The stability of ionized calcium (CaI) concentration and pH in sera (n = 14) stored at 23 or 4 C for 6, 9, 12, 24, 48, or 72 hours, or -10 C for 1, 3, 7, 14, or 30 days was evaluated. Also studied were the effects of oxygen exposure, cold handling, and feeding on CaI and pH values. Results indicated that serum CaI concentration was stable throughout 72 hours of storage at 23 or 4 C, and for 7 days at -10 C. Serum CaI concentration significantly (P < 0.05) decreased by 14 days of storage at -10 C. Serum pH was stable for 6 hours at 23 or 4 C, and for 24 hours at -10 C, but significantly (P < 0.05) increased by 9 hours of storage at 23 or 4 C and by 3 days at -10 C. Exposure of the surface of the serum to air immediately before measurement had no effect on CaI or pH values, but mixing serum with air resulted in significantly (P < 0.05) decreased CaI concentration and increased pH. Handling of blood on ice resulted in significantly (P < 0.05) higher serum pH, compared with blood handled at 23 C, but serum CaI concentration was unaffected. Serum obtained at 2 hours after feeding did not have any significant changes in CaI, total calcium, or pH values. It appears that if canine serum is obtained, handled, and stored anaerobically, CaI concentration can be accurately measured after 72 hours at 23 or 4 C, or after 7 days at -10 C.

A commercially available radioimmunoassay kit for measurement of human osteocalcin was validated for use in horses. For accurate measurement of equine serum osteocalcin, blood samples may be collected at a temperature between 20 and 25 C, then centrifuged within 90 minutes; serum may be stored at - 20 C in plastic tubes for up to 26 weeks. Serum may be thawed and refrozen up to 5 times without significant change in measured equine serum osteocalcin concentration. Assay sensitivity was 0.16 ng/ ml. Recovery of bovine osteocalcin standard added to equine serum was linear. Intra-assay coefficient of variation (x 100) for 2 equine serum pools was 6.9 (mean +/- SD, 13.9 +/- 1.0 ng/ml) and 7.5 (10.6 +/- 0.8 ng/ml) %. Interassay coefficient of variation for 3 equine serum pools measured in 12 assays was 12.5 (16.1 +/- 2.0 ng/ml), 12.7 (11.5 +/- 1.5 ng/ml), and 24.6 (3.0 +/- 0.7 ng/ml) %. Dilutional parallelism was documented by assaying pooled equine serum at 4 dilutions and correcting the mean result for dilution. Significant change was not observed in equine serum osteocalcin concentration for various time-of-day blood sample collections in horses housed under continuous lighting.

OBJECTIVE To evaluate the effect of presurgical storage conditions on leakage pressures of enterotomy sites closed with unidirectional barbed suture material in fresh, chilled, and frozen-thawed cadaveric canine jejunal specimens. SAMPLE 36 grossly normal jejunal segments obtained from 4 dog cadavers. PROCEDURES 9 jejunal segments were harvested immediately from each euthanized dog and randomly assigned to be tested within 4 hours after collection (fresh segments), stored at 4°C for 24 hours before testing (chilled segments), or stored at −20°C for 7 days and thawed at 21°C for 6 hours before testing (frozen-thawed segments). For leakage pressure testing, a 3-cm-long antimesenteric enterotomy was performed and repaired with 3-0 unidirectional barbed suture material in a simple continuous pattern in each segment. Time to complete the enterotomy, initial leakage pressure, maximum intraluminal pressure, and leakage location were recorded for each segment. RESULTS Mean ± SD initial leakage pressure for fresh, chilled, and frozen-thawed segments was 52.8 ± 14.9 mm Hg, 51.8 ± 11.9 mm Hg, and 33.3 ± 7.7 mm Hg, respectively. Frozen-thawed segments had significantly lower mean initial leakage pressure, compared with findings for fresh or chilled segments. Time to complete the enterotomy, maximum intraluminal pressure, and leakage location did not differ among groups. CONCLUSIONS AND CLINICAL RELEVANCE Leak pressure testing of cadaveric jejunal segments that are fresh or chilled at 4°C for 24 hours is recommended for enterotomy studies involving barbed suture material in dogs. Freezing and thawing of cadaveric jejunal tissues prior to investigative use is not recommended because leak pressure data may be falsely low.

The objective of this study was to assess whether concentrations of epidermal growth factor (EGF), fibronectin, and alpha(α)-2-macroglobulin in canine serum remain stable under different storage conditions. Serum was obtained from 10 adult dogs and stored for 7 d at room temperature (RT) and at 4°C and for 1, 3, and 6 mo at -20°C. Bacterial cultures of serum were carried out after 7 d at 4°C and at RT. For each dog and time point, EGF, fibronectin, and α-2-macroglobulin were quantified in duplicate by enzyme-linked immunosorbent assay (ELISA). Mean concentrations of each factor at each time point were used for statistical analysis. No bacterial growth was observed in any samples. Compared to baseline (232.24 ± 49.47 pg/mL), EGF concentration was significantly lower after 1 wk of storage at 4°C (135.39 ± 27.12 pg/mL, P = 0.006), but not at RT (315.85 ± 79 pg/mL, P = 0.6) or after 1, 3, or 6 mo of storage at -20°C (220.84 ± 41.07 pg/mL, P = 0.7; 220.98 ± 78.26 pg/mL, P = 0.8; 266.06 ± 20.39 pg/mL, P = 0.4, respectively). Compared to baseline, concentrations of fibronectin after 1 wk of storage at 4°C or at RT and 1, 3, or 6 mo of storage at -20°C were not statistically different. Compared to baseline (186.67 ± 45.20 mg/dL), the concentration of α-2-macroglobulin after 1 wk of storage at 4°C was significantly increased (244.61 ± 58.27 mg/dL, P = 0.002), but not at RT (177.09 ± 26.99 mg/dL, P = 0.2). The differences in concentration after 3 and 6 mo of storage at -20°C were significant compared to baseline (243.32 ± 42.64 mg/dL, P = 0.005 and 56.39 ± 21.78 mg/dL, P < 0.0001, respectively), but not after 1 mo of storage at -20°C (136.79 ± 25.61 mg/dL, P = 0.1). One week of storage at RT has little effect on the stability of EGF, fibronectin, and α-2-macroglobulin in canine serum. Measured factors remain stable for 3 mo of storage at -20°C.

Eleven pairs of canine metacarpal bones, 10 pairs of metatarsal bones, and 7 pairs of ribs were harvested cleanly and prepared for banking at -20 C for 1 year. One bone of each pair was randomly assigned to 1 type of storage: plastic pack vs immersion in a normal solution of sodium chloride. The contralateral bone was assigned to the opposite treatment. Six pairs of metacarpal bones and 5 pairs of metatarsal bones were tested in torsion to failure. No significant difference was found within pairs. All ribs, 5 pairs of metacarpal bones, and 5 pairs of metatarsal bones were loaded in 4-point bending to failure. The energy absorbed at failure and the ultimate displacement of ribs and metacarpal and metatarsal bones were increased by 25 to 30% and 18 to 24%, respectively, when the bones were frozen in isotonic saline solution. Corticocancellous grafts frozen in normal saline solution are biomechanically less fragile and brittle than grafts stored in plastic without saline solution.

Effects of temperature and storage time on canine bone-transfixation pin specimens were tested by comparing pin pull-out forces. A total of 16 femurs from 8 mature dogs were tested. Five nonthreaded Steinmann pins were placed through both cortices in the diaphysis of each femur. The femurs were then sectioned transversely between each pin, with a bonepin specimen placed evenly into each of 5 groups prior to biomechanical testing. Four bone-pin specimen groups were stored at -20 or -70 C for 14 or 28 days, while 1 specimen group was immediately tested. Pull-out forces for frozen groups were compared with pull-out forces for the fresh group. Using two-way ANOVA, there was no statistical difference in mean axial-extraction forces among bonepin specimen in any of the tested groups. It is concluded that acute pin pull-out forces are not significantly affected by freezing temperature or time. However, specimens stored at -20 C for as few as 14 days had a trend for increased pull-out forces, compared with freshly harvested specimens. Therefore, the authors recommend storage of bone-pin specimens at -70 C when possible.

Eight Holstein cows, 4 inoculated intracisternally in 1 quarter of the mammary gland with Escherichia coli and 4 noninfected controls, were administered ceftiofur sodium (3 mg/kg of body weight, IV, q 12 hours) for 24 hours, beginning at 14 hours after inoculation of infected cows. All challenge-exposed cows became infected, with mean +/- SEM peak log10 bacterial concentration in milk of 5.03 +/- 0.69 colony-forming units/ml. The infection resulted in systemic signs (mean peak rectal temperature, 41.5 +/- 0.3 C; anorexia; signs of depression) and local inflammation (mean peak albumin concentration in milk, 7.89 +/- 1.71 mg/ml). Ceftiofur was detectable in milk from all challenge-exposed cows, compared with only 1 of 4 noninfected cows, and the mean period after inoculation that ceftiofur was detectable in milk was longer (P < 0.05) in infected (147.7 +/- 27.5 hours) than noninfected cows (1.3 +/- 1.3 hours). However, maximal ceftiofur concentration attained in milk for all cows was 0.28 micrograms/ml, and was 0.20 micrograms/ml or less for all but 2 milk samples collected for 10 days after challenge exposure. Mean serum concentration of ceftiofur peaked at 1.0 +/- 0.3 micrograms/ml and 0.7 +/- 0.1 micrograms/ml for infected and noninfected COWS, respectively. After each ceftiofur dose, mean peak and trough concentrations of ceftiofur in serum did not differ between groups; however, concentration of ceftiofur in serum was higher at 7 hours after each dose in noninfected cows, suggesting more rapid clearance of the drug in infected cows. Ceftiofur was not detected in serum (< 0.05 micrograms/ml) of any cow at or after 120 hours following inoculation of infected cows. Storage of serum samples at -20 C for 3 weeks resulted in a 98.8% decrease in ceftiofur activity, compared with that in fresh serum samples. Eighty-seven percent of this loss occurred 30 minutes after mixing serum and ceftiofur; thus, about 13% of the original activity was lost in storage. Storage of milk samples under similar conditions did not result in loss of ceftiofur activity. Despite acute inflammation, the dosage of ceftiofur used in this trial would not result in drug concentrations in milk above FDA safe concentrations, or above the reported minimum inhibitory concentration for coliform bacteria.

A commercially available radioimmunoassay (RIA) kit for measurement of human adrenocorticotropin (hACTH) was validated for use in dogs. Assay sensitivity was 3 pg/ml. Intra-assay coefficient of variation (X 100; CV) for 3 canine plasma pools was 3.0 (mean +/- SD, 33 +/- 0.99 pg/ml), 4.2 (71 +/- 2.4 pg/ml) and 3.7 (145 +/- 3.7 pg/ml) %. Interassay CV for 2 plasma pools measured in 6 assays was 9.8 (37 +/- 3.6 pg/ml) and 4.4 (76 +/- 3.4 pg/ml) %, respectively. Dilutional parallelism was documented by assaying 2 pools of canine plasma at 3 dilutions and correcting the measured result for dilution. Corrected mean concentrations for the first pool were 33 (+/- 0.99), 36 (+/- 4.3), and 33 (+/- 6.8) pg/ml; corrected mean concentrations for the second pool were 145 (+/- 5.4), 141 (+/- 10.8) and 125 (+/- 3.4) pg/ml. Recovery of 1-39hACTH added to canine plasma (6.25, 12.5, 25.0, 50.0, and 100.0 pg/ml) was linear and quantitative (slope = 0.890, R2 = 0.961). To test whether anticoagulant or the protease inhibitor, aprotinin, influences ACTH concentration in canine plasma, ACTH was measured in canine blood collected in 4 tubes containing anticoagulant: heparin (H), heparin + 500 kallikrein inhibitor units (KIU) of aprotinin/ml (HA), EDTA (E), and EDTA + aprotinin (EA). Plasma ACTH concentration was the same when samples containing H and HA, or HA and E were compared, and was significantly (P < 0.01) lower in samples containing EA. Plasma storage at -20 C for 1 week or 1 month was not associated with significant change in ACTH concentration in canine plasma, using any of the 4 anticoagulant treatments. Plasma ACTH concentration measured after 6 months' storage at -20 C was significantly (P < 0.01) lower for all anticoagulants used. Synthetic 1-39hACTH added to canine blood was accurately recovered (88 to 109%, n = 3) from plasma containing EDTA, with or without aprotinin, whereas percentage recovery was overestimated by 18 to 91% in heparinized plasma. Plasma ACTH concentrations in EDTA-treated canine blood kept at 4 or 22 to 25 C for 15 to 90 minutes prior to centrifugation at 8 C were not significantly different. Plasma ACTH concentration in canine plasma was affected by storage tube material. Concentration of ACTH in canine plasma stored in borosilicate glass tubes for 1 week or 1 month at -70 C was significantly higher than initial ACTH values (P less than or equal to 0.01), but was unchanged over time in plasma stored in polypropylene or polystyrene tubes. Sample handling procedures affect canine plasma ACTH concentration measured by use of the RIA kit. Optimal sample handling conditions for plasma ACTH measurement in dogs include use of EDTA anticoagulant, blood collected at 20 to 25 C (room temperature) followed by centrifugation within 15 to 90 minutes, and plasma storage in plastic (not glass) tubes for not longer than 1 month at -20 C.

This study was conducted to determine the incidence of microorganisms associated with dressed broiler with intact skin and without skin at different frozen storage periods such as 0, 10, 20, 30 days and to demonstrate the role of packaging and pretreatment chilling on the changes of carcass quality. The values of total viable count (TVC), total coliform count (TCC), total streptococcal count (TStC) and total staphylococcal count (TSC) were determined for meat samples of thigh and breast and swab samples of visceral surfaces of the broilers with intact skin and without skin.