Methods for the Measurement of a Bacterial Enzyme Activity in Cell


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Biological Procedures Online · Vol. 1 No. 1 · May 14, 1998 ·
Methods for the Measurement of a Bacterial Enzyme Activity in Cell
Lysates and Extracts
Brendan P. Burns1*, George L. Mendz2 , Stuart L. Hazell1
1School of Microbiology and Immunology and 2School of Biochemistry and Molecular Genetics, The
University of New South Wales, Sydney 2052, Australia. *To whom correspondence should be
addressed. E-mail:
The kinetic characteristics and regulation of aspartate carbamoyltransferase activity were studied in
lysates and cell extracts of Helicobacter pylori by three different methods. Nuclear magnetic resonance
spectroscopy, radioactive tracer analysis, and spectrophotometry were employed in conjunction to
identify the properties of the enzyme activity and to validate the results obtained with each assay. NMR
spectroscopy was the most direct method to provide proof of ACTase activity; radioactive tracer
analysis was the most sensitive technique and a microtitre-based colorimetric assay was the most costand
time-efficient for large scale analyses. Freeze-thawing was adopted as the preferred method for cell
lysis in studying enzyme activity in situ. This study showed the benefits of employing several different
complementary methods to investigate bacterial enzyme activity.
Studies of bacterial enzyme activities provide fundamental information relevant to microbial physiology,
and to a more complete understanding of cell metabolism, bacterial evolution, and to the interactions
that occurs between bacterium and host. Characterizing enzyme activities helps to elucidate regulatory
mechanisms and pathways in an organism, and to identify proteins essential for cell survival. The ability
to investigate enzyme activities in whole cell lysates or crude extracts is important for the initial
identification of a particular activity, and for obtaining information on enzyme function in an
environment close to the cellular milieu.
The activities of enzymes in the intracellular milieu depend on their intrinsic physico-chemical properties
and on their interactions with other cellular components. Modulation of enzyme activities of functional
relevance occurs not only among the constituents of recognised multienzyme clusters, but also among
soluble enzymes and other cellular components (1, 2). Proteins have also been known to be more stable
in concentrate than in dilute solutions (3). The basic mechanisms of action of soluble enzymes
established in purified preparations are not expected to change from the in vivo conditions, however
their activities are modulated by the intracellular milieu. Thus, in situ investigations of enzyme activities
may reveal properties which would otherwise remain undetected.
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Biological Procedures Online · Vol. 1 No. 1 · May 14, 1998 ·
The investigation of enzyme activity with a variety of methods results in synergies in the understanding
of the characteristics of an enzyme. Application of different techniques can validate data obtained by one
method by the results of the others, yield more specific information on a particular aspect of a system,
and provide results in a range of assay conditions ordinarily not available to a single method. The
benefits and disadvantages of particular techniques, such as sensitivity, efficiency, and cost, need to be
considered to determine which are the most useful methods for a specific situation.
In a previous investigation (4), we used three different techniques to study aspartate
carbamoyltransferase (ACTase) activity in Helicobacter pylori. This bacterium is an important human
pathogen (5, 6), and the enzyme of interest is a key regulatory step in bacterial de novo pyrimidine
nucleotide metabolism (7). As this enzyme appears to be essential for the survival of the bacterium, it
provides a potential site for therapeutic intervention. Consequently, an in-depth understanding of the
enzyme activity and regulation in situ would serve toward more rational design of therapeutic agents.
Preparation of cell free extracts
Cells were grown at 37°C in an atmosphere of 10% CO2 in air, and 95% humidity. Bacteria were
harvested during the late logarithmic growth phase from agar plates into 0.1 M Tris buffer (pH 8.0). H.
pylori cells change shape as cultures grow older. The bacteria have spiral-rod forms when they are in log
phase, and this shape becomes spherical with the formation of coccoids which tend to aggregate in the
stationary phase. Cultures on plates will produce cells of different ages, but based on the morphology of
the bacteria, phase contrast microscopy allows easy determination of the predominant growth phase.
Under our experimental conditions, inspection of cultures showed that at 24h growth more than 95% of
the cells were in the spiral-rod form, and after 72h growth more than 95% of the cells were coccoids. It
should be noted that the formation of coccoid aggregates poses a problem for the estimation of cell
growth by spectrophotometry at 600 nm. The number of scattering centres can decrease significantly
upon aggregation, and the measurements could underestimate bacterial growth.
Harvested cells suspended in Tris buffer were centrifuged at 17,000g for 8 min at 4°C. The resulting
pellet was washed twice and resuspended in the same buffer. Cells were lysed by twice freezing in liquid
nitrogen and thawing the suspensions, which were allowed to thaw completely before re-freezing. To
obtain crude extracts, the lysates were centrifuged at 27,000g for 8 min at 4°C, and the supernatant
carefully separated from the pellet made up of cell-envelope debris. The lysates and the crude extracts
could be stored at -20°C for several months without loss of activity.
Measurement of ACTase activity
Nuclear magnetic resonance spect roscopy (NMR). The unique potential of NMR spectroscopy for
monitoring simultaneously the concentrations of several metabolites in complex milieux makes it one of
the most powerful techniques available to carry out this type of study.
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For NMR measurements, lysates or cell-free extracts were resuspended in 0.15 M NaCl constituted in
5:1 H2O/2H2O buffer mixtures to provide deuterium frequency lock for the spectrometer. Substrate
concentrations were 40 mM L-aspartate and 50 mM carbamoyl phosphate, in 0.1 M HEPES buffer (pH
8.0). The reaction was started by adding 100 μl of cell free extract to a total sample volume of 600 μl.
To allow for efficient dispensing of the assay mixture into the 5 mm narrow bore NMR tube (Wilmad,
Buena, NJ), the lysate or extract suspension and the substrates were mixed in an Eppendorf tube;
diluting viscous cell lysates helped to place them into the tube.
Free induction decays were collected using a Bruker AM-500 spectrometer, operating in the Fourier
transformation mode. Measurements were carried out at 37°C, and sequential spectra were acquired
automatically at 500.11 MHz with presaturation of the water resonance. The instrumental parameters
were: spectral width 5347 Hz, memory size 8 K, recycling time 3.5 s, number of transients 144, and
pulse angle 50° (8 μs). To improve signal-to-noise, exponential filtering of 1 Hz was applied prior to
Fourier transformation. Depending on the viscosity of the sample and the concentrations of the
substrates, it is often advisable to employ resolution-enhancing window functions at the expense of
losing some signal intensity; for example, a Gaussian window function with typical parameters: -1 Hz
line broadening, and 0.19 Gaussian parameter.
The time evolution of the utilisation of substrates and appearance of product was followed by acquiring
sequential spectra of the reactions. Progress curves were obtained by measuring the integrals of
substrate and product resonances at each point in time. Maximal rates were calculated from good fits
(correlation coefficients ³ 0.99) of the data to straight lines for 30 min of the reactions.
Radioactive tracer analysis . ACTase activity was also determined using a radioactive assay that
measures the incorporation of [14C]carbamoyl phosphate into carbamoyl aspartate (8). Typical enzyme
assay conditions were 40 mM L-aspartate, 0.4 mM carbamoyl phosphate containing [14C]carbamoyl
phosphate (0.1 μCi μmole-1), Tris buffer (0.1 M, pH 8.0), and cell-free extract in a final volume of 200
μl. The reaction was initiated by adding 10 μl of cell-free extract, and incubated for 10 min at 37°C in a
water bath. Reactions were terminated by adding 100 μl of 3 M formic acid and heating at 80°C for 6
min, during which time any unreacted [14C]carbamoyl phosphate decomposed into phosphate and carbon
dioxide, and the label was evaporated as 14CO2. Heating for 6 minutes was found to be the optimum;
less time was insufficient for complete conversion to 14CO2 of the unreacted [14C]carbamoyl phosphate,
and longer times were unnecessary. Ten milliliters of scintillation fluid (2,5-diphenyloxazole/toluene;
0.5% w/v) were added, and the radioactive decay measured on a Packard Tricarb scintillation counter
(Packard Instrument Co., USA).
Spectrophotometry. A colorimetric measurement of carbamoyl aspartate production using a microtitre
protocol was employed for kinetic studies of the enzyme reaction (9), and all measurements were
performed in triplicate. The method was based on that of Prescott and Jones (10) which uses a
monoxime and antipyrine colour reagent for the detection of CAA. Two parts antipyrine (5 g/l in 50%
(v/v) sulfuric acid) were mixed with 1 part monoxime (8 g/l in 5% (v/v) acetic acid). The colour reagent
B.P. Burns et al 20
Biological Procedures Online · Vol. 1 No. 1 · May 14, 1998 ·
was prepared immediately before use. Owing to the instability of carbamoyl phosphate (11), it is
essential that solutions are prepared fresh before each reaction. The reaction mixture contained the same
concentrations of substrates and buffer as described above but without the [14C]carbamoyl phosphate, in
a final volume of 200 μl. To start the enzyme reaction 50 μl of freshly prepared carbamoyl phosphate
was added to the wells, and the plate incubated for 10 min at 37°C. One hundred microlitres of the color
reagent was then added to each well to stop the reaction, and the plate incubated in the dark in a water
bath at 60°C for 2 h. Else and co-workers (9) recommend covering the plate with adhesive film while
incubating; however when this was done water droplets would sometimes form on the underside of the
film and mix into the wells. To avoid this problem another microtitre plate was used as a cover. The
plate was then allowed to cool to room temperature in the dark for 15 minutes, and the absorbance
measured at 450 nm.
This method was adapted in the present study to provide an activity stain for ACTase in native PAGE
gels. Native proteins were separated by modifying the procedure of Laemmli (12), where gel
electrophoresis was performed under non-reducing and non-denaturing conditions. The gel was then
sliced into 1mm sections, and each section placed into a microtitre well containing the substrates of the
ACTase reaction. An identical lane of electrophoretically separated proteins were also sliced and then
incubated without aspartate, as a negative control. The microtitre plate was then processed as before.
ACTase activity and cellular localisation
ACTase activity could be localised to the soluble fraction obtained by freeze-thawing cell suspensions
and separating the cell envelope and cytosolic fractions by high-speed centrifugation. The use of freezethawing
as a technique of cell lysis has several advantages over other methods in the study of enzyme
activities. Sonication may not be desirable on account of the heat generated at the tip of the probe, that
could result in denaturation of proteins. It has also been shown that sonication gives rise to H. and OH.
radicals which can lead to protein fragmentation (13). Similarly, chemical lysis may affect enzyme
activity detrimentally by disrupting enzyme complexes, etc. Some techniques, such as sonication and
rupturing with glass beads, employ relatively harsh disruption procedures whose action may dislodge
membrane-associated proteins, and thus yield positive activity in the soluble fraction. Freeze-thawing is
an efficient and relatively ‘gentle’ process of cell disruption, which appears to open holes in the cell
envelope and have minimal effects on its overall integrity, ensuring little shearing of membrane-bound
proteins and good recovery of soluble proteins (14, 15). It is important when lysing cells to take into
account not only the efficiency of the lysis procedure in breaking open the cells, but also of any effects
the method may have on processes one may subsequently wish to study. For these reasons freezethawing
was adopted as the preferred method for producing cell free extracts for studying ACTase
activity in H. pylori.
Different enzyme assays for ACTase study in H. pylori
ACTase properties were studied in situ in cell-free extracts to obtain information on enzyme function in
an environment that was closer to the bacterium’s physiological conditions. There are numerous and
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Biological Procedures Online · Vol. 1 No. 1 · May 14, 1998 ·
unique advantages in studying enzyme activities in purified preparations, but the use of simpler milieux
may mask properties such as protein-protein interactions, that could be observed in more complex
conditions. Investigation of an enzyme activity in crude extracts allows for its characterisation, and at
the same time retains the possibility of discovering some of its interactions with other cell components.
Three different methods of studying ACTase activity were employed in this investigation. The results
obtained by each technique were compared with the data from the others, to validate them and/or gather
additional information. Comparison of the rates in Table 1, indicated that each method represented a
valid procedure for studying ACTase activity in H. pylori.
Table 1. Comparison of ACTase activity determined by three different methods
Assay method ACTase activity (μmole min-1 mg protein-1)
NMR spectroscopy 0.228 ± .029
Radioactive tracer analysis 0.256 ± .023
Spectrophotometry 0.272 ± .024
NMR spectroscopy was the first method employed to identify ACTase in H. pylori. 1H-NMR spectra
showing the time-course of substrates and products of the ACTase reaction in incubations with H. pylori
cell-free extracts were given in Figure 1 of the original article (1). Decrease of the peaks of the aspartate
substrate, and appearance of peaks corresponding to a metabolic product were observed. The protons of
the substrate carbamoyl phosphate are in chemical exchange with those of the aqueous solvent at rates
which render them invisible in the 1H-NMR spectrum under the experimental conditions employed. The
1H-NMR resonances arising from H. pylori lysates or cell-free extracts were very small compared to
those of the aspartate and the product, thus it was possible to follow the evolution of these metabolites
over time. Assignment of the resonances from the product carbamoyl aspartate (CAA) was achieved by
adding this metabolite to assay mixtures and comparing the spectral position of its resonances with those
of the product in the bacterial preparations.
Metabolites have NMR spectral signatures characterized by their chemical structures. In the case of
CAA the resonances arising from a-CH and b-CH2 groups of the aspartate moiety have chemical shifts
and coupling constants different from those of the amino acid aspartate. Thus, it was possible to make
an unambiguous assignment of CAA. This definite identification of a reaction product, is a feature
lacking in other enzyme assays, particularly when complex preparations are assayed. For example, the
radioactive or colorimetric assay measures the formation of a radioactive product and a ureido product,
respectively, without yielding sufficient information about its identity. Although proper controls could
ensure the validity of these methods, NMR spectroscopy gave direct evidence for the presence of
ACTase activity. However, this technique has limitations, some of which are: a) sensitivity, because often
relatively high concentrations of substrates are needed to obtain clear NMR signals; b) spectral overlap,
under certain conditions some peaks may appear superimposed on others thus masking specific effects;
and c) cost of operating NMR spectrometers, which may preclude their use in large scale enzyme
analysis. Notwithstanding its limitations, this technique has proven most effective in the study of enzyme
characteristics in H. pylori (4, 16-18).
Radioactive tracer analysis was another useful method to study ACTase activity in cell-free extracts. A
B.P. Burns et al 22
Biological Procedures Online · Vol. 1 No. 1 · May 14, 1998 ·
main advantage of this technique is its sensitivity; measurement of radioactive decays allowed for
determination of ACTase activity at relatively small enzyme concentrations. This could prove especially
useful in studies on purified proteins, where yield is often quite low. Radioactive tracer analyses
provided information on ACTase activity that could not be obtained by NMR analysis or the
colorimetric assay. Specifically, to study the effects of carbamoyl aspartate, the ACTase end-product, on
enzyme activity. In these experiments the added exogenous CAA will mask the product formed through
the activity of the enzyme in both the NMR and colorimetric assays; the higher the concentration of
added CAA, the less accurate are the measurements of the activity of the enzyme by NMR spectroscopy
or spectrophotometry. The effects of CAA on ACTase activity were assessed confidently employing
radioactive tracer analysis and, as shown in Figure 1, the observed product-inhibition of the enzyme
activity was consistent with the inhibitory effects CAA had on H. pylori growth.
0 10 20 30 40 50 60
[Carbamoyl aspartate] (mM)
ACTase activity (% control)
Viability (% control)
Figure 1. Effects of carbamoyl aspartate (CAA) on ACTase activity and viability of H. pylori. The
activity or viability are taken as 100% in the absence of CAA. (·) ACTase activity; ( ) H. pylori
Because the labelled carbamoyl phosphate is used only in trace amounts and is an unstable compound it
is very important to monitor its purity continuously. At one point in the work, significant radioactive
counts were measured in samples only containing [14C]carbamoyl phosphate, even after addition of
strong acid and heating. As mentioned in the methods, this treatment should allow for the removal of
any unreacted [14C]carbamoyl phosphate as 14CO2. The finding indicated the presence of an acid-stable
contaminant in the stock [14C]carbamoyl phosphate. Apart from the obvious expense in obtaining fresh
radioactive substrate, the possibility of further contamination and false positive results suggested this
method may not the most appropriate for large scale characterisation of ACTase activity. Another less
technical disadvantage of this technique is the safety in the handling of radioactive products. Although
B.P. Burns et al 23
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due care should prevent any problems, the element of risk involved may favour the use of a technique
with less hazardous materials.
The use of a micro-titre based colorimetric assay provided a third method for the study of ACTase
activity, and the most useful for large scale measurements. The method was essentially as described
previously (9), but several findings proved useful in optimising the assay for this work. Although the
authors suggest that the colour development reagents can be stored for a short period of time (10), it
was found in the present study that the most consistent results were obtained when the reagents were
made fresh immediately prior to use. In constructing the CAA standard curve in Figure 2, it was found
that the absorbance values increased linearly with respect to CAA up to 0.15 μmole CAA, slightly more
than the 0.125 μmole reported previously. At CAA quantities higher than this, the readings suffered
from noise and were no longer linear with respect to CAA. This was important in assessing that this
technique could not be used to study the effects of CAA on ACTase activity, as described above. For
CAA quantities below 0.015 μmole, the readings were below the limits of sensitivity of the assay.
Comparison of colour development times between 2 and 7 h, revealed that a two hour development
time yielded the most consistent and reproducible results.
0.00 0.05 0.10 0.15 0.20
[Carbamoyl aspartate] (μmole)
A 450 nm
Figure 2. Measurement of carbamoyl aspartate employing the microtitre assay.
The colorimetric assay was adopted as the preferred method, because its protocol was the more efficient
and inexpensive. Employing this assay, several 96-well plates could be used, allowing for the processing
of hundreds of samples simultaneously. Experiments could be performed routinely in at least triplicate,
increasing their precision. The colorimetric technique provided significant efficiency improvements
compared with the processing time of both NMR and the radioactive analysis methods. A novel
adaptation of this technique was also seen with the use of this method to detect ACTase activity in a
native PAGE gel. A single area of ACTase activity was localised approximately 8 mm down on a 6%
B.P. Burns et al 24
Biological Procedures Online · Vol. 1 No. 1 · May 14, 1998 ·
alcrylamide gel (Figure 3), suggesting the presence of a high molecular weight protein. This
modification of this microtitre assay for use as an enzyme activity stain may be prove useful particularly
for protein isolation and purification.
0 10 20 30 40
Distance along gel (mm)
ACTase activity (A 450 nm)
Figure 3. ACTase activity stain. Profile of ACTase activity of 1 mm sections of a 6% native PAGE gel
of electrophoretically separated H. pylori proteins. · Sections incubated with carbamoyl phosphate and
aspartate; Sections incubated with carbamoyl phosphate alone.
The ability to include a good number of controls in the measurements also ensured very reliable
measurements of ACTase activity. No significant increase in optical density was observed between
controls with only enzyme extracts and samples with enzyme extracts together with one of the
substrates, suggesting there was little or no formation of ureido products other than carbamoyl
aspartate. No increase in optical density was observed when carbamoyl phosphate and aspartate were
incubated without cell-free extracts, suggesting that any chemical formation of carbamoyl aspartate was
below the detection limits of the method. Table II is a summary of the results on ACTase activity using
this method, showing the significant data generated using this technique.
Table II. Summary of ACTase activity characteristics in H. pylori
Enzyme characteristic Data from H. pylori ACTase
Km (carbamoyl phosphate) 0.6 mM
Km (L-aspartate) 11.6 mM
B.P. Burns et al 25
Biological Procedures Online · Vol. 1 No. 1 · May 14, 1998 ·
Enzyme characteristic Data from H. pylori ACTase
Vmax (carbamoyl phosphate) 0.68 μmole min-1 (mg protein)-1
Vmax (L-aspartate) 0.64 μmole min-1 (mg protein)-1
pH optimum 8.0
Temperature optimum 45°C
Substrate specificity Specific for L-aspartate
Effect of aspartate analogues Succinate and malate inhibit
Effect of phosphate analogues Phosphonoacetate and acetyl phosphate inhibit
Effect of PALA 50% inhibition at 0.1 μM; Ki of 0.245 μM
Effect of nucleotides All, tri-, di-, and monophosphate nucleotides
The main limitation found with the colorimetric assay was already described, namely the inability to
study the effect of the end-product CAA on enzyme activity. Lack of results in detecting ACTase
activity in whole cells was thought to be another limitation of the method. However, the finding that
[14C]carbamoyl phosphate was not transported into the cells, indicated that the lack of detection of
enzyme activity in whole cells was a feature of the permeability of the H. pylori cell membrane, and not
a deficiency of the colorimetric assay.
In conclusion, this study has demonstrated the benefits of using several enzyme assays to investigate in
situ the functioning of the ACTase enzyme in H. pylori. The use of a direct NMR method, a sensitive
radioactive assay, and a very efficient micro-titre assay allowed for a comprehensive understanding of
this activity in a complex background. Further use of such a combination of methods should allow for
more complete understandings of bacterial enzyme systems in situ.
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