An improved microfluorometric enzymatic assay for the determination of ammonia

ANALYTICAL

95, 507-511 (1979)

BIOCHEMISTRY

An Improved

Microfluorometric Determination

Enzymatic of Ammonia1

Assay for the

BARRY L. NAZARAND ANTON C. SCHOOLWERTH Departments of Medicine Medical Center,

and Physiology. The Pennsylvania

Renal State

and Electrolyte Division, The Milton S. Hershey University, Hershey, Pennsylvania 17033

Received December 10, 1978 A simple, rapid microfluorometric enzymatic method for the determination of ammonia is described. The basis for the assay is the enzymatic conversion of ammonia and a-ketoghttarate to glutamate by glutamate dehydrogenase and the measurement of disappearance of NADH. The assay sensitivity is l-50 nmol of ammonia.

Numerous approaches for determining ammonia have been reported and used successfully in certain applications. The Conway microdiffusion technique (l), the nesslerization method (2), the ammoniaspecific electrode (3,4), the formalin-titrimetric method (5,6), the Bertholet or phenate hypochlorite reaction (7), and the indophenol reaction (8,9) have been utilized for urinary ammonia and pollution control applications. These methods have limitations to other applications because they require either large sample sizes (0.2- 1.0 ml), lack specificity and sensitivity, or they are slow and cumbersome to process on a large scale. Sensitivity is greatly increased by the mercury-sensitized luminescence technique (lo), but this approach requires rather sophisticated and expensive apparatus and is not suitable for large-scale processing. Enzymatic methods utilizing glutamate dehydrogenase seem to overcome most of the limitations described above. Ammonia and cy-ketoglutarate react in the presence of glutamate dehydrogenase to form glutamate and oxidize a stoichiometric quantity of NADH to NAD. Several investigators 1 This research was supported by Grant AM-19714 from the National Institutes of Health. 507

have employed this reaction for determining ammonia by monitoring spectrophotometrically (A 340“,,,) the oxidation of NADH (I I- 13). The sensitivity of this approach is further enhanced by using a fluorometric measurement of NADH oxidation. Rubin and Knott (14) have reported a fluorometric approach for blood ammonia analysis. The fluorometric method presented here describes a modification of the method of Rubin and Knott (14) which makes even greater use of the sensitivity availed by fluorescence, and is well suited to measuring 50-60 samples in duplicate simultaneously. METHODS

a-Ketoglutarate, ethylenediamine tetracetic acid (EDTA), reduced nicotinamide adenine dinucleotide (NADH), and bovine liver L-glutamate dehydrogenase (Type II, approximately 500 units/ml and substantially free of ammonium ions) were purchased from Sigma Chemical Company. All other reagents were purchased from Fisher Scientific Company and were of analytical grade. Deionized water (high resistance, greater than 5 MQ and with low fluorescence background) were used for all solutions. Glass0003-2697/79/080507-05$02.00/0 Copyright All rights

0 1979 by Academic Press. Inc. of reproduction I” any form reserved.

508

NAZAR AND SCHOOLWERTH

ware was treated as described by Lowry and Passoneau (15) and cuvettes (10 x 75mm Pyrex test tubes) were made of low fluorescence glass. Standard stock solutions containing 100 mM a-ketoglutarate, 2 mg/ml NADH, pH 7-8, and ammonia standard (O.Ol- 1.0 mM NH,Cl) were stored at -30°C. Potassium phosphate buffer, 0.2 M, pH 7.6, 0.1 M EDTA and L-glutamate dehydrogenase were stored at 4°C. Fluorometric measurements were made with a Farrand A4 fluorometer using Corning glass filter No. 5860 as primary filter, which isolates the mercury line at 365 nm, and secondary filters Nos. 4303 and 3387, which have maximal transmission at 470 nm. A quinine sulfate standard (0.1 pg/ml) was used each day to standardize the fluorometer for the assay. The incubation buffer was prepared in the following proportions.

strutted by plotting the absolute values of change in fluorescence for the standard concentrations of ammonium chloride. When a large number of samples was being measured, it was more efficient to compute the slope of the standard curve and multiply the coefficient of the slope times the change in relative fluorescence of each sample minus the change of the blank: (coefficient

of slope)

x (ARF sample - ARF blank) = concentration

of ammonia.

Trouble shooting. The order of mixing reagents described above was designed to eliminate extraneous sources of ammonia. Although commercial sources of glutamate dehydrogenase claim to be “substantially free” of ammonia, there may likely be traces of ammonium ions which could affect measurements at low concentrations. Aqueous 100 ml 0.2 M phosphate buffer, pH solutions can also dissolve ammonia from 7.6 the atmosphere and this can influence the 6 ml 0.1 M cu-ketoglutarate (final con- outcome of the assay. By mixing all reacentration approximately 6 gents in the buffer prior to the addition of mM) the standard or samples, these sources of 2 mg/ml NADH (final concen0.6-4 ml ammonia will be consumed before the ontration 0.015-o. 1 mM) set of the assay. 1.2 ml glutamate dehydrogenase (fiA possible source of error with this nal concentration 6 units/ml) sequence is high fluorescence in the sample 0.1 M EDTA (final concentra0.6 ml itself. If samples contain high fluorescence, tion 0.5 IriM) there are several approaches to eliminate this factor. The first is to include a sample This mixture was well stirred and brought to room temperature or near 25°C. One blank (i.e., sample plus buffer without glutamilliliter was placed into each cuvette and mate dehydrogenase) and subtract fluoresthe initial fluorescence for each tube was cence due to chromogens or quenching agents in the sample. If a limited supply of measured and recorded. Then 50 ~1 of blank sample material precludes this, then one (H,O), ammonia standard (0.01-1.0 mM), may eliminate glutamate dehydrogenase or unknown sample was added to the cufrom the buffer mixture and measure the vettes, respectively. After a 30-min incubainitial fluorescence with the sample and buftion period the final fluorescence was measfer minus glutamate dehydrogenase in each ured. The change in relative fluorescence tube. After the initial fluorescence measure(ARF)2 was computed by subtracting the ment, the reaction in each tube is begun final fluorescence from the initial fluoresby adding 10 ~1 of the enzyme suspension cence. A standard curve was then conwith care that an equal quantity of gluta2 Abbreviations used: RF, Relative fluorescence. mate dehydrogenase is added to each tube.

MICROFLUOROMETRIC

AMMONIA

509

ASSAY 50

The water blank, then, will correct for fluorescence changes due to the enzyme mixture.

SAMPI

E VOLUME

02

05

50~1

40 30

RESULTS

Preliminary studies were performed to determine the optimal conditions for the assay system. These studies indicated that a glutamate dehydrogenase concentration of 6 units/ml resulted in the greatest stability and reproducibility of the assay. The effect of pH and time of incubation on the attainment of equilibrium in the assay system was also evaluated. At pH 6.8 and 7.2 the time of equilibration required 60 min or more and tended to result in high blanks. At pH 7.6 and 8.0, equilibrium was attained within 30 min. Although equilibrium was quite rapid at pH 8.0, this pH level occasionally resulted in erratic results due to. loss of ammonia from solution in gaseous form. Since equilibration at pH 7.6 was attained within 30 min with very consistent results, this pH was adopted for the standard assay procedure. A pH of 7.6 was sufficiently low to prevent loss of ammonia in the gaseous state, and yet near enough to the pH optimum of glutamate dehydrogenase to allow rapid equilibration. The measurement range and sensitivity of the assay can be varied from I-50 nmol ammonia by adjusting the volume of standards and samples used in the procedure and by altering the initial level of NADH in the reaction medium. To process a large number of samples of widely divergent ammonia concentrations, a high NADH level (0.060.1 mM final concentration) was generally utilized (Fig. 1). With these levels of NADH, the assay was quite reproducible with approximately the same fluorometric readings and change in relative fluorescence obtained each day for a given concentration of ammonia and the same instrument setting. At this level of NADH in the incubation buffer a sensitivity of 2-5 nmol ammonia could be achieved consistently. Increased

20 10 p 005

ARF

07

10

50SAMPLE

VOLUME

100 ~1

40

30 20 10 I_I 002501

05

02 AMMONIA

(mM)

FIG. 1. Standard curve-reproducibility at different sample volumes. Each point represents the mean zSD of change in relative fluorescence (ARF) plotted against ammonia concentration in 12 standard curves utilizing 0.1 mM (final concentration) NADH in the incubation buffer. Sample volumes of 50 ~1 are shown in the upper panel and of 100 ~1 in the lower panel. Concentrations are those of the original samples added.

sensitivity can be obtained by decreasing the initial NADH level to approximately 0.015 mM. As shown in Fig. 2, ammonia in the range of 0.5-2 nmol can be measured by this change in NADH level, if ammonia contamination can be kept low. It is important that this assay be performed in carefully timed sequence. Although the destruction of NADH is slight at pH 7.6, a small but significant decrease in fluorescence was detected under the conditions of the assay. This change was readily corrected by careful attention to the time sequence (30 min incubation for each sample) and the use of appropriate blank controls. This assay was conducted both with and without EDTA in the incubation buffer. The absence of EDTA had little effect on the

510

NAZAR AND SCHOOLWERTH 10

known samples. The samples were perchloric acid extracts of rat kidney mitochondria incubated with 1 mM glutamine as described by Schoolwerth et al. (16). Recovery of ammonia in these samples approximated 100% at the levels tested.

r

a6-

DISCUSSION 0

.05

.I AMMONIA

.15

.2

.25

(mM)

FIG. 2. Assay sensitivity. Each point represents the average of two determinations of change in relative fluorescence minus blank (ARF) plotted against ammonia concentration. The concentrations are those of the original samples added. In the determination of these standard curves, a lower concentration (0.015 mM) of NADH was utilized to increase the sensitivity of the assay. Sample size was 50 ~1.

outcome of the standard curve. Linear regression analysis showed r = 0.97 without EDTA and r = 0.99 with EDTA. For the measurement of unknown samples, however, EDTA was found to increase the stability of the assay. The mechanism by which EDTA stabilizes the assay of samples is not known but may relate to the fact that divalent cations can enhance the native fluorescence of NADH (15). Although this enhancement occurs at alkaline pH levels (15) and has not been shown at neutral pH, as used in this assay, Mg2+ or Cu*+ could contribute uncertain variability to the assay. To test this possibility, micromolar concentrations of Mg2+ and CU*+ were added to the incubation buffer. No effect was observed with Mg*+. Addition of lo-100 pM Ct.P+, however, did affect NADH fluorescence in this reagent system. This effect is shown in Fig. 3 as an inhibition by Cu*+ of the change (decrease) in relative fluorescence (ARF) resulting from increasing concentrations of ammonia. The effect was abolished with the addition of EDTA to the incubation buffer. Table 1 shows the percentage recovery of known amounts of ammonia added to un-

The use of enzymatic procedures for measuring ammonia provide specificity, sensitivity, and efficiency for multiple sample processing. The use of fluorometric procedures not only enhances the sensitivity over spectrophotometric procedures, it allows one to conduct the entire assay in cylindrical test tubes, eliminating the need for transfers to expensive square cuvettes 5Or 40

a 20 I.& 10 30

,-“: NO EDTA

rl

0 0.05

5or 40

.25

.5

.75

c

1.0

II

0.5 mM EDTA

/

P

2 30 a 20 10 v Of

0.05

I

I

I

A

.25

.5

.75

1.0

AMMONIA

(mM)

FIG. 3. Effect of EDTA on the fluorescence enhancement caused by CtP+. Ammonia standard curves were plotted as in the previous figures. The change or decrease in relative fluorescence (ARF) was plotted in the absence (0) and in the presence of 1 (B), 10 (O), and 100 (A) PM cupric sulfate. Standard curves obtained without EDTA are shown in the upper panel and with 0.5 mM EDTA in the lower panel.

MICROFLUOROMETRIC

AMMONIA

TABLE RECOVERY

OF AMMONIA

Initial sample ammonia” (nmol)

5 nmol Ammonia added to samples (nmol)

4.69 16.66 4.68 17.71 4.17 18.23 2.09 15.62 Mean t SD

9.56 21.79 9.64 22.52 9.40 23.59 6.81 20.26

Recovery (%) 97.3 102.5 99.2 96. I 104.6 107.2 94.3 92.8 99.3 2 5.1

for absorbance measurements. It is possible to perform 50-60 determinations in duplicate simultaneously within 3 h. The present assay represents a modification of the fluorometric procedure described by Rubin and Knott (14). The changes consist of (i) reducing the volume from 3 ml to 1 ml, (ii) adding the sample last in order to allow any contaminating ammonia to react ahead of time, and (iii) addition of EDTA to the incubation buffer to reduce variability and enhance stability of the assay. Rubin and Knott (14) previously reported a method with a sensitivity of 5-6 nmol ammonia. The range of the present assay can be adjusted from 1- 50 nmol. The assay was found to be quite reproducible with little variation from day to day. The use of EDTA in the assay system was found to stabilize the conduct of fluorescence changes in tissue sample extracts which contain varying amounts of divalent cations. REFERENCES I. Conway, E. J., and O’MalIey, them. J. 36, 655-661.

1 ADDED

o Perchloric acid extracts of rat kidney mitochondria Extracts were neutralized with 0.5 M 4-morpholinopropane stored at -20°C until the assay.

E.

(1942)

Bio-

511

ASSAY

TO SAMPLES

25 nmol Added to samples (nmol) 30.12 42.69 28.03 42.04 28.69 43.85 28.77 39.32

Recovery (%I 101.7 104.1 93.4 97.3 98.1 102.5 106.7 94.8 99.8 + 4.7

incubated either zero or 4 min in 1 mht glutamine. sulfonic acid (MOPS), 3 M KOH to pH 6.7-7.0 and

2. Seligson, D., and Hirahara, K. (1957) J. Lab. Clin. Med. 49, %2-974. 3. Dewolfs, R., Broddin, G., Clysters, H., and Deelstra, H. (1975) Z. Anal. Chem. 275, 337341. 4. Hoge, J. H. C., Hazenberg, H. J. A., and Gips, C. H. (1974) Clin. Chim. Acta 55, 273-279. 5. Jorgensen, K. ( 1957) Stand. J. C/in. Lab. Invest. 9, 287-291. 6. Chan, J. C. M. (1972) C/in. Biochem. 5, 94-98. 7. Kaplan, A. (1%9) Methods Eiochem. Anal. 17, 31 l-324. 8. Chaney, A. L., and Marbach, E. P. (1962) Clin. Chem. 8, 130-132. 9. Gips, C. H., Reitsema, A., and Wibbens-Alberts, M. (1970) Clin. Chim. Acta 29, 501-505. 10. Ho, W., and Harker, A. B. (1976) Anal. Chem. 48, 1780- 1784. 11. Kirsten, E., Gerez, C., and Kirsten, R. (1963) Biochem. Z. 337, 312-319. 12. DeHaan, E. J., Tager, J. M., and Slater, E. C. (1967) Biochim. Biophys. Acta. 131, l-13. 13. Buttery, P. J., and Rowsell, E. V. (1971) Anal. B&hem. 39, 297-310. 14. Rubin, M., and Knott, L. (1967) C/in. Chim. Acta 18, 409-415. 15. Lowry, 0. H., and Passoneau, J. V. (1972) A Flexible System of Enzymatic Analysis, Academic Press, New York. 16. Schoolwerth, A. C., Nazar, B. L., and LaNoue, K. F. (1978) J. &al. Chem. 253, 6177-6183.