The influence of storage time and temperature on the measurement of serum,plasma and urine osmolality

Abstract

Background

Many clinical laboratories require that specimens for serum and urine osmolality determination be processed within 3 h of sampling or need to arrive at the laboratory on ice. This protocol is based on the World Health Organization report on sample storage and stability, but the recommendation lacks good supporting data. We studied the effect of storage temperature and time on osmolality measurements.

Methods

Blood and urine samples were obtained from 16 patients and 25 healthy volunteers. Baseline serum, plasma and urine osmolality measurements were performed within 30 min. Measurements were then made at 3, 6, 12, 24 and 36 h on samples stored at 4–8℃ and room temperature. We compared baseline values with subsequent measurements and used difference plots to illustrate changes in osmolality.

Results

At 4–8℃, serum and plasma osmolality were stable for up to 36 h. At room temperature, serum and plasma osmolality were very stable for up to 12 h. At 24 and 36 h, changes from baseline osmolality were statistically significant and exceeded the total allowable error of 1.5% but not the reference change value of 4.1%. Urine osmolality was extremely stable at room temperature with a mean change of less than 1 mosmol/kg at 36 h.

Conclusions

Serum and plasma samples can be stored at room temperature for up to 36 h before measuring osmolality. Cooling samples to 4–8℃ may be useful when delays in measurement beyond 12 h are anticipated. Urine osmolality is extremely stable for up to 36 h at room temperature.

Introduction

Serum and urine osmolality are frequently measured in clinical practice to assist with the diagnosis and management of polyuria, hypernatraemia and hyponatraemia. It is also applied in the calculation of serum or urine osmolal gaps and can be used in the estimation of urinary ammonium excretion. ¹ Specimens for serum and urine osmolality determination are often required to be processed within 3 h of sampling or must be transported and delivered to the laboratory on ice. This protocol has been in place in many laboratories since 2009 and is based on the World Health Organization (WHO) report on sample storage and stability. ² Implementation of this protocol results in the rejection of many samples with obvious cost implications and a negative impact on effective clinical decision making.

According to the WHO report, serum/plasma osmolality is stable for 3 h at 20 to 25℃, for 1 day at 4–8℃ and for three months at −20℃. It also states that urine osmolality is stable for 3 h at 20–25℃, for seven days at 4–8℃ and for more than three months at −20℃. The recommendations are based on a study by Zhang et al. ³ on the effects of prolonged serum-clot contact time on the determination of commonly measured analytes. They found that potassium and glucose concentrations had clinically relevant changes at 3 h and that phosphate concentration was elevated by 6 h. Sodium, calcium, urea and chloride concentrations were stable over 24 h. The WHO recommendations for plasma and serum osmolality stability are therefore based on the instability of analytes that contribute to osmolality, rather than on stability studies which measured osmolality directly. ³

Boyanton and Blick ⁴ demonstrated a decrease in glucose and HCO₃⁻ concentration and an increase in lactate and CO₂ concentrations when serum and plasma separation is delayed. These changes are attributed to on-going glycolysis by red blood cells in vitro. The formation of lactate during glycolysis yields H⁺ ions that are buffered by HCO₃⁻. HCO₃⁻ concentration consequently decreases and CO₂ concentration increases over time. Changes in the concentrations of HCO₃⁻ and CO₂ were shown to influence plasma osmolality by Boning and Maassen. ⁵ It is therefore reasonable to expect osmolality to change over time when separation from cellular components is delayed.

Osmolality stability studies where serum or plasma is separated from cellular components almost immediately after sampling and then stored have limited relevance for clinical practice where there are often delays in getting samples to the laboratory and where plasma or serum has not yet been separated.^6,7 The direct effect on osmolality of delaying serum separation was examined by Redetzki et al. ⁸ who showed that the osmolality of serum samples stored for 1–4 h at room temperature before separation was, on average, 2.8 mosmol/kg higher than that of baseline samples which were cooled immediately and measured within 20 min of sampling. They demonstrated that this difference was attributable to the formation of lactate. In another set of samples, they also demonstrated an average increase in serum osmolality of 3.5 mosmol/kg over 5 h at room temperature and 1.5 mosmol/kg at 0℃. The changes from baseline correlated very well with the changes in lactate concentration. The effect of storage temperature and delayed serum separation was, however, not studied beyond 5 h.

Most laboratories consider plasma and serum osmolality to be identical.^6,9 Serum is considered to be more stable than plasma for the measurement of many analytes, when there is prolonged contact with cells. In Boyanton and Blick’s stability study, ⁴ the changes seen over time in lactate, glucose, potassium, pH, CO₂ and HCO₃⁻ were more pronounced in plasma than in serum. This suggests that clotted serum samples might be the preferred medium for measuring osmolality when separation is delayed.

To the best of our knowledge, the only stability data available for urine osmolality is a white paper from an osmometer manufacturing company, which reported good stability over 24 h. ¹⁰ The WHO report refers to Cooper et al., ¹¹ but this study has no data on the stability of urinary analytes or osmolality. It is generally advised that urine specimens be refrigerated or chemically treated to prevent bacterial growth if samples are to be stored before analysis. Bacteria may cause conversion of glucose into acids and alcohols, and urea into ammonia. They also utilize glucose and ketones, causing a decrease in their concentration. For determining urine osmolality, refrigeration, but not the addition of preservatives, is advised. ¹²

In summary, guidelines for the handling of samples for osmolality lack good supporting data. The stability of chemical analytes contributing to serum and plasma osmolality has been studied well, but only one study directly investigated the effect of temperature and delayed serum separation on the determination of osmolality, and then only over a 5-h period. No studies have directly examined the stability of urine and plasma osmolality. In view of this knowledge gap, we investigated the effect of delayed serum and plasma separation, and cooling, on osmolality measurements. We also compared serum with plasma osmolality to determine the most stable medium. Last, we investigated the effect of storage time and temperature on the stability of urine osmolality.

Materials and methods

Assessment of the sample rejection rate based on current laboratory protocols

The sample rejection rate for serum and urine osmolality was determined by a search of the Tygerberg Hospital NHLS database for the period 1 January to 31 December 2012.

Participants and samples for osmolality measurements

We obtained a total of 30 blood samples and 40 urine samples over a period of six weeks. These were provided by 16 patients and 25 healthy volunteers. The patients were selected on the basis of having conditions such as hyperglycaemia, hyponatraemia and renal failure, which are known to affect osmolality. To ensure a good spread of urine osmolality, some volunteers were sampled after an overnight fast and others after taking a water load.

Specimen collection and handling

After obtaining written informed consent, 45–50 mL of blood was drawn from a forearm vein. The sample was immediately divided among 11 serum-separating tubes and 11 lithium heparin tubes (BD Vacutainer®, Becton, Dickinson South Africa, Gauteng, South Africa). Serum-separating tubes were each filled with 1 mL of blood and heparin lithium tubes with 3 mL of blood. One serum and one plasma tube were centrifuged, and osmolality was determined within 30 min to provide a baseline value for each medium. Five tubes from each medium were stored at 4–8℃ and the remaining five tubes at room temperature. Pairs of serum and plasma samples from the cooled and room temperature sets of samples were centrifuged and measured at 3, 6, 12, 24 and 36 h.

The stability of urine osmolality was assessed in a similar manner. A fresh urine sample was divided amongst 11 non-sterile, plastic 5 mL tubes containing no additives. A baseline sample was centrifuged and measured within 30 min. The remaining samples were split into a group of tubes stored at 4–8℃ and a group stored at room temperature. Samples from each group then had osmolality determinations at 3, 6, 12, 24 and 36 h.

Osmometry

All osmolality measurements were determined by freezing-point depression. Samples from the 16 patients were analysed on the Advanced Micro-Osmometer 3320 (Advanced Instruments, Inc., Norwood, MA, USA). The instrument has a within-day coefficient of variation (CV) of 0.9% for measurements below 400 mosmol/kg and a CV of 0.5% for measurements between 400 and 2000 mosmol/kg. Baseline measurements falling outside the 280–295 mosmol/kg reference intervals for serum/plasma osmolality were repeated as per the standard operating procedure in our laboratory. Measurements at later time points were repeated if they differed by more than 10 mosmol/kg from the baseline value to minimize random errors.

A Micro-Osmometer 3300 (Advanced Instruments, Incorporated Norwood, MA, USA) was used to measure the samples from the healthy volunteers. This instrument has a within-day CV of 0.87% for serum and 0.36% for urine. Each measurement was performed in duplicate. When measurements differed by more than 5 mosmol/kg, a third measurement was made. The average of the two closest values was taken to be the true value.

Statistical analysis

Repeated-measures analysis of variance was used to compare baseline values with subsequent measurements at each time point, to compare serum and plasma, and cooled and room temperature specimens. We used Bland–Altman difference plots to illustrate the deviation from baseline osmolality at different time points. The total allowable error for serum osmolality of 1.5% as specified in the Westgard Biological Variation Database, and the reference change value of 4.1% was used to determine serum and plasma sample stability at the various time points. ¹³ The reference change value (RCV) indicates whether a true clinical change in the value has been observed. The RCV was calculated using a bidirectional Z-score of 1.96, a within-subject biological variation of 1.3% and an analytical imprecision of 0.7%. ¹⁴ The biological variation in urine osmolality is large, hence the large total allowable error of 39.5% quoted in the Westgard Database ¹³ and the RCV of 88%. This is too large to be clinically useful and, in the absence of other guidelines, we considered a deviation from baseline values of >5% to be of clinical significance. A P-value of <0.05 was used to indicate statistical significance.

Results

There were 483 requests for serum osmolality and 628 requests for urine osmolality in 2012 at Tygerberg Hospital. One-hundred and nine (22%) of the serum samples and 132 (21%) of the urine samples were rejected because they were not transported on ice and had not reached the laboratory within 3 h of sampling.

In the samples obtained from our participants, blood osmolality measurements ranged from 252 to 311 mosmol/kg (n = 30). No haemolysed specimens were received. Urine osmolality ranged from 49 to 1157 mosmol/kg (n = 40).

Table 1 shows the changes in serum, plasma and urine osmolality over time. The average room temperature during the period of data collection was 22℃.

Table 1.

Changes in serum, plasma and urine osmolality in stored samples over time.

Time from sampling (h)	Serum		Plasma		Urine
	4–8℃	Room temperature	4–8℃	Room temperature	4–8℃	Room temperature
3	−0.2 ± 3.3	0.4 ± 2.9	−2.0 ± 4.5	−1.1 ± 4.9	−1.6 ± 4.0	−0.1 ± 6.1
6	−0.5 ± 3.0	1.0 ± 4.4	−1.8 ± 4.3	−0.1 ± 4.7	−1.0 ± 4.6	−1.7 ± 4.4
12	−0.7 ± 3.1	1.1 ± 2.5	−2.1 ± 4.3	0.7 ± 4.2	−0.3 ± 4.5	−0.1 ± 5.1
24	0.2 ± 3.7	3.5 ± 3.6  a	0.6 ± 5.7	3.5 ± 5.7  a	−1.5 ± 3.7	−0.7 ± 4.3
36	0.9 ± 4.4	6.4 ± 4.2  a	0.9 ± 10.1	3.3 ± 3.3  a	−1.2 ± 3.2	0.5 ± 5.2

Mean differences from baseline values ± SD are shown in mosmol/kg.

a

Indicates statistically significant changes from baseline values.

Figure 1 demonstrates the serum and plasma osmolality at 4–8℃ and room temperature at each time point. In serum samples kept at room temperature, there was no difference between measurements at baseline and those made at 3, 6 or 12 h. The average change from baseline was 0.4 mosmol/kg at 3 h (P = 0.60), 1.0 mosmol/kg at 6 h (P = 0.20) and 1.1 mosmol/kg at 12 h (P = 0.15). Statistically significant changes were present at 24 and 36 h. At 24 h, the average change from baseline was an increase of 3.5 mosmol/kg (P < 0.05), and by 36 h, the mean increase was 6.4 mosmol/kg (P < 0.05).

Figure 1.

Serum and plasma osmolality at each time point. Mean values and 95% confidence intervals are shown. In serum samples at room temperature, there was an increase of 3.5 mosmol/kg (P < 0.005) at 24 h and 6.4 mosmol/kg (P < 0.005) by 36 h. In cooled samples, there were no significant changes from baseline measurements for up to 36 h. In plasma samples at room temperature, the average change from baseline at 24 h was 3.5 mosmol/kg (P = 0.001), with no further increase at 36 h. In cooled samples, there was a fall in plasma osmolality at 12 h of 2.1 mosmol/kg (P = 0.04) but no significant changes at the other time points.

In plasma samples kept at room temperature, no significant change from baseline measurements was observed up to 12 h. Like serum, the average change from baseline at 24 h was 3.5 mosmol/kg, but at 36 h, there was no further increase.

Serum and plasma samples stored at 4–8℃ showed no significant change from baseline osmolality measurements (see Table 1). For serum, the average change from baseline in the cooled samples was less than 1 mosmol/kg at all time points. For plasma, there was a small decease in osmolality from baseline values of approximately 2 mosmol/kg at 3, 6 and 12 h, and then at 24 and 36 h small increases of less than 1 mosmol/kg from baseline values were recorded.

Figures 2 and 3 illustrate the changes in serum osmolality at 24 and 36 h for samples stored at room temperature. Figure 4 illustrates the changes in plasma osmolality at 36 h at room temperature. Dashed lines indicate the limits of the total allowable error and dotted lines indicate the reference change value.

Figure 2.

Serum osmolality of samples at room temperature: deviation from baseline at 24 h. Dashed lines indicate the limits of the total allowable error and dotted lines indicate the reference change value.

Figure 3.

Serum osmolality of samples at room temperature: deviation from baseline at 36 h. Dashed lines indicate the limits of the total allowable error and dotted lines indicate the reference change value.

Figure 4.

Plasma osmolality of samples at room temperature: deviation from baseline at 36 h. Dashed lines indicate the limits of the total allowable error and dotted lines indicate the reference change value.

Urine osmolality was found to be extremely stable. Analysis of variance with repeated measures for the factor time indicated no significant differences between baseline measurements and measurements made at 3, 6, 12, 24 and 36 h at 4–8℃ or at room temperature (P = 0.21). The average deviations from baseline osmolality measurements did not exceed 2 mosmol/kg at any time point (see Table 1). Figure 5 illustrates percentage changes from baseline urine osmolality at 36 h. Absolute differences are shown in Figure S4 of the supplementary materials. Most measurements do not deviate by more than 2% (or 10 mosmol/kg) from baseline values.

Figure 5.

Urine osmolality of samples at room temperature: percentage deviation from baseline at 36 h. Most measurements do not deviate by more than 2% from baseline values.

Discussion

The investigation of salt and water disorders and the calculation of serum and urine osmolal gaps depend on the accurate measurement of osmolality. Our study has demonstrated good stability of osmolality in blood and urine samples stored at room temperature for up to 36 h. The wide range of blood and urine osmolality included in the study makes the results applicable to a variety of clinical settings, and it therefore appears that the rejection of so many samples based on the current recommendations can be avoided.

There were no changes in the osmolality of serum and plasma samples stored at 4–8℃ for up to 36 h. In the samples stored at room temperature, we found no changes in the osmolality of samples stored for up to 12 h. At 24 and 36 h, there were statistically significant increases in the osmolality of the samples kept at room temperature. At 24 h, the average increase for both serum and plasma osmolality was 1.2%. At 36 h, there was no further change for plasma samples but the osmolality of the serum samples increased to 2.1% above baseline values, exceeding the total allowable error. Importantly, these changes did not exceed the reference change value and were not regarded as clinically significant.

In our setting, most samples for osmolality reach the laboratory within 12 h. Sending samples on ice or storing them at 4–8℃ before measurement is therefore not necessary. In rural clinical settings, the time from sampling to measurement may well exceed 24 or even 36 h. In such cases, it would be acceptable to do serum osmolality measurements up to 36 h, but cooling serum specimens to 4–8℃ or sending a heparinized specimen for measurement of plasma osmolality may be preferable if delays beyond 12 h are anticipated.

Our data revealed excellent stability of urine osmolality over 36 h. There was no difference in osmolality measurements between urine samples stored at 4–8℃ and those kept at room temperature. The average deviation from baseline at 36 h was 0.46 mosmol/kg in samples at room temperature. The total allowable error for urine osmolality is 39.5% and the RCV is 88%. ¹² Most of our samples measured at 36 h were within 2% (or within 10 mosmol/kg) of baseline osmolality values. Sending urine samples on ice or storing them at 4–8℃ before measurement is therefore not required.

The effect on osmolality of storing samples at higher ambient temperatures was not evaluated in this study. We therefore cannot comment on the stability of samples when very warm transportation or storage conditions exist.

In conclusion, we suggest that existing recommendations for sample stability and storage for osmolality be amended. Serum and plasma samples can be stored for up to 36 h before separation and measurement of osmolality. Cooling blood samples to 4–8℃ is advised when delays in measurement beyond 12 h are anticipated. Samples for urine osmolality can be stored for up to 36 h before measurement and cooling of these samples is not required.

Note

Since the acceptance of this article the authors have become aware of the letter by Abbadi et al.¹⁵ reporting good stability of osmolality in heparinised whole blood samples kept at room temperature for up to 24 h.