 min.
Blood glucose concentration is often used as a biological evaluation index in studies that evaluate the effects of dietary components on rearing conditions and health abnormalities. Moreover, in research involving blood glucose, measurements have been performed using whole blood (Burns et al., 2002; Bafundo et al., 2018), plasma (Smith and Baranowski-Kish, 1976), or serum (Brake et al., 1981; Daly and Peterson, 1990; Latour et al., 1996; Peebles et al., 2012; Sahin et al., 2018), and it is difficult to find consistency from blood collection techniques to measurement methods and results. Glucose is the main carbohydrate in circulation in birds and its concentration is controlled by pancreatic hormones, glucagon, and insulin (Scanes, 2015). Typical avian blood glucose levels fall within a range higher than is normal for mammals and are almost unaffected by dietary intake (Stevens, 1996). In broilers, Spiking Mortality Syndrome (SMS) is associated with severe hypoglycemia. In the diagnosis of this syndrome, an abnormally low blood glucose level is an important indicator in addition to typical clinical findings (Davis, 2013). The measured value of glucose in blood biochemical tests is affected by the processing method and processing time after blood collection. After blood collection, glucose in the blood is consumed rapidly by blood cells, mainly for glycolysis (Chan et al., 1989). To avoid this effect, it is necessary to examine the blood immediately after collection and to separate the cell components from the liquid component. However, when blood is collected at the site where the chicken is raised, it is not only difficult to carry out the test immediately, but in most cases, it is inevitable that the post-treatment process requires a certain amount of time. The glucose value measured in blood immediately after blood collection is lower than that measured using plasma and is affected by the hematocrit value (Kim, 2016). Serum is the liquid component of blood that remains after the cell components have coagulated. In our clinical experiences, for chicken blood samples, the time required for serum to separate out sufficiently exceeds 1 h at ambient temperature (15–35°C). Therefore, when glucose is measured in serum, there is a possibility that levels are lower than those measured in blood and plasma.
In this study, we examined the effect of processing and the elapsed time after blood collection on glucose measurements in chicken blood.
In this study, animal handling and sampling were conducted in accordance with the Japan Veterinary Medical Association guidelines for industrial animal medicine.
Blood samples (3–4 mL) were collected individually from twenty-four 41-day-old broiler chickens (ROSS308) via the wing vein. Each blood sample was immediately divided by dispensing half into a Spitz tube that contained lithium heparin and the other half into a plain Spitz tube. Eleven of the lithium heparin samples were immediately gently agitated, centrifuged at 1200 × g for 10 min, and 0.2 mL of separated plasma from each blood sample was placed in a microtube and chilled with iced water (0 h plasma). The same Spitz tubes (containing the remaining plasma plus blood cell components) were cooled with iced water and 0.2 mL samples of the plasma that had separated from the blood cells were removed at 1 h, 9 h, and 24 h after blood collection and transferred to microtubes (1 h/9h/24h plasma). The 11 blood samples in plain Spitz tubes were incubated in a water bath at 37°C for 1 h and 0.2 mL of each separated serum was placed in a microtube and cooled with iced water (1 h serum). The blood samples that remained in the Spitz tubes were allowed to stand at room temperature (24–28°C), which allowed serum production to continue and the blood clot to become sufficiently retracted. After 9 h and 24 h, serum samples (0.2 mL at each time point) were transferred into microtubes and cooled with iced water (9 h/24 h serum). The plasma and the serum collected in microtubes at each time point were measured for glucose on the day of collection. The 0 h plasma was refrigerated (between 2°C and 5°C ) for 7 d after blood collection and the plasma glucose was measured (0 h+7 d:rfg plasma). The refrigerated 0 h plasma was frozen at −30°C for 23 d and thawed at room temperature (23–25°C). After thorough shaking and stirring, the plasma glucose level was measured (0h+7 d:rfg+23d:fz plasma).
Plasma and serum obtained by these treatments were measured for glucose with a dry clinical chemistry analyzer (Spotchem D-00 and D-02, Arkray, Kyoto).
Comparative Analysis of Glucose MeasurementsThe glucose levels for 1 h, 9 h, and 24 h plasma were compared with those of 0 h plasma to investigate changes over time for plasma stored with iced water without separation from blood cell components. In addition, to investigate the effect of storage by refrigeration and freezing, the glucose levels for 0h+7 d:rfg plasma and 0h+7 d:rfg+23d:fz plasma were compared with those of 0 h plasma. To track changes in glucose levels over time for separated serum at room temperature, measurements in 1 h, 9 h, and 24 h serum were compared with those in 0 h plasma.
The statistical analysis in these comparisons used the Wilcoxon signed-rank test.
The glucose levels for 0 h, 1 h, 9 h, and 24 h plasma are shown in Table 1. The mean glucose level of plasma separated from blood cell components immediately after blood collection (0h plasma) was 236.1±15.9 mg/dL, whereas mean levels in 1 h, 9 h, and 24 h plasma were 229.9±11.1 mg/dL, 224.1±10.3 mg/dL, and 216.7±7.7 mg/dL, respectively. No significant difference was observed between 1 h and 0h plasma glucose levels (p≥0.05), but there were significant differences between 9 h and 24 h plasma glucose levels and 0h plasma (both p<0.01).
| Blood samples | 0h | 1h | 9h | 24h |
|---|---|---|---|---|
| mg/dL | ||||
| BL01 | 255 | 237 | 226 | 215 |
| BL02 | 252 | 243 | 243 | 231 |
| BL03 | 235 | 234 | 220 | 214 |
| BL04 | 236 | 223 | 216 | 213 |
| BL05 | 241 | 243 | 214 | 208 |
| BL06 | 253 | 242 | 242 | 225 |
| BL07 | 244 | 235 | 230 | 220 |
| BL08 | 233 | 222 | 222 | 221 |
| BL09 | 229 | 220 | 218 | 220 |
| BL10 | 204 | 214 | 214 | 204 |
| BL11 | 215 | 216 | 220 | 213 |
| Mean±S.D. | 236.1±15.9 | 229.9±11.1 | 224.1±10.3 | 216.7±7.7 |
| Max | 255 | 243 | 243 | 231 |
| Min | 204 | 214 | 214 | 204 |
| p-value | 0.0606 | 0.0059 | 0.0020 | |
p-value: Significance between 0 h plasma glucose level.
The glucose levels for 0h+7d:rfg plasma and 0h+7d:rfg+3d:fz plasma were 235.7±10.6 mg/dL and 226.3±10.0 mg/dL, respectively (Table 2), with no significant difference (p≥0.05). However, mean levels in 0h+7 d:rfg+23d:fz plasma were significantly lower than those in 0 h plasma (p<0.05).
| Blood samples | 0h | 0h+7 d:rfg | 0h+7 d: rfg+23d:fz |
|---|---|---|---|
| mg/dL | |||
| BL01 | 255 | 258 | 241 |
| BL02 | 252 | 240 | 242 |
| BL03 | 235 | 228 | 216 |
| BL04 | 236 | 226 | 235 |
| BL05 | 241 | 242 | 215 |
| BL06 | 253 | 245 | 229 |
| BL07 | 244 | 233 | 226 |
| BL08 | 233 | 228 | 224 |
| BL09 | 229 | 239 | 227 |
| BL10 | 204 | 220 | 212 |
| BL11 | 215 | 234 | 222 |
| Mean±S.D. | 236.1±15.9 | 235.7±10.6 | 226.3±10.0 |
| Max | 255 | 258 | 242 |
| Min | 204 | 220 | 212 |
| p-value | 0.8496 | 0.0186 | |
p-value: Significance between 0h plasma glucose level.
0h: Plasma of the day separated immediately after blood collection.
0h+7 d:rfg: 0h Plasma refrigerated for 7 days after blood collection.
0h+7 d:rfg+23d:fz: 0h Plasma stored for 23 days at −30°C after refrigeration for 7 days.
The glucose levels for 1 h, 9 h, and 24 h serum are shown in Table 3, with mean glucose levels of 206.4±9.2 mg/dL, 170.7±16.2 mg/dL, and 108.3±30.0 mg/dL, respectively, all significantly different from 0 h plasma (all p<0.01).
| Blood samples | 0h plasma | 1h | 9h | 24h |
|---|---|---|---|---|
| mg/dL | ||||
| BL01 | 255 | 208 | 163 | 131 |
| BL02 | 252 | 227 | 183 | 108 |
| BL03 | 235 | 199 | 174 | 134 |
| BL04 | 236 | 206 | 150 | 96 |
| BL05 | 241 | 202 | 189 | 94 |
| BL06 | 253 | 218 | 182 | 127 |
| BL07 | 244 | 209 | 192 | 128 |
| BL08 | 233 | 198 | 171 | 117 |
| BL09 | 229 | 196 | 153 | 63 |
| BL10 | 204 | 204 | 144 | 50 |
| BL11 | 215 | 203 | 177 | 143 |
| Mean±S.D. | 236.1±15.9 | 206.4±9.2 | 170.7±16.2 | 108.3±30.0 |
| Max | 255 | 227 | 192 | 143 |
| Min | 204 | 196 | 144 | 50 |
| p-value | 0.0020 | 0.0010 | 0.0010 | |
p-value: Significance between 0h plasma glucose level.
Avian blood glucose levels are higher than in mammals and steady-state levels are less likely to change, even under starvation (Stevens, 1996). On the other hand, an abnormal drop in blood glucose (hypoglycemia) occurs as one of the clinical signs of SMS in broilers (Brown et al., 1991; Davis et al., 1995; Burns et al., 2002; Davis, 2013). An accurate measurement of blood glucose level is essential for the clinical diagnosis of SMS. Sidebottom et al. (1982) suggested that blood samples taken for measuring blood glucose levels should be analyzed promptly; otherwise, the incidence of hypoglycemia might be overestimated. Human medical laboratory tests indicate that whole blood or plasma should be used in the diagnosis of diabetes, where blood glucose measurement results are particularly important clinically (Kim, 2016). However, it is still unclear how differences in post-collection treatment ofavian blood affect glucose levels. In general, a considerable amount of time is inevitably required from the time of blood collection in a poultry house until a biochemical test is performed. It may take several hours to catch chickens in a poultry house, collect blood, and transport the samples to a laboratory equipped with biochemical testing equipment. Consequently, for accurate results, it is important to know how differences in processing methods affect blood glucose levels. In this study, we evaluated the effect of various post-collection blood treatments on glucose levels, by comparing the results against plasma levels measured immediately after collection. Levels in plasma separated from blood cell components were stable for at least 1 week in refrigerated storage. However, after further, frozen, storage at −30°C, mean plasma glucose levels decreased by approximately 4%, which showed that once the plasma was frozen, glucose levels slightly decreased.
Serum is the fluid that remains after blood clots over time at room temperature. Empirically, when chicken blood is placed at room temperature, clot retraction generally progresses slowly and it may take several hours before serum is produced. In this experiment, to accelerate clot retraction, samples were incubated in a water bath (37°C) for 1 h immediately after blood collection to promote serum production. Even by this time, the mean glucose level had fallen to 206.4±9.2 mg/dL, significantly lower than the levels for 0h plasma (236.1±15.9 mg/dL; p<0.01), a decrease of over 10%. Glucose levels in serum produced at room temperature decreased further over time, to less than half the mean levels of 0 h plasma after 24 h. Therefore, we concluded that serum was not suitable for the measurement of chicken blood glucose. The time required for blood clotting in birds, including chickens, is considerably longer than that in mammals, requiring 38–124 min (Scanes, 2015) and the reduction in glucose concentration in human whole blood ex vivo averages 5%–7% per hour (Chan et al., 1989). The blood-clotting mechanism in birds is presumed to be the same as that in mammals, with prothrombin activated to thrombin by stimulation with surface contact or tissue damage (Scanes, 2015). In human erythrocytes, glucose degradation is promoted by phosphofructokinase activated by stimulation of inorganic phosphate in the blood (Tsuboi and Fukunaga, 1965). Additionally, thrombin and epinephrine induce platelet destruction secondary to aggregation and contraction and increase glucose uptake, thereby promoting glycolysis by hexokinase activity (Karpatkin, 1967). In birds, erythrocytes have phosphofructokinase activity, pyruvate kinase activity, and hexokinase activity and the glycolytic mechanism functions in the same way as human erythrocytes (Kalomenopoulou and Beis, 1990). The decrease in glucose levels in chicken plasma stored for 9–24 h with iced water without separation of blood cell components was less than that in serum and these levels slightly decreased compared with that in plasma separated from blood cell components immediately after blood collection followed by refrigeration. Blood glycolysis can be delayed by treatment with sodium fluoride but progresses to the same extent in blood treated with heparin for 1 h after treatment (Chan et al., 1989). In addition, it is necessary to exercise care when using sodium fluoride because hydrogen fluoride may be generated during the measurement of glucose level with the potential to damage the biochemical test apparatus.
In conclusion, when collecting blood to measure blood glucose, the optimal method is to immediately heparinize the sample, remove the blood cell components within at least 30 min after collection, and rapidly store the resulting plasma in a cooler box with iced water or a refrigerator. When collecting a small amount of blood in a poultry house, it is possible to separate plasma and blood cell components immediately after blood collection using a small battery-powered centrifuge. The results in this study were obtained using blood collected from apparently healthy chickens of a single breeding brand, breeding method, and blood collection age. Therefore, it is necessary to further investigate the relationship between different breeding brands, breeding methods, blood collection ages, or differences in the pathophysiological state of chickens and the effects of blood treatment on blood glucose levels and their mechanisms.
The authors declare no conflict of interest.
The authors would like to thank the poultry farm of Rakuno Gakuen cooperated with collect the blood samples.
This study was funded by KAWAVET LLC, the owner company of Research Office Concerning the Health of Humans and Birds.
