-------
24
~ 16
1
o
8
0 0.2 0.4 0.6 0.8 1.0
ALKALINE PHOSPHATASE (/jg)
Figure 26. Kinetic assay of alkaline phosphatase. p-Nitrophenol
production was measured spectrophotometrically at
410 nm using a chart recorder calibrated for a full-
scale deflection for one absorbance unit. The recorder
was operated at 5 cm/min and the minimum change detect-
ed was aAA of 0.005/min. The sample holder was kept
at 30 C for the assays. The molar absorptivity_co-
efficient of_p_-nitrophenol is 1.62 x 10 M
cm
-1
77
-------
60
o
o 40
10
X
0.
o
a:
20
10 20 30 40 50
ALKALINE PHOSPHATASE (ng)
Figure 27. One-hour incubation assay of alkaline phosphatase.
p-Nitrophenol production was measured spectrophoto-
metrically at 410 nm after a 1-hr incubation of 0.1 ml
samples with 0.9 ml of reagent (0.0066 M disodium
p-nitrophenyl phosphate in 0.6 M Tris-HCl, pH 8.2)
at 37 C. The values have been corrected for blank
hydrolysis of substrate.
78
-------
3.0
2.0
1.0
24 48 72
TIME (hours)
96
CD
O
-1.0
B
-1.0 0 1.0
LOG ALKALINE PHOSPHATASE (ng)
Figure 28. Long-term incubation assay of alkaline phosphatase.
Samples (0.5 ml) were mixed with 0.5 ml reagent
(0.0135 M disodium p_-nitrophenyl phosphate in 1.0
M Tris-HCl/opH 8.0) in capped plastic vials and incu-
bated at 37 C. Duplicate samples were removed and
£-nitrophenol production measured spectrophotometric-
ally at 410 nm at 24-hr intervals. Reagent blanks were
also read each d so that nonenzymatic £-nitrophenol
formation could be determined and used to correct the
results of the enzyme samples.
A. Time course of product formation by alkaline phos-
phatase - 8.8 ng (0), 0.88 ng (t), and 0.09 ng (C),
measured at 24-hr intervals for 4 d.
B. Plot of enzyme amounts after a 24- hr incubation
(•) and 72-hr incubation (0).
79
-------
range of alkaline phosphatase tested. Therefore, a long
incubation of one or more days can be used to increase the
sensitivity of the alkaline phosphatase determination at least
10-fold over a 1-hr incubation assay.
c. Measurement of Alkaline Phosphatase in J2. coli
Cells.
Alkaline phosphatase was measured in E. coli cells (8x10
-8x10 /ml) treated with 0.1% toluene ari3 assayed in an 1-hr
incubation assay. Cell samples .(0.1 ml) were mixed with 0.9 ml
reagent and incubated 1-hr at 37 C. Cell blanks (cell + buffer)
were used so that absorbance caused by the cells could be
subtracted from the final absor±>ance readings of the samples. The
limit of detection was 8.5x10 cells/ml, containing one ng of
alkaline phosphatase (from Figure 27).
3. Catalase
Hydrogen peroxide produced by cellular oxidases is scavenged
by catalase (E. C. No. 1.1.11.6). Catalase activity is measured
by quantitation of residual hydrogen peroxide, either kinetically
in a spectrophotometer or after an incubation period.
a. Kinetic Assay
Catalase was measured by spectrophotometric determination
(240 run) of the rate of hydrogen peroxide destruction (Figure
29). A 3-ml assay volume was used because oxygen bubbles
produced by the catalase caused fewer problems in 3-ml reaction
volumes than in 1 ml. The limit of detection was 0.2 pg and the
range tested was 0.2 - 10 jjg catalase.
b. Incubation Assay
An 1-hr incubation assay for catalase was used to increase
the sensitivity of catalase determination (Figure 30). The
amount of hydrogen peroxide utilized was determined by the
difference in absorbance between hydrogen peroxide-containing
blanks and samples containing catalase. The limit of detection
was 1 ng and the range 1-100 ng catalase.
c. Measurement of Catalase in E._ coli.
4 9
Catalase was measured in £. coli cells (10 -10 /ml)
suspended in distilled water. Toluene treatment was not used
since toluene absorbs strongly at 240 nm. An assay was used in
which 0.1 ml sample was added to 0.9 ml reagent in capped vials
and incubated 1 hr. The limit of detection was 10 cells/ml
which had the equivalent of 1 ng catalase/10 cells.
80
-------
12.0
E 8.0
\
o
E
4.0
02468
CATALASE (jug)
10
Figure 29. Kinetic assay of catalase. Catalase samples (2 ml)
were mixed with 1.0 ml buffered hydrogen peroxide in a
3-ml cuvette and the decrease in absorbance was measur-
ed spectrophotometrically at 240 nm using a chart re-
corder calibrated at 1 absorbance unit/full scale de-
flection and running at 5 cm/min. The limit of de-
tection was a A A of 0.005/min. Sample temperature
was held at 30 C for the assays.
81
-------
2.0
1.5
o
E
CM 1-0
O
CM
0.5
10 20 30 40
CATALASE (ng)
50
Figure 30. Incubation assay of catalase. Catalase samples
(0.1 ml) were mixed with 0.9 ml buffered hydrogen per-
oxide and incubated 1 hr at 37 C. The hydrogen per-
oxide used was measured as the difference in absorb-
ance between samples and blanks.
82
-------
4. Diaphorase
Figure 31 shows the standard curve obtained with dia-
phorase (E.G. No. 1.6.4.3) assayed as described in the Experi-
mental Procedure section (Section 5).
5. Lactate Dehydrogenase
The glycolytic enzyme lactate dehydrogenase (E. C. No.
1.1.1.27) catalyzes the reduction of pyruvate to lactate and
oxidation of NADH to NAD . Lactate dehydrogenase from some
microbes can use other electron donors such as cytochrome c,
ferricyanide, and quinone in addition to NADH, but only the most
common substrates, NAD and NADH, were used in this study. In
incubation assays, lactate dehydrogenase was coupled to dye
reduction by diaphorase, a non-specific enzyme that uses NADH as
substrate.
a. Kinetic Assay
Initial velocity measurements of lactate
dehydrogenase were made spectrophotometrically by using a chart
recorder to monitor changes in absorbance (Figure 32). The limit
of detection was 0.1 ug of lactate dehydrogenase or 1.5 nmol NADH
oxidized/ min and the range 0.1-15 pg of lactate dehydrogenase.
b. Incubation Assay
A 1-hr incubation assay based on lactate and NAD as
substrates was used to measure lactate dehydrogenase. Activity
was determined by a coupled enzyme reaction in which NADH was
oxidized with concomitant reduction of the dye INT by diaphorase.
The use of a dye such as INT or TTC is preferred for many
determinations because the product to be measured can be
extracted from turbid or soil samples with organic solvents, thus
improving the sensitivity of the measurement. INTF formation was
measured, both before and after extraction with
tetrachloroethylene:acetone (Figure 33). The limit of detection
was 1.5 ng and 1.0 ng of lactate dehydrogenase for the unextracted
and extracted samples, respectively. The increased response of
the extracts, even though the INTF concentration was one-half
that of the unextracted samples, was the result of complete
solution of INTF in the extract. Precipitation of the water-
insoluble INTF was observed in the samples prior to extraction,
and this reduced the response for each sample.
c. Measurement of Lactate Dehydrogenase in £. coli.
Lactate dehydrogenase was measured in toluene-treated £._
co^i using the incubation assay. Cells (10 -10 /ml or 10 -
10 /assay) were treated with 0.1% toluene prior to addition of
83
-------
1.0
0.8
0.6
c
1
0.4
0.2
10 20 30 40 50 60
jug DIAPHORASE
Figure 31. Assay of diaphorase. The reduction INT was measured
at 490 nm as described in Section 5.
84
-------
0.40
0.30
o 0.20
E
0.10
24 6 8 10 12 14
LACTATE DEHYDROGENASE (jug)
Figure 32. Kinetic assay of lactate dehydrogenase. Absorbance
measurements were made at 340 nm. The chart recorder
was calibrated so that a full-scale deflection was 1
absorbance unit. The chart speed was 5 cm/min and the
minimum change detected was aAA of 0.005 A/min. The
sample holder on the spectrophotometer was maintained
at 30 c for the assays.
85
-------
0.9 ml of the reagent used for the incubation assay. Samples were
incubated 1-hr and the INTF produced was extracted with
tetrachloroethylene : acetone and read spectrophotometrically
at 490 nm. Lactate dehydrogenase activity was observed in all
samples, the limit of detection being 10 cells/ml. This
corresponds to the INTF produced by a 1-hr incubation of 1.0 ng
lactate dehydrogenase using the standard curve for extracted
samples in Figure 33.
d. Enzymatic Cycling Assay
As stated earlier, the limit of detection for an enzymatic
activity is the limit of detection for a change in the amount of
substrate or product being measured. An increase in sensitivity
of product measurement also increases the sensitivity of enzyme
measurement. This concept was tested by using enzymatic cycling
to measure NAD produced by lactate dehydrogenase and so improve
the sensitivity of detection of the enzyme.
i. Destruction of Excess NADH — All unreacted
NADH must be destroyed prior to enzymatic cyling so that the
response will be proportional to the enzymatic activity. This
was accomplished by treatment of samples w^th hydrochloric acid,
which destroys NADH without affecting NAD , the product of the
lactate dehydrogenase reaction. The half-life of NADH at pH 2 at
37 C is 0.07 min so the time required for 99.99% destruction of
NADH under assay conditions, using the equation in Table 8, is
0.93 min. The reagent used contained 1 nmol NADH, and a 10-min
incubation at pH 2 was used, which was sufficient to decrease the
amount of NADH present to 1 x 10" mol without changing the
amount of NAD produced by the reaction.
ii. Measurement of Lactate Dehydrogenase
Lactate dehydrogenase samples (Ipg - 1 ng) were incubated
with substrates pyruvate and NADH for 1 hr at 37 C. Unreacted
NADH was destroyed by acidification of the samples with
hydrochloric acid and by incubation for 15 min. The samples were
then neutralized with NaOH and the NAD was amplified by the
enzymatic cycling procedure. The results are shown in Figure 34.
The limit of detection was 1 pg and the range of the as|ay was 1
pg-1 ng lactate dehydrogenase. Enzymatic cycling of NAD improv-
ed the sensitivity of lactate dehydrogenase detection 1000-fold
over the 1-hr incubation assay.
iii. Measurement of Lactate Dehydrogenase in E_.
coli Using Enzymatic Cycling.
Samples containing J2. coli cells (10 -10 /sample)
were treated with 0.1% toluene and then lactate dehydrogenase mea-
sured by a 1-hr incubation with the pyruvate-NADH reagent followed
86
-------
0.10
0.08
O 0-06
o>
0.04
0.02
4 6
LDH (ng)
8
10
Figure 33. Measurement of lactate dehydrogenase using an incuba-
tion assay . INTF formation was measured spectrophoto-
metrically at 490 nm before (0) and after (•) extrac-
tion with 2 ml tetrachloroethylene:acetone (1.5:1).
Samples (0.1 ml) were mixed with 0.9 ml reagent and
incubated 1 hr at 37 c.
87
-------
TABLE 8. NADP+ DESTRUCTION WITH ALKALI (pH 12.5)
Temperature
( C)
38
65
Half-Life
(min)
21.0
5.5
N = N e
Destruction
for 99.9%
209
54.8
- 0.693 t
fcl/2
Time (min)
for 99.99%
279.1
73.0
where
N = the amount present
N = the starting amount
t°= the incubation time
t,/2 = half-life under the incubation
conditions
88
-------
10'
h;
CO
LU
k-
- io3
1.0 10 10* 10'
LACTATE DEHYDR06ENASE (pg)
Figure 34. Measurement of lactate dehydrogenase using enzymatic
cycling of NAD . Picogram amounts of lactate dehydro-
genase were measured by enzymatic cycling of NAD pro-
duced by a 1-hr incubation of 20 jal samples with 10 pi
of LDH reagent (10 nmol NADH and 30 nmol gyruvate in
0.09_M phosphate buffer, pH 7.4) at 37 C. Unreacted
NADH was^ destroyed by treatment with 10 jul 1 N HC1 and
the NAD was measured by the enzymatic cycling proce-
dure already discussed.
89
-------
by NADH destruction and enzymatic cycling of NAD . Using the
lactate dehydrogenase standard curve in Figure 34, the cell
samples read as follows:
10_ cells - 5.4 pg;
10i? cells - 56 pg;
10 cells - 390 pg.
The sample containing 10 cells did not have a value above
the blanks which were run with the samples. The concentration of
the 10 cells/assay was 5 x 10 cells/ml. This represented
only a two-fold increase in sensitivity of lactate dehydrogenase
measurement in £. coli over a 1-hr incubation assay using the
dye INT and is not worth the effort required.
6. Microtox
a. Basic Measurements
Figure 35 shows the standard curve for determining the
toxicity of toluene with the Microtox assay. The experiment was
done as described in Section 5. The EC50 is 50 ppm. Table 9
shows the EC50 values for a number of compounds and the broad
range of concentration of inhibitors that can be measured in the
system—ranging from 47 x 10 ppm for ethanol to 2.5 ppm for
cyanide. Although the light output from a sample of bacteria was
reduced to 1/3 during 5 hr of storage after hydration, the EC50
values determined for m-cresol with fresh and 5-hr-old samples
were 11 and 12 ppm, respectively. An example of the
reproducibility of experiments is that with a concentration of 9
mg/L of m-cresol the normalized light loss was 44.0% for the
first experiment and 44.9% for the second.
b. Environmental Samples
The results of typical application of the Microtox method to
environmental samples are shown in Table 10. In Part A various
local water sources were tested, and in Part B various spring
water samples from eastern Oklahoma were tested. The amounts of
lipopolysaccharides present in these samples as determined by the
Limulus amebocyte lysate assay are also shown. Part C shows the
results of testing various oil refinery effluents. The LC50 for
the fathead minnow 96-hr bioassay is also shown for some.
c. Toxicity of Pesticides
A number of commonly used pesticides were tested in the
Microtox bioassay (Table 11). The luminescent bacteria were
fairly sensitive to most of the compounds tested. Thus, the
Microtox assay is an effective method for their determination.
The sensitivity with which light production can be measured
with commercially available instrumentation and the availability
90
-------
LU
UJ
O
100
75
50
25
10
7.5
5
2.5
20 40 60
% LIGHT LOSS
Figure 35. Standard curve for toxicity of toluene with the
Microtox assay. The inhibition of light production
by various concentrations of toluene was determined as
described in Section 5. The logarithm of toluene con-
centration is plotted against the per cent of light
production loss by the luminescent bacteria.
91
-------
TABLE 9. TYPICAL EC , LD and LC FOR SELECTED COMPOUNDS
50 50 50
Microtox Rat (oral) Fish toxicity
Typical Compounds EC,.,, ppm LD g/kg LC,-^ mg/L
50 50 50
Ethanol 47,000 14 13,000-
1-Butanol 44,000 4.4 1,9002
Benzene 200 5.7 502
Toluene 50 . 5.0 23 -
Phenol 26 0.53 5.0 -
m-Cresol 11 2.0 19 (p_ cresol)
Formaldehyde 8.7 0.80 250
Respiratory Inhibitors
Amytal 1,000
Thenoyltrifluoroacetone 3.5
Cyanide 2.5
Azide 400
Arsenate 94
Lethal dose • From the Merck Index, Merck & Co., Rahway,
N.J. 50
2
Lethal concentration .... BRUNGS, W. A., J. H. McCORMICK, T. W.
NEIHEISEL, R. L. SPEHAS, C. E. STEPHAN, and G. N. STOKES:
J. Water Poll. Control 49, 1425 (1977).
McKIM, J. M., R. L. ANDERSON, D. A. BENOIT, R. L. SPEHAR, and
G. N. STOKES: J. Water Poll. Control 48, 1544 (1976).
92
-------
TABLE 10. APPLICATION OF MICROTOX TO ENVIRONMENTAL SAMPLES
Water Sample
A. Local Water
Barstead still system
Burned field water
Essenberg well
Glass-distilled water
Spring (obtained locally)
Tap water
Water fountain
Microtox Assay
LL.
LL,
15
100
0
0
93
35
2
27
100
2
0
96
35
7
Lipopolysaccharide
pg/mL
4.4 x
5.0 x
10
5.0 x
1.0 x
75
0 i
1.0 x 10;
10;
10"
B. Eastern Oklahoma
Park Spring
Sparrow Hawk Spring
Speed's Spring House
Stream
Wilson Ranch Spring
0
-15
-24
0
-14
0
-15
-20
-1
-14
5.0 x 10_
2.5 x 10'
4
2.5 x 10*
Percent required for
Microtox Fathead Minnow
LL,
EC
50
LC
50
C. Oil Refinery Effluents
ETE - 55
17 - 51 - 80
LNX
UQB - 3
UQB - 4
The sample to be tested constituted 45% of the Microtox assay
volume. For the ECc,, determination in part C the concentration
of the effluent was varied.
46
34
0
90
0
58
74
100
1.8
100
65
65
65
42
75
93
-------
TABLE 11. MICROTOX TOXICITY OF SELECTED PESTICIDES
Common Name of Compound ECq0 PPm LDR0 m<3/k9
Microtiox Rat (oral)
Captafol 7 6200
Carbaryl 2 500
Cyhexatin 10 540
Diazinon 1.7 300
Dichloran 3 5000
DDT 7 110
Glyphosate 7.7 4300
Malathion 10 1400
Paraquat 780 150
Ridomil 120 670
Thiabendazole 3400 3100
From Farm Chemical Handbook, 1980, Meister Publishing Co.,
Willoughby, OH.
94
-------
of suitable freeze-dried preparations of luminescent bacteria
enhance the usefulness of the Microtox bioassay. The ease of
performance of the basic measurement and the range of
concentrations of toxic and other compounds that can be
determined are also attractive features of the Microtox bioassay.
Table 9, Part A shows a variety of compounds with differing
toxicities assayed in three systems. These three measures of
toxicity have good correlations. The Microtox has a correlation
coefficient of 0.90 with the rat toxicity and 1.0 with the fish
toxicity. Advantages of the Microtox test are the short time (30
min) required for an assay and the statistical advantage in using
over 10 bacteria instead of a small number of rats or fish in
the other bioassays. Because the luminescent pathway is a branch
of the electron transport chain, the effects of various
respiratory inhibitors were determined. The relationship between
the electron transport chain and the luminescent pathway would
suggest that inhibitors acting after the divergence of the two
pathways would be less effective in blocking light production and
even increasing it. This is not observed; therefore, the
metabolic interrelationships must be more complicated.
The local water samples varied greatly in both their
inhibition of light production and in their lipopolysaccharide
content. There is no correlation between these two parameters,
but there is no reason to expect that there should be. Some of
the eastern Oklahoma spring waters stimulate light production
(shown by the minus sign). Certain of the compounds (at low
concentrations) such as ethanol also stimulate light production.
These compounds could either be substrates or change the
availability of components.
Table 10 shows little relation between light inhibition by
selected pesticides and their LD,-« orally for the rat. The
correlation coefficient between these variables is 0.09.
G. Lipopolysaccharide Determination
Limulus Amebocyte Lysate Assay
The unusual reaction of the hemocyte lysate from the
horseshoe crab in which a proclotting enzyme is activated by
lipopolysaccharides (LPS) producing a clot has been exploited in
several biomedical applications (39). Since the
lipopolysaccharides of gram-negative bacteria are pyrogenic, and
are endotoxins, their detection has great clinical significance.
Sullivan, Valois, and Watson (94) have reviewed the development
of the Limulus amebocyte lysate system (LAL) as a test for
endotoxins.
A method for the detection of as little as 100 pg of
endotoxin has been described by Jorgensen and Smith (95). A slide
test has been developed which reduces the volume required
(Frauch, (96) and Flowers (97)).
Watson, Novitsky, Quinby and Valois (98) have used the LAL
95
-------
determination of LPS as an indirect assay of the number of
bacteria in the marine environment. The biochemical basis of the
clotting has been established through the studies of Liu and
coworkers (99-101). A MW of 150 K J^as found for the proclotting
enzyme which, when activated by Ca and endotoxins, is a serine
protease (possible MW 79-84 K). The purified coagulogen has a MW
of 24.5 K; when acted upon by the clotting enzyme, it is
converted to a 17.5-K protein and a 5.5-K peptide by hydrolysis
at a single arginine-lysine linkage. There is a LPS binding
protein of MW 80 K which is found on the amebocyte membranes.
Correlation ofthe LAL test with numbers of coliform,
enteric, gram-negative, and heterotrophic bacteria was found by
Evans, Schillinger, and Stuart (102) for stream water. Goto and
Nakamura (103) used a dry-up method on a glass slide to reduce
the volume of JLAL needed. Picogram quantities of LPS were
measured with ( I)-labeled coagulogen by Munford (104); this
procedure also required less LAL.
An alternative assay using a chromogenic peptide substrate
for the active clotting enzyme was useful over the
LPS range of 0.2 to 10 ng/ml (105).
Figure 36 shows the standard curve that we obtained when
LPS-induced coagulation was measured photometrically. Using E._
coli cells as the standard with their number being determined by
plate counts and turbidity measurements, we obtained the results
shown in Figure 37.
H. ATP Determinations
Adenosine triphosphate may be measured directly with firefly
luciferase or indirectly with a coupled enzyme system using NADPH
for spectrophotometrie or fluorometric determination. Enzymatic
cycling can be used to amplify the response and increase the
sensitivity of measurement of ATP.
1. Coupled Enzyme Assay
A coupled enzyme reaction of hexokinase-glucose 6- phosphate
dehydrogenase was used to measure ATP by the fluorescence of
NADPH. Tests of the assay using both ATP and glucose 6-phosphate
standards showed that the reaction was complete and that all ATP
was used up in less than 5 min at 37 C. A typical standard.curve
is shown in Figure SS.iThe limit,,of detection was 2 x 10 mol
and the range 2 x 10 -2 x 10 mol of ATP.
2. Enzymatic Cycling
The sensitivity of detection of ATP was improved by
enzymatic cycling+of the NADPH produced by the coupled enzyme
assay. Excess NADP in the coupled enzyme reagent was destroyed
by alkali treatment prior to cycling. This approach was selecte^
instead of an ADP/ATP cycle (Figure 39) because the NADP
96
-------
200 400 600 800
fg LPS
20 40 60 80
E. coli cells
Figure 36. LPS standard curve. The Limulus amebocyte
lysate assay for lipopolysaccharide was performed as
described in Section 5. The reaction was followed by
the spectrophotometric assay.
Figure 37. E_. coli determination using LAL. The results
of the more sensitive LPS procedure with a bromosulfa-
lein determination of the gelled protein are shown.
The E^. coli cells were diluted by another worker so
that the determinations were not prejudiced. The pro-
cedure is described in Section 5.
97
-------
60
40
UJ
h-
20
05 10 15
ATP (nmol)
Figure 38. Coupled enzyme assay of ATP. ATP was measured by
reaction with hexokinase and glucose-6-phosphate to
produce NADPH, which was measured fluorimetrically
(excitation 340 nm, 1-mm slit; emission 455 run, 2-mm
slit). ATP samples (0.1 ml) were mixed with the
coupled enzymes reagent (0.9 ml) and incubated 15 min
at 37°C.
98
-------
NADP*
6-P-gluconate
glucose
ATP
pyruvate
glucose-6-P
glucose-6-P
dehydrogenose
•> NADPH
ADP
PEP
Figure 39. Reaction scheme for the enzymatic cycling of ATP.
99
-------
cycling procedure had been developed and could be applied to the
measurement of another metabolite, ATP.
a. Destruction of Excess NADP
For enzymatic cycling to be useful, all unreacted NADP
must be destroyed so that only the NADPH formed by the coupled
enzyme assay of ATP participates in the cycling reactions (Figure
40). This was accomplished by alkali treatment prior to addition
of the cycling reagent, since NADP +is labile at high pH (Table
(8)). A 99.99% destruction of NADP wgs accomplished by a 1-hr
incubation of samples at pH 12.5 (65 C) which means that 5 x
10 mol of NADP remained of the 1 £10 y mol in the
reagent after treatment. The residual NADP was a major contri-
butor to the blank fluorescence and thus largely determined the
sensitivity of ATP detection.
b. ATP Measurement by Enzymatic Cycling
ATP was measured using enzymatic cycling of NADPH after
alkali treatment (Figure 41},., The limit of ..detection was 2x10
mol and the range 2 x 10 to 2 x 10 mol of ATP. NADP
standards produced a parallel curve but with a lower blank and so
a greater sensitivity. The limit of detection of ATP by
enzymatic cycling was improved 1000-fold over the coupled enzyme
assay method.
3. Measurement of ATP in E. coli
ATP was extracted from E_. coli using boiling Tricine
buffer and measured by the enzymatic cycling procedure discussed
earlier. The extraction procedure yielded 90% of the ATP in
standard samples so no correction was made for recovery. Since
extracts contained ATP and pyridine nucleotides, the response was
the sum of ATP and reduced pyridine nucleotides.
When 5 x 10 cells were extracted with 5 ml boiling Tricine
buffer and 0.02-ml portions of the-extract were used in the
cycling procedure, there was 5.4x10 mol of ATP equivalent/20
pi. This may be expressed as_1.35 x 10 mol ATP and reduced
pyridine nucleotides / 5 x 10 cells. When the amount of the
pyridine nucleotides were subtracted, the ATP content was 1.23 x
10 y mol/5 x 10' cells.
4. Bioluminescent ATP Assay
The bioluminescence produced when firefly luciferase (EC
1.13.12.7) reacts with ATP, luciferin, and oxygen is the basis of
an extremely sensitive method for ATP determination. A whole
technology has developed (Leach, F.R., Review in preparation ).
We examined a number of procedures for ATP determination using
firefly luciferase (1). Because of the diversity of conditions
100
-------
glucoM
ATP
ydrog»f>as»
NADPH
ADP
glucoM-6-P
6-P-gluconot* 6-P-gluconott H+ + NADPH
^
•drogtnost
NAOP*
C02
ribulo«e-5-P
Figure 40. Reaction scheme for the measurement of ATP using
enzymatic cycling of NADP "t
101
-------
>. I0
H
(0
z
UJ
H
Z
10s
j I
j I
10"
IO"12 10"" 10"
ATP (mol)
Figure 41. Enzymatic cycling assay of ATP. ATP was measured by
enzymatic cycling of NADPH produced by the coupled
enzyme reactions discussed earlier (Figure 17).
Samples (20 /ul) were incubated with 10 jjl of coupled
enzyme reagent 15 min, treated with NaOH and incubated
1 hr at 65 C^ neutralized, and carried through the
standard NADP cycling procedure.
102
-------
used and the current availability of a number of commercial
luciferase preparations, we optimized the assay (44) and compared
the properties of the various commercial preparations (41).
Conditions were established for the determination of as little as
50 fg of ATP (43). Tricine was found a superior buffer (42).
Since these publications contain the details of much of our work
on ATP analysis, we will present only minimal information here.
Figure 42 shows a typical standard curve obtained by the
procedure described in Section 5. The position of the curve can
be varied depending upon the amounts of luciferase and luciferin
used (42). The details of optimizing the assay are in the paper
by Webster and Leach (44).
5. Comparison of Enzymatic Cycling and Bioluminescent Assays
Since this laboratory has studied ATP determination
both with firefly luciferase and with enzymatic cycling, a
comparison of the two methods for the determination of ATP was
made (Table 12). The parameters compared were range,
sensitivity, cost, productivity (number of samples measured, not
counting standards), inhibitors, equipment, turnaround time (time
required to get a result and repeat an assay), and specificity.
The firefly luciferase was clearly as good or better in every
category except inhibitors, which is of vital importance in the
measurement of environmental samples. Enzymatic cycling could be
used for those samples that contain inhibitors preventing
determination by firefly luciferase; as such it is a useful
procedure.
103
-------
tooo n r
100
en
H
z:
^>
H
CD
,„
O BOEHRINGER-MANNHEIM
O LUMAC PM
• FIRELIGHT
10
100
1000
10000
fg ATP
Figure 42. Standard curve for ATP determination by firefly luci-
ferase. Light production is determined using a Packard
Model 6100 photometer maintained at 25 ±0.1 c. The
light production without added ATP is subtracted. The
reaction system (0.2 ml) contained 0.025 M Tricine
buffer, pH 7.8, 5 mM MgSO , 0.5 mM EDTA, and 0.5
DTT. With Boehringer-Mannheim luciferase, 100 pg of
bovine serum albumin and 50 jug of luciferin were added.
Various amounts of ATP contained in 20 jul were injected
into the cuvette and the light emission was counted for
30 sec after a 1-sec delay. The following luciferases
were used: Lumac's Lumit PM Lot Nos. 5041 and 5045
(130 units), Analytical Luminescence Laboratory's Fire-
light Lot No. 6621 (102 units), and Boehringer-Mannheim
luciferase Lot No. 1210301 (361 units).
mM
104
-------
TABLE 12. COMPARISON OF ATP MEASUREMENT TECHNIQUES
Parameter Luciferase Cycling
Range , 0.2 pmol - 100 pmol 0.3 pmol - 10 pmol
Sensitivity 0.2 fmol (0.1 pg) 0.1 pmol (50 pg)
Cost 6 cents/assay 9.5 cents/assay
Productivity 25/hour or 96/5 hours or
200/8-hour day_3 192/8-hour day
Inhibitors Metal ions, PO. None encountered
to date
Equipment Photometer Fluorimeter
Turnaround Time <30 minutes 5 hours
Specificity ATP only ATP, NADH, NADPH
, Useful range of ATP amounts which can be routinely measured.
Smallest amount of ATP detected by the assay.
, Based on 1979 prices.
An optimistic statement.
105
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SECTION 7
ENVIRONMENTAL SAMPLES
Since the goal of the research reported herein is
applicability to environmental samples, the described procedures
were tested with such samples. As described previously in
Section 5, several of the procedures were eliminated because they
were 1) not sensitive enough, 2) involved complicated procedures,
or 3) required equipment not commonly available. For the
remaining procedures, techniques were developed for their
application to environmental samples. These procedures and their
application are now described.
A. ATP Extractions
1. Comparison of Procedures
Many methods are available for the extraction of ATP from
various types of environmental samples. The media for the
initial extraction can be divided into four categories: 1)
aqueous buffers, 2) inorganic acids, 3) organic acids and other
solvents and 4) commercially available preparations (Table 13).
We have applied many of these extractants to produce ATP
solutions and utilized those that seemed the most promising to
liberate ATP from environmental samples.
In our hands boiling Tricine containing Mg-EDTA seems to be
the simplest and one of the more effective methods for microbial
cells. Sulfuric acid and dimethyl sulfoxide are effective for
soil samples and cells. Several commercial releasing agents, NRB
Reagent from Lumac and Releasing agent N (cat # 04-40-40) from
SAI, were not satisfactory when the manufacturers' recommended
procedure was followed. Some preliminary experiments indicate
that centrifugation to remove the soil and analysis in the
Pico-Lite luminometer might make them applicable. We have not
used the commercial releasing agents for microbial cell ATP
extraction.
The detailed procedures for boiling Tricine, DMSO and
sulfuric acid extractions of ATP are found in Section 5.
106
-------
TABLE 13. METHODS AVAILABLE FOR EXTRACTION OF ATP
Media
A. Aqueous Buffers
Condition for Extraction Recovery
Concentration Temp Time %
• M C Min
Reference
Tris
Tricine
Tricine-EDTA
Tricine-Mg
Tris-EDTA-Mg
Tris-EDTA
Glycylglyc ine-EDTA
Phospha te-EDTA
Water
0.02 or 0.05
0.05
0.05-0.002
0.05-0.01
0.05-0.002-0.01
0.05-0.002
0.05-0.002
0.05-0.002
Tap-distilled
100
100
100
100
100
100
100
100
100
2 59-65
2 59-63
1.5 93
1.5 82
1.5 89
1.5 100
1.5 100
1.5 100
2 87-100
(106)
(42)
(107)
(107)
(108)
(109,110)
B. Organic Acids and Solvents
Ammonium Formate
1
n-Bromosuccinimide-EDTA 0.01-0.01
25
25
12-21
62-96
(111)
Chloroform (saturated with aqueous)
CHC1 -Tricine
CHCl3-NaHCO
CHC13-PO "* 3
Dime thylsolf oxide
Dimethylformamide
Methanol
Ethanol
1-Butanol
2-Butanol
Oc tano 1-Bu tano 1
TCA
TCA-EDTA
C. Inorganic Acids
Sulfuric Acid
Nitric Acid
100%, 0.1
100%, 0.5
100%, 0.01
90%
90%
100%
95%
100%
100%
1:8 v/v
10%
0.51-0.017
0.6 N
1.5 N
0.1 N
25
25
25
100
100
100
100
25
0
0-4
0
78
35-46
63-72
48-65
17-20
59-94
64-86
64
68
71-99
3-13
9
80
-
-
(112,113)
(114)
(115,116)
(117)
(118)
(107)
(112,119)
(120)
(121)
Perchloric Acid
0.4
(110)
Based on ATP standard solutions with data obtained in this lab.
107
-------
2. Optimizing Conditions
Many reagents are capable of extracting ATP from
environmental samples. However, we have found that many of these
reagents subsequently interfere with the assay and consequently
must be removed. Sample size, type, concentration, and volume of
extracting reagent, methods of mixing, temperature, etc., need to
be considered in developing an optimum extraction method.
The components of the culture medium for cells interfered
with the assay; this interference was eliminated by harvesting
the cells by centrifugation and washing twice with buffer or
minimal medium. If heat is used for cell lysis the temperature
must exceed 96 C inside the test tube for complete extraction.
Heating the tubes prior to addition of the cells and injecting a
small volume of cells (usually 1 part into 10) into the buffer
helps to accomplish this. If boiling buffers are used for
extraction additional EDTA enhances the yield of ATP from
solutions; if cells are extracted, additional sensitivity is
achieved by adding both Mg and EDTA.
Dimethyl sulfoxide is a very good extractant for cellular
and soil ATP. The method is easy and very quick. The major
drawback is the inhibition of the luciferase assay by small
amounts of DMSO. This inhibition can be seen in the data
presented in Table 14. If the samples contain enough ATP to be
diluted 10- or 100-fold the DMSO inhibition can be effectively
eliminated.
Sulfuric acid is an excellent extractant for ATP from soil
and cells. Many investigators use 0.6 N H-SO., but this
concentration is too low for some types of alkaline soils, so we
use 1.5 N H2SO.. For the soils that we have tested 5-10 ml of
acid per gram or soil is optimum, and samples of from 1-5 g are
adequate. We use 2 g of soil/10 ml acid. The use of a Polytron
and wrist-action shaker facilitate the extraction of ATP from
larger soil samples (1.3-fold increase in activity for each
separately and 2.3-fold increase in measured ATP activity when
both are used) by aiding in the breakdown of soil and allowing
good mixing.
The greatest difficulty encountered with the use of acids
for extracting ATP is the neutralization of these extracts prior
to analysis since the luciferase assay is very pH sensitive.
Acids also extract unknown compounds from the soil which inhibit
the assay. We have used strong bases (NaOH, KOH, ethanolamine),
buffers containing strong bases, 0.5 M Alamine in Freon, cation
exchange columns, charcoal adsorption, and dilution as a means of
neutralizing and/or cleaning up acid extracts. Experiments have
shown that compounds of high ionic strength, ethanol, and
Alamine/Freon are inhibitory so we have avoided using them for
neutralization. The acid extract of soil is diluted 1:10 with
buffer, a small aliquot of the diluted extract is taken, and the
pH adjusted to 7.5 with 10 N KOH or 5 N ethanolamine using phenol
red indicator (0.1%). From this the amount of base required to
108
-------
TABLE 14. DMSO INHIBITION OF FIREFLY LUCIFERASE
DMSO Inhibition
% %
0.23 2
0.9 21
4.5 39
9.0 59
18.0 86
The % of DMSO is the final concentration in a 1-ml reaction
volume.
109
-------
neutralize 1 ml of extract can be calculated.
Inhibitory compounds extracted with the ATP can be removed
by cation exchange and charcoal adsorption. These methods will
be discussed later.
3. Stability of Samples
Soil samples were stored in sterile test tubes at three
temperatures, 25 , 4 , and -15 C, to determine the optimum
method of treating them before extraction. Frozen samples (-15
C) retained ATP activity equivalent to samples extracted
initially for 16 wk. Samples stored at 4 C showed more variation
from week to week than frozen samples but retained 81% of their
initial activity. Samples stored at room temperature (25 C)
retained only 22% of their activity for 1 wk. These
room-temperature samples showed a 122-147% increase in activity
during the first 72 hr suggesting that this could perhaps be used
as an amplification technique. These experiments all used the
sulfuric acid-Tricine dilution method for extraction.
The stability of sulfuric acid extracts of soil, ATP
solutions and soil spiked with ATP was monitored for 12 wk. The
soil and ATP solutions showed no significant decrease in activity
over the 12 wk when extracted, diluted 1:10 with 0.1 M Tricine,
pH 9, neutralized with 5 N ethanolamine and stored frozen. The
spiked soil samples were more variable, probably due to
adsorption of the ATP by the clay type soil used.
Tricine extracts of £. coli were stable for 1-2 wk when
stored frozen.
4. Recovery of ATP
Three different sources of ATP were used to determine
recovery when the various extraction methods were applied to
solutions, cell suspensions, or soil samples. These sources were
a known quantity of ATP in solution, ( H) -ATP, and a known
quantity of E. coli cells. These results are shown in Table 15.
These results suggest that further research is required to
optimize extraction and recovery procedures.
5._ Purification Using Charcoal and Ion Exchange
One of the major difficulties encountered in measuring ATP
extracted from soil is the concurrent extraction of several
unknown inhibitors of the firefly luciferase reaction. Dilution
techniques, charcoal adsorption of the ATP, and cation exchange
have been studied using ( H)-ATP Extract cleanup was tested
with polypropylene columns containing charcoal, Amberlite or
Dowex. The ( H)-ATP is retained (97-99%) on the Amberlite
IR-120, charcoal or Dowex 50 after one pass. The cation exchange
columns do not release the ATP readily without the use of eluants
of high ionic strength (5 N NaOH for Amberlite and 1 M sodium
110
-------
TABLE 15. RECOVERY OF ATP USING VARIOUS EXTRACTION METHODS
Method
A. Boiling Tricine
Sample
Spike
Recovery
%
B.
C.
D.
E.
F.
0.05 M, 2 min
0.05 M, 90 sec
Tricine-EDTA
Tr ic ine-Mg-EDTA
Tricine
0.05 M
Tricine-EDTA
Tricine-Mg
Tr icine-Mg-EDTA
Tr icine-Mg-EDTA
Tr icine-Mg-EDTA
Butanol-o c tano 1
Dimethyl sulfoxide
TCA-EDTA
Nitric acid (0.1
Sulfuric acid (0
(0.
(0.
(0.
(1.
(1.
(1.
N)
.2 N)
6 N)
2 N)
6 N)
5 N)
5 N)
5 N)
Solution
Solution
Solution
Solution
Solution
E. coli
IS. coli
E_. coli
E. coli
B. subtilis
Soil
E. coli
Solution I
E. coli i
Soil i
E. coli
E. coli
E. coli
E. coli
Soil
Soil
Soil
Solution
Soil
ATP standard solution 63
ATP standard solution 126
ATP standard solution 104
ATP standard solution 99
ATP standard solution 97
ATP standard solution 93
ATP standard solution 75
ATP standard solution 82
ATP standard solution 89
ATP standard solution 53
ATP standard solution 0
ATP standard solution 98
ATP standard solution 59
ATP standard solution 31
ATP standard solution 7
ATP standard solution 48
ATP standard solution 50
ATP standard solution
ATP standard solution
ATP standard solution
ATP standard solution
ATP, standard solution
j
H-ATP
3H-ATP
94
99
<2
22
63
55
12
111
-------
formate for Dowex), and these inhibit luciferase. Three washes
with 50% arnmoniacal ethanol releases 61% of the ATP from charcoal,
and since ethanol is volatile its inhibitory effects can be
minimized by evaporation. Some commercial luciferase
preparations appear less susceptible to ethanol inhibition than
others. This is being investigated further.
Column purifications are laborious and time consuming;
comparisons were made of Joatchwise charcoal cleanup versus
column purification using ( H)-ATP in the presence and absence
of soil. About 97-99% of ATP is adsorbed by charcoal whether in
columns or batch. The presence of inert cellulose does not
adversely affect the adsorption or release of J^TP and it
facilitates the packing of columns. Some 73-74% of ( H)- ATP was
either lost in the extraction procedure or adsorbed by the soil.
Less ATP is adsorbed by the charcoal in presence of soil (75-80%)
than without (95%). The recovery of ATP was the same (46-55%)
from either columns or batchwise treatments. Since the columns
are more time consuming and not significantly better, the
batchwise treatment is the method of choice.
6. Applications
ATP has successfully been measured in J3. coli, EJ^
subtilis cultures, in plant tissues, in spring water samples,
in microcosm samples and soil samples (both locally obtained
ones and core samples and column samples from Dr. John Wilson of
RSKERL, Ada, OK). The amount of ATP measured and the extraction
method used are summarized in Table 16.
B. Lipopolysaccharide Extraction
1. Soil and Core Samples
Soil was extracted with 44% phenol at 68 C in a
shaking water bath for 10 min. The pooled aqueous layers were
extracted 2 X with ether to remove residual phenol. Then the LPS
was determined as described in Section 5. Certain soil samples
were spiked with known amounts of LPS to permit an internal
standard measurement that was used for correction.
In a core sample collected by Dr. John Wilson we found 1.85
ng/g of material; in a Webster garden soil sample 85 ng/g of
soil was found.
2. Water Samples
When an extensive dilution series was made on eastern
Oklahoma spring water samples, we found that the previously
determined values were underestimated. This occured because the
signal due to gel formation determined photometrically saturates
at a low absorbance value, unlike that observed in other
spectrophotometric determinations.
112
-------
TABLE 16. APPLICATION OF VARIOUS EXTRACTION PROCEDURES FOR ATP
Extraction Method
Sample
ATP Measured * . Recovery
Boiling Tricine E. coli
B. subtilis
DMSO
E. coli
Soil sample
E_. coli
Soil sample
Wilson core
Wilson column
5.7 fg/cell
4.6 fg/cell
0.74 ng ATP/ 106 cells
3.8 ng/g
1.6 fg/cell
12 ng/g
8 ng/g
2 ng/g
84
55
39
74
59
57
63
*Corrected for inhibition and dilution.
113
-------
What is required for accurate estimation of LPS is an
extensive dilution series. When this was done we found 3.6 ng/ml
in Grotto-Eureka water and 10 pg/ml in Park Spring water. For
these environmental samples the less sensitive lots of LAL
reagents may be useful.
C. Other Procedures — Application to Spring Water
Spring water was collected in 8 locations in eastern
Oklahoma and western Arkansas in sterile and pyrogen-free
ampules. The samples were stored in a cold room (4 C) until
assayed. Samples (1.0 and 0.1 ml) of the various spring waters
were plated on nutrient agar for plate counts.
Sample preparation for colorimetric determinations consisted
of adding 10 ml of 10% trichloroacetic acid and 5 mg of
Hyflo-filter aid to 10 ml of water. The tubes were heated 15 min
at 90 C and centrifuged, and the precipitate washed with 5%
trichloroacetic acid. Centrifugation was repeated and the
combined solutions were used for nucleic acid determinations
(DNA with diphenylamine and RNA with orcinol). The protein
in the precipitate was determined by the bromosulfalein method.
The amounts of several biochemicals found in spring water
from several locations are shown in Table 17. Plating on
nutrient agar showed a bacterial content of 2-473/ml. Since
plating conditions could not be selected that would support the
growth of all types of organisms, the plate count does not
reflect the true bacterial count. There was no detectable DNA,
RNA, or protein; the amounts were <1, <1, and <2.6 /ug/ml,
respectively (that indicates the lowest amount of the substances
that could be detected). A fairly constant amount of
lipopolysaccharide was found, but these samples were probably not
diluted sufficiently; preliminary experiments suggest that this
substance remains stable in the environment for a considerable
time. ATP (determined by the firefly luciferase procedure)
varied from 7 to 58 pg/ml, and heme (determined using luminol)
from less than 2 to 200 pg/ml. Samples were measured for heme
both before and after treatment with 0.5% hydrogen peroxide so
that the response due to intact cells could be determined.
Treatment of samples with 0.5% hydrogen peroxide destroys
extracellular porphyrins and oxidizes reduced metal ions that may
give a response but does not damage viable or intact cells (34).
Three conclusions can be drawn from the heme iron data.
First, environmental samples contain measurable amounts of iron
porphyrins. Second, hydrogen peroxide treatment substantially
reduces the luminescence response of the samples. Last, there
is no clear correlation between cell numbers measured by
nutrient agar plate counts and amounts of iron porphyrins present
in the samples. Each sample of spring water stimulated bacterial
bioluminescence in the Microtox assay (data not shown).
There was no high correlation between any two of the
components measured. This lack of correlation suggests that
114
-------
TABLE 17. LEVELS OF SELECTED BIOCHEMICAL INDICATORS IN SPRING
WATER SAMPLES
Components (amount / ml)
Bacterial DNA RNA Protein LPS ATP Heme
Source Count pg pg pg fg pg pg
Whaley's 59 <1 <1 <2.5 11 58 9
Behind Speedy's 29 <1 <1 <2.5 9.9 30 20
Billy Brown's 219 <1 <1 <2.5 10.5 7 14
Grotto-Eureka 4 <1 <1 <2.5 10.5 16 17
Charlie Tanihill's 2 <1 <1 <2.5 9.8 34 200
Park Spring 5 <1 <1 <2.5 9.8 22 25
Harding-Eureka 114 <1 <1 <2.5 10.5 33 2
Murrell Home 473 <1 <1 <2.5 9.8 45 2
115
-------
additional parameters will be required to adequately characterize
these spring water samples. Further data must be obtained
before standards may be set.
116
-------
REFERENCES
1. Dermer, 0. C., V. S. Curtis, and F. R. Leach. Biochemi-
cal Indicators of Subsurface Pollution. Ann Arbor
Science, Ann Arbor, Michigan, 1980. 203 pp.
t
2. Lowry, 0. H., N. J. Rosebrough, A. L. Farr, and R. J.
Randall. Protein Measurement with the Folin Phenol
Reagent. J. Biol. Chem., 193:265-275, 1951.
3. McKnight, G. S. A Colorimetric Method for the Determina-
tion of Submicrogram Quantities of Protein. Anal. Biochem.,
78:86-92, 1977.
4. McGuire, J., P. Taylor, and L. A. Greene. A Modified Bromo-
sulfalein Assay for the Quantitative Estimation of Protein.
Anal. Biochem. 83:75-81, 1977.
5. Kutchai, H., and L. M. Geddis. Determination of Protein in
Red Cell Membrane Preparations by o-Phthalaldehyde Fluo-
rescence. Anal. Biochem., 77:315-319, 1977.
6. Butcher, E. C., and 0. H. Lowry. Measurement of Nano-
gram Quantities of Protein by Hydrolysis followed by
Reaction with Orthophthalaldehyde for Determination of
Glutamate. Anal. Biochem., 76:502-523, 1976.
7. Abraham, G. N., C. Scaletta, and J. H. Vaughan. Modifi-
ed Diphenylamine Reaction for Increased Sensitivity. Anal.
Biochem., 49:547-549, 1972.
8. Setaro, F., and C. D. G. Morley. A Rapid Colorimetric
Assay for DMA. Anal. Biochem., 81:467-471, 1977.
9. Cattolico, R. A., and S. P. Gibbs. Rapid Filter Method
for the Microfluorometric Analysis of DMA. Anal. Biochem.,
69:572-582, 1975.
10. LePecq, J.-B., and C. Paoletti. A New Fluorometric Method
for RNA and DNA Determination. Anal. Biochem., 17:100-107,
1966.
11. Boer, G. J. A Simplified Microassay of DNA and RNA Using
Ethidium Bromide. Anal. Biochem., 65:225-231, 1975.
117
-------
12. Karsten, U.f and A. Wollenberger. Improvements in the
Ethidium Bromide Method for Direct Fluorometric Estimation
of DMA and RNA in Cell and Tissue Homogenates. Anal.
Biochem., 77:464-470, 1977.
13. El-Hamalawi, A.-R., J. S. Thompson, and G. R. Barker. The
Fluorometric Determination of Nucleic Acids in Pea Seeds by
Use of Ethidium Bromide Complexes. Anal. Biochem.,
67:384-391, 1975.
14. Kapuscinski, J., and B. Skoczylas. Simple and Rapid Fluori-
metric Method for DMA Microassay. Anal. Biochem.,
83:252-257, 1977.
15. Ceriotti, G. Determination of Nucleic Acids in Animal
Tissues. J. Biol. Chem., 214:59-70, 1955.
16. Going, J., S. Wenzel, and J. Thompson. Spectrophotometrie
Determination of Phosphate by Extraction of Reduced Molyb-
doantimonylphosphoric Acid with Acetophenone-Chloroform.
Microchem. J., 20:126-131, 1975.
17. Environmental Protection Agency. Methods for Chemical
Analysis of Water and Waste. 1974. 249 pp.
18. Sottocasa, G. L., B. Kuylenstierna, L. Ernster, and A.
Bergstrand. Separation and Some Enzymatic Properties of the
Inner and Outer Membranes of Rat Liver Mitochondria. In:
Methods in Enzymology, R. W. Estabrook and M. E . Pullman,
eds. 10:448-463, 1967.
19. Rasmussen, H. N. Preparation of Partially Purified Firefly
Luciferase Suitable for Coupled Assays. In: Methods in
Enzymology, M. A. DeLuca, ed. 57:28-36, 1978.
20. Lowry, 0. H., and J. V. Passonneau. A Flexible System of
Enzymatic Analysis. Academic Press, New York, 1972. 291 pp.
21. Malamy, M. H., and B. L. Horecker. Purification and Crys-
tallization of the Alkaline Phosphatase of Escherichia
coli. Biochemistry, 3:1893-1897, 1964.
22. Torriani, A. Alkaline Phosphatase of Escherichia coli. In:
Methods in Enzymology, L. Grossman and K . Moldave, eds.
128:212-218, 1968.
23. Worthington Biochemical Corp. Worthington Enzyme Manual.
Freehold, New Jersey, 1972. pp. 41-42.
24. Worthington Biochemical Corp. Worthington Enzyme Manual.
118
-------
Freehold, New Jersey, 1972. p. 36.
25. Wbrthington Biochemical Corp. Wbrthington Enzyme Manual.
Freehold, New Jersey, 1972. pp. 7-10.
26. Curl, H., Jr., and J. Sandberg. The Measurement of Dehydro-
genase Activity in Marine Organisms. J. Mar. Res.,
19:123-138, 1961.
27. Wieser, W., and M. Zech. Dehydrogenases as Tools in the
Study of Marine Sediments. Mar.. Biol., 36:113-122, 1976.
28. Dowry, 0. H., J. V. Passonneau, D. W. Schulz, and M. K.
Rock. The Measurement of Pyridine Nucleotides by Enzymatic
Cycling. J. Biol. Chem., 236:2746-2755, 1961.
29. Chi, M. M.-Y., C. V. Lowry, and 0. H. Lowry. An Improved
Enzymatic Cycle for Nicotinamide-Adenine Dinucleotide
Phosphate. Anal. Biochem., 89:119-129, 1978.
30. Bulich, A. A., and M. W. Greene. The Use of Luminescent
Bacteria for Biological Monitoring of Water Quality. In:
International Symposium on Analytical Applications of Bio-
luminescence and Chemiluminescence, E. Schram and P.
Stanley, eds. State Printing and Publishing, Inc., Westlake
Village, CA, 1979. pp. 193-211.
31. Beckman Instruments Inc., Microbics Operations. Beckman
Microtox Model 2055 Toxicity Analyzer System. Bulletin 6984.
32. Stanley, P. E. Quantitation of Picomole Amounts of NADH,
NADPH, and FMN Using Bacterial Luciferase. In: Methods in
Enzymology, M. A. DeLuca, ed. 57:215-222, 1978.
33. Okrend, H./ R. R. Thomas, J. W. Deming, E. W. Chappelle,
and G. L. Picciolo. Methodology for Photobacteria Lucifer-
ase FMN Assay of Bacterial Levels. In: 2nd Bi-Annual ATP
Methodology Symposium, G. A. Borun, ed. SAI Technology Co.,
San Diego, CA., 1977. pp. 525-546.
34. Thomas, R. R., G. L. Picciolo, E. W. Chappelle, E. L.
Jeffers, and R. E. Taylor. Use of the Luminol Assay for the
Determination of Bacterial Iron Porphyrins: Flow Techni-
ques for Wastewater Effluent. In: 2nd Bi-Annual ATP Metho-
dology Symposium, G.A. Borun, ed. SAI Technology Co., 1977.
pp. 569-579.
35. Neufeld, H. A., C. J. Conklin, and R. D. Towner. Chemilumin-
escence of Luminol in the Presence of Hematin Compounds.
Anal. Biochem., 12: 303-309, 1965.
119
-------
36. Oleniacz, W. S., M. A. Pisano, M. H. Rosenfeld, and R. L.
Elgart. Chemiluminescent Method for Detecting Microorganisms
in Water. Environ. Sci. Technol., 2:1030-1033, 1968.
37. Searle, N. D. Applications of Chemiluminescence to
Bacterial Analysis. In: Analytical Applications of Bio-
luminescence and Chemiluminescence, E. W. Chappelle and
G. L. Picciolo, eds. NASA SP-388, Washington, D. C., 1975.
pp. 95-103.
38. Ewetz, L., and A. Thore. Factors Affecting the Specificity
of the Luminol Reaction with Hematin Compounds. Anal.
Biochem., 71:564-570, 1976.
39. Cohen, E., ed. Biomedical Applications of the Horseshoe
Crab (Limulidae). Alan R. Liss, Inc., New York, 1981. 688 pp.
40. Associates of Cape Cod, Inc. Limulus Amebocyte Lysate
(Pyrotell) for The Detection and Quantitation of Endotoxins.
Woods Hole, Mass., 1981.
41. Webster, J. J., J. C. Chang, J. L. Howard, and F. R. Leach.
Some Characteristics of Commercially Available Firefly Luci-
ferase Preparations. J. Appl. Biochem., 1:471-478, 1979.
42. Webster, J. J., J. C. Chang, E. R. Manly, H. 0. Spivey, and
F. R. Leach. Buffer Effects on ATP Analysis by Firefly
Luciferase. Anal. Biochem., 106:7-11, 1980.
43. Webster, J. J., J. C. Chang, and F. R. Leach. Sensitivity
of Various Commercial Luciferase Preparations. J. Appl.
Biochem., 2:516-517, 1980.
44. Webster, J. J., and F. R. Leach. Optimization of the Fire-
fly Luciferase Assay for ATP. J. Appl. Biochem., 2:469-479,
1980.
45. Mehl, J. W. The Biuret Reaction of Proteins in the Presence
of Ethylene Glycol. J. Biol. Chem., 157:173-180, 1945.
46. Gornall, A. G., C. J. Bardawill, and M. M. David. Determin-
ation of Serum Proteins By Means of the Biuret Reaction. J.
Biol. Chem., 177:751-766, 1949.
47. Ma, T. S., and G. Zuazaga. Micro-Kjeldahl Determination of
Nitrogen, a New Indicator and an Improved Rapid Method.
Ind. Eng. Chem., Anal. Ed., 14:280-282, 1942.
48. Peterson, G. L. Review of the Folin Phenol Protein Quanti-
tation Method of Lowry, Rosebrough, Farr and Randall. Anal.
Biochem., 100:201-220, 1979.
120
-------
49. Bradford, M. M. A Rapid and Sensitive Method for the Quan-
titation of Microgram Quantities of Protein Utilizing the
Principle of Protein - Dye Binding. Anal. Biochem.,
72:248-254, 1976.
50. Sedmak, J. J., and S. E. Grossberg. A Rapid, Sensitive,
and Versatile Assay for Proteins Using Coomassie Brilliant
Blue G 250. Anal. Biochem., 79:544-552, 1977.
51. Pierce, J. and C. H. Suelter. An Evaluation of the
Coomassie Brilliant Blue G-250 Dye-Binding Method for Quan-
titative Protein Determination. Anal. Biochem., 81:478-480,
1977.
52. Van Kley, H. and S. M. Hale. Assay for Protein by Dye
Binding. Anal. Biochem., 81:485-487, 1977.
53. Chiappelli, F., A. Vasil, and D. F. Haggerty. The Protein
Concentration of Crude Cell and Tissue Extracts as Estimated
by the Method of Dye Binding: Comparison with the Lowry
Method. Anal. Biochem., 94:160-165, 1979.
54. Greif, R. L. Use of Bromosulphalein for the Measurement of
Proteolytic Activity. Proc. Soc. Exp. Biol. Med.,
75:813-815, 1950.
55. Nayyar, S. N., and D. Click. Studies in Histochemistry XXXI.
A Method for the Determination of Protein in Millimicrogram
Quantities. J. Histochem. Cytochem., 2:282-290, 1954.
56. Bonting, S. L., and M. Jones. Determination of Microgram
Quantities of Deoxyribonucleic Acid and Protein in Tissues
Grown "in Vitro". Arch. Biochem. Biophys., 66:340-353, 1957.
57. Flores, R. A Rapid and Reproducible Assay for Quantitative
Estimation of Proteins Using Bromophenol Blue. Anal.
Biochem., 88:605-611, 1978.
58. Schultz, R. M. and P. M. Wassarman. ( H)-Dansyl Chloride:
A Useful Reagent for the Quantitation and Molecular Weight
Determination of Nanogram Amounts of Protein. Anal.
Biochem., 77:25-32, 1977.
59. Burzynski, S. R. Quantitative Analysis of Amino Acids and
Peptides in the Femtomolar Range. Anal. Biochem., 65:93-99,
1975.
60. Bohlen, P., S. Stein, W. Dairman, and S. Udenfriend. Fluoro-
metric Assay of Proteins in the Nanogram Range. Arch.
Biochem. Biophys., 155:213-220, 1973.
121
-------
61. Roth, M. Fluorescence Reaction for Mino Acids. Anal.
Chem., 43:880-882, 1971.
62. Weidekamm, E., D. F. H. Wallach, and R. Fluckiger. A New
Sensitive, Rapid Fluorescence Technique for the Determina-
tion of Proteins in Gel Electrophoresis and in Solution.
Anal. Biochem., 54:102-114, 1973.
63. Benson, J. R., and P. E. Hare. o-Phthalaldehyde: Fluoro-
genic Detection of Primary Amines in the Picomole Range.
Comparison with Fluorescamine and Ninhydrin. Proc. Nat.
Acad. Sci. USA, 72: 619-622, 1975.
64. Butcher, E. C., and 0. H. Lowry. Measurement of Nanogram
Quantities of Protein by Hydrolysis followed by Reaction
with Orthophthalaldehyde or Determination of Glutamate.
Anal. Biochem., 76:502-523, 1976.
65. Goldberg, M. L. Quantitative Assay for Submicrogram Amounts
of Protein. Anal. Biochem., 51:240-246, 1973.
66. Kinoshita, T., F. linuma, and A. Tsuji. Microassay of
Proteins on Membrane Filter in the Nanogram Range Using
Cycloheptaamylose-Dansyl Chloride Complex. Anal. Biochem.,
66:104-109, 1975.
67. Viets, J. W., W. M. Deen, J. L. Troy, and B. M. Brenner.
Determination of Serum Protein Concentration in Nanoliter
Blood Samples Using Fluorescamine or £-Phthalaldehyde.
Anal. Biochem., 88: 513-521, 1978.
68. Robrish, S. A., C. Kemp, and W. H. Bowen. The Use of the
o-Phthalaldehyde Reaction as a Sensitive Assay for Protein
and to Determine Protein in Bacterial Cells and Dental
Plaque. Anal. Biochem., 84:196-204, 1978.
69. Castell, J. V., M. Cervera, and R. Marco. A Convenient
Micromethod for the Assay of Primary Amines and Proteins
with Fluorescamine. A Reexamination of the Conditions of
Reaction. Anal. Biochem., 99:379-391, 1979.
70. Mroz, E. A., and C. Lechene. Fluorescence Analysis of Pico-
liter Samples. Anal. Biochem., 102:10-96, 1980.
71. Leaback, D. H., and S. Creme. A New Experimental Approach
to Fluorometric Enzyme Assays Employing Disposable Micro-
reaction Chambers. Anal. Biochem., 106:314-321, 1980.
72. Dische, Z. Uber einige neue charakteristische Farbreaktionen
der Thymonukleinsaure und eine Mikromethode zur Bestimmung
122
-------
derselben in tierischen Organen mit Hilfe dieser Reaktionen.
Mikrochemie, 8:4-32, 1930.
73. Burton, K. A Study of the Conditions and Mechanism of the
Diphenylamine Reaction for the Colorimetric Estimation of
Deoxyribonucleic Acid. Biochem. J., 62:315-323, 1956.
74. Giles, K. W., and A. Myers. An Improved Diphenylamine Method
for the Estimation of Deoxyribonucleic Acid. Nature, 206:93,
1965.
75. Richards, G. M. Modifications of the Diphenylamine Reaction
Giving Increased Sensitivity and Simplicity in the Estima-
tion of DMA. Anal. Biochem., 57:369-376, 1974.
76. Labarca, C., and K. Paigen. A Simple, Rapid, and Sensitive
DMA Assay Procedure. Anal. Biochem., 102:344-352, 1980.
77. Martin, R. F., D. C. Donohue, and L. R. Finch. New Analy-
tical Procedure for the Estimation of DMA with £-Nitro-
phenylhydrazine. Anal. Biochem., 47:562-574, 1972.
78. Gold, D. V., and D. Shochat. A Rapid Colorimetric Assay for
the Estimation of Microgram Quantities of DMA. Anal.
Biochem., 105:121-125, 1980.
79. Cesarone, C. F., C. Bolognesi, and L. Santi. Improved Micro-
fluorometric DNA Determination in Biological Material Using
33258 Hoechst. Anal. Biochem., 100:188-197, 1979.
80. Brunk, C. F., K. C. Jones, and T. W. James. Assay for Nano-
gram Quantities of DNA in Cellular Homogenates. Anal. Bio-
chem., 92:497-500, 1979.
81. Shmookler Reis, R. J. A Sensitive Microassay of Nucleic
Acids. Anal. Biochem., 90:821-823, 1978.
82. Markovits, J., P. Roques, and J.-B. LePecq. Ethidium Dimer:
A New Reagent for the Fluorimetric Determination of Nucleic
Acids. Anal. Biochem., 94:259-264, 1979.
83. Bial, M. Die Diagnose der Pentosurie. Deut. Med. Wochschr.,
28:253-254, 1902.
Bial, M. Uber die Diagnose der Pentosurie mit dem von mir
angegebenen Reagens. Deut. Med. Wochschr., 29:477-478. 1902.
84. Barrenscheen, H. K., and A. Peham. Zur photometrischen
Bestimmung der Nucleoside und Nucleotide auf Grund der
Orcinreaktion. Z. Physiol. Chem., 272:81-86, 1942.
123
-------
85. Dische, Z. Color Reactions of Nucleic Acid Components. In:
The Nucleic Acids I , E. Chargaff and J. N. JDavidson, eds.
Academic Press, Inc., New York, 1955. pp. 285-305.
86. Almog, R., and T. L. Shirey. A Modified Orcinol Test for the
Specific Determination of RNA. Anal. Biochem., 91:130-137,
1978.
87. Beers, P. C., and J. L. Wittliff. Measurements of DMA and
RNA in Mammary Gland Homogenates by the Ethidium Bromide
Technique. Anal. Biochem., 63:433-441, 1975.
88. Fiske, C. H., and Y. SubbaRow. Phosphocreatine. J. Biol.
Chem., 81:629-679, 1929.
89. Lazarus, L. H., and S.-C. Chou. Modifications of the Anal-
ysis of Phosphorus and Kinetics of the Reaction. Anal.
Biochem., 45:557-566, 1972.
90. Chen, P. S., Jr., T. Y. Toribara, and H. Warner. Microdeter-
mination of Phosphorus. Anal. Chem., 28:1756-1758, 1956.
91. Brunette, M. G., N. Vigneault, and G. Danan. A New Fluoro-
metric Method for Determination of Picomoles of Inorganic
Phosphorus. Application to the Renal Tubular Fluid. Anal.
Biochem., 86: 229-237, 1978.
92. Lowry, 0. H., N. R. Roberts, and J. I. Kapphahn. The Fluoro-
metric Measurement of Pyridine Nucleotides. J. Biol. Chem.,
224:1047-1064, 1957.
93. Lee, J., and H. H. Seliger. Quantum Yields of the Luminol
Chemiluminescence Reaction in Aqueous and Aprotic Solvents.
Photochem. Photobiol., 15:227-237, 1972.
94. Sullivan, J. D., Jr., F. W. Valois, and S. W. Watson. Endo-
toxins: The Limulus Amebocyte Lysate System. In: Mechan-
isms in Bacterial Toxicology, A. W. Bernheimer, ed. John
Wiley and Sons, New York, 1976. 263 pp.
95. Jorgensen, J. H., and R. F. Smith. Preparation, Sensitivity,
and Specificity of Limulus Lysate for Endotoxin Assay.
Appl. Microbiol., 26:43-48, 1973.
96. Frauch, P. Slide Test as a Micromethod of a Modified
Limulus Endotoxin Test. J. Pharma. Sci., 63:808-809, 1974.
97. Flowers, D. J. A Microtechnique for Endotoxin Assay by
Using Limulus Lysate. Med. Lab. Sci., 36:171-176, 1979.
98. Watson, S. W., J. Novitsky, H. L. Quinby, and F. W. Valois.
124
-------
Determination of Bacterial Number and Biomass in the Marine
Environment. Appl. Environ. Microbiol., 33:940-946, 1977.
99. Tai, J. Y., and T.-Y. Liu. Studies on Limulus Amoebocyte
Lysate, Isolation of Pro-Clotting Enzyme. J. Biol. Chem.,
252:2178-2181, 1977.
100. Tai, J. Y., R. C. Seid, R. D. Huhn, and T.-Y. Liu. Studies
on Limulus Amoebocyte Lysate II, Purification of the
Coagulogen and the Mechanism of Clotting. J. Biol.
Chem., 252:4773-4776, 1977.
101. Liang, S.-M., T. P. Sakmar, and T.-Y. Liu. Studies on
Limulus Amoebocyte Lysate III. Purification of an
Endotoxin-Binding Protein from Limulus Moebocyte
Membranes. J. Biol. Chem., 255:5586-5590, 1980.
102. Evans, T. M., J. E. Schillinger, and D. G. Stuart. Rapid
Determination of Bacteriological Water Quality by Using
Limulus Lysate. Appl. Environ. Microbiol., 35:376-382,
1978.
103. Goto, H., and S. Nakamura. Dry Up Method as a Revised
Limulus Test with a New Technique for Gelation Inhibitor
Removing. Japan. J. Exp. Med., 49:19-25, 1979.
104. Munford, R. S. Quantitative Limulus Lysate Assay for
Endotoxin Activity: Aggregation of Radioiodinated Coagulogen
Monomers. Anal. Biochem., 91:509-515,1978.
105. Maeda, M., and N. Taga. Chromogenic Assay Method of Lipo-
polysaccharide (LPS) for Evaluating Bacterial Standing Crop
in Seawater. J. Appl. Bacteriol., 47:175-182, 1979.
106. Cheer, S., J. H. Gentile, and C. S. Hegre. Improved Methods
for ATP Analysis. Anal. Biochem., 60:102-114, 1974.
107. Lundin, A., and A. Thore. Comparison of Methods for Extrac-
tion of Bacterial Adenine Nucleotides Determined by Firefly
Assay. Appl. Microbiol., 30:713-721, 1975.
108. Bullied, N. C. An Improved Method for the Extraction of
Adenosine Triphosphate from Marine Sediment and Seawater.
Limnol. Cceanogr., 23:174-178, 1978.
109. Aledort, L. M., R. I. Weed, and S. B. Troup. Ionic Effects
on Firefly Bioluminescence Assay of Red Blood Cell ATP.
Anal. Biochem., 17:268-277, 1966.
125
-------
110. Bagnara, A. S., and L. R. Finch. Quantitative Extraction
and Estimation of Intracellular Nucleoside Triphosphates of
Escherichia coli. Anal. Biochem., 45:24-34, 1972.
111. MacLeod, N. H., E. W. Chappelle, and A. M. Crawford. ATP
Assay of Terrestrial Soils: A Test of an Exobiological Ex-
periment. Nature, 223:267-268, 1969.
112. Forsberg, C. W., and K. Lam. Use of Adenosine S'-Triphos-
phate as an Indicator of the Microbiota Biomass in Rumen
Contents. Appl. Environ. Microbiol., 33:528-537, 1977.
113. Dhople, A. M., and J. H. Hanks. Quantitative Extraction of
Adenosine Triphosphate from Cultivable and Host-Grown Micro-
bes: Calculation of Adenosine Triphosphate Pools. Appl.
Microbiol., 26:399-403, 1973.
114. Paul, E. A., and R. L. Johnson. Microscopic Counting and
Adenosine 5'-Triphosphate Measurement in Determining Micro-
bial Growth in Soils. Appl. Environ. Microbiol., 34:263-269,
1977.
115. Afghan, B. K., J. F. Ryan, and R. S. Tobin. Improved Method
for Quantitative Measurement of Adenosine Triphosphate in
Lake Waters, Activated Sludges and Sediments. In: Environ-
mental Analysis, G. W. Ewing, ed. Academic Press, Inc.,
New York, 1977. pp. 29-45.
116. Mathis, R. R. and 0. R. Brown. ATP Concentration in
Escherichia coli during Oxygen Toxicity. Biochim.
Biophys. Acta, 440:723-732, 1976.
117. St. John, J. B. Determination of ATP in Chlorella with the
Luciferin-Luciferase Enzyme System. Anal. Biochem., 37:409-
416, 1970.
118. Sharpe, A. N., M. N. Woodrow, and A. K. Jackson. Adenosine-
triphosphate (ATP) Levels in Foods Contaminated by Bacteria.
J. Appl. Bact., 33:758-767, 1970.
119. Lee, C. C., R. F. Harris, J. D. Williams, D. E. Armstrong,
and J. K. Syers. Adenosine Triphosphate in Lake Sediments: I.
Determination. Soil Sci. Soc. Amer. Proc., 35:82-91, 1971.
120. Eiland, F. An Improved Method for Determination of Adenosine
Triphosphate (ATP) in Soil. Soil Biol. Biochem., 2:31-35,
1979.
126
-------
121. Knust, E. A., E. W. Chappelle, and G. L. Picciolo. A Compar-
ison of Certain Extraction Agents for Extracting of Adenosine
Triphosphate (ATP) from Microorganisms for Use in the Fire-
fly Luciferase ATP Assay. In: Analytical Applications of
Bioluminescence and Chemiluminescence, E. W. Chappelle and
G. L. Picciolo, eds. NASA SP-388, Washington, D. C., 1975.
pp. 27-34.
127
-------