Modified alkaline phosphatase

This invention provides a proteolytically modified alkaline phosphatase which is irreversibly inactivated upon removal of divalent ions. The modified enzyme is particularly useful as a molecular biological or immunological reagent.

FIELD OF THE INVENTION 
This invention relates to the field of enzymology. More specifically, this 
invention relates to a modified alkaline phosphatase which is particularly 
useful in various molecular biological and immunological protocols where 
the alkaline phosphatase enzyme is required. 
BACKGROUND OF THE INVENTION 
Alkaline phosphatase (orthrophosphoric-monoester phosphohydrolase, alkaline 
optimum, EC. 3.1.3.1) is an enzyme widely distributed in nature. The 
enzyme has been isolated from a variety of eukaryotic sources such as 
human placenta, liver, bone, leukocytes and serum; bovine bone, intestine 
(particularly calf intestinal) and kidney; and rat liver; as well as a 
variety of prokaryotic sources including Escherichia coli, Bacillus 
subtilis, Bacillus, lichenoformis, Micrococcus sodonensis, 
Thermoactinomyces, vulgaris, and Lysobacter enzymogenes. 
The enzymes have been reasonably well characterized and all appear to be 
ZnII metalloenzymes which catalyze the hydrolysis of monoesters by means 
of the formation of a phosphoseryl intermediate. A number of excellent 
literature reviews exist (See for example: Reid, T. W. and I. B. Wilson, 
Enzymes 4: 737 (1971); Coleman, J. E. and J. F. Chlebowski, Adv. Inorg. 
Biochem. 1:1 (1979); McComb, R. B. et al. "Alkaline Phosphatase", Plenum 
Press, New York, (1979); Coleman, J. E. and P. Getlins, Adv. in Enzymol. 
55: 381 (1983); and Wyckoff, H. W. et al., Adv. in Enzmol. 55: 453 
(1983)). 
The enzyme from E. coli has been particularly well studied and complete 
amino acid sequence as well as three dimensional structural data are 
available. The enzyme exists as a dimer of approximately 94,000 molecular 
weight, composed two identical approximately 47,000 molecular weight 
monomer subunits. The monomer is an unglycosylated single chain of 449 
amino acids in its mature form. The enzyme is localized in the periplasmic 
space and as is common with such extra-membrane protein products, it is 
initially synthesized on membrane bound polysomes in a precursor form and 
secreted through the membrane with the assistance of a signal peptide 
region which is cleared from the enzyme as a consequence of its deposition 
within the periplasmic region. 
In its native form, the protein possesses remarkable stability as reflected 
by its resistance to thermal and chemical denaturation (Chlebowski, J. F., 
et al., J. Biol. Chem. 252: 7053 (1977)). The enzyme has also previously 
been reported to be completely resistant to proteolytic modification 
(Schlesinger, M. J. et al. Ann. N.Y. Acad. Sci. 166: 368 (1969); Reid, I. 
M. and I. B. Wilson, The Enzymes 4: 373 (1971)). Stabilization of the 
protein is due prrncipally to the association of Zn(II) and Mg(II) ions in 
the holoenzyme. As isolated from the periplasmic space of the E. coli 
bacterium, the enzyme has bound up to 4 eq of Zn(II) and 2 eq of Mg(II). 
Three metal ion-binding sites, designated A, B, and C, are located on each 
subunit in a cluster, lying within 4 to 7 .ANG. of one another. Since the 
binding a minimum of 2 eq of Zn(II)/dimer is required for activity 
(phosphate monoester hydrolysis), the metal ion cluster appears to define 
the active site region. The location of Ser-201, which is covalently 
phosphorylated in the course of the reaction, at the metal ion cluster 
permits an unequivocal location of the active center. 
The protein is reported to display cooperative subunit interactions 
affecting metal ion association with the metal-free apoenzyme and ligand 
association with the active metalloenzyme. Since the active centers of the 
holoenzyme lie 32 .ANG. apart across the 2-fold symmetry axis relating the 
subunits of the dimer, such cooperative effects would appear to involve 
the transmission of conformational information through the interconnecting 
polypeptide structure. Consistent with this depiction is the extensive 
array of intersubunit contacts at the monomer-monomer interface. The 
existence of cooperative phenomena has, however, been a source of 
continuous controversy in the literature. This has, at least in part, 
detracted from the plausibility of allosteric interactions as playing a 
role in modulating the structure and reactivity of the enzyme. 
As mentioned above, the effects of certain ions on the stability of the 
enzyme has been studied in some detail not only with respect to the E. 
coli enzyme but also with respect to a number of different alkaline 
phosphatases isolated from a variety of sources. 
Ensinger, et al. (Biochem. et Biophys. Acta. 527: 432 (1978)) disclose the 
inactivation of calf intestine alkaline phosphatase by chelating agents. 
The inactivation was shown to be reversible (i.e., the activity was 
restored by readdition of Zn.sup.++) at pH 8.0. It was also shown that, at 
more alkaline pHs, the inactivation became irreversible and that complete 
irreversible inactivation occurred at pH 9.8. 
In an investigation of structural-functional domains of bacterial alkaline 
phosphatase, McCracken and Meighen (J. Biol. Chem. 256 (8): 3945 (1981)) 
provide evidence that certain histidine residues are responsible for metal 
ion binding and that by chemically blocking (derivatizing) the histidine 
moieties, the stability of the enzyme subunit structure is affected. 
Sinha et al. (Indian J. Exp. Biol. 19: 453 (1981)) disclose the cation 
requirements of an alkaline phosphatase from a thermophile, 
Thermoactinomyces vulgaris. These researchers demonstrate that the 
presence of Mg.sup.++ is necessary. 
Ueda (Biol. J. Clin. Pathol. 80(3): 342 (1983)) discloses certain 
physiocochemical properties of an alkaline phosphatase isolated from 
leukocytes. The enzyme is relatively heat labile, being 100% inactivated 
at 56.degree. C. after 2 minutes whereas the placental form of alkaline 
phosphatase is 100% stable at the same temperature after 15 minutes. The 
leukocyte alkaline phosphatase is inhibited by 50% after incubation with 
0.02M EDTA. 
Yamashita, et al. (J. Biochem. (JAPAN) 80: 129 (1976) teach that tryptic 
digestion of serum protects human serum alkaline phosphatase from 
histidine-mediated heat inactivation. The authors show that trypsin does 
not affect the heat stability of alkaline phosphatase in the absence of 
histidine. 
In a recent report vonTigerstrom (Appl. and Environ. Microbiol. 47 (4): 693 
(1984) discloses a potential new source of alkaline phosphatase. One of 
the forms of the enzyme is apparently extracellular. The Lysobacter enzyme 
can be distinguished from other bacterial alkaline phosphatases and calf 
intestine alkaline phosphatases in that chelating agents have little or no 
effect. 
In direct contrast to the reports of Schlesinger et al. (supra) and 
Yamashita et al. (supra), it has now been surprisingly discovered that the 
alkaline phosphatase from E. coli is susceptibile to modification by 
trypsin. It is further demonstrated that the modified enzyme displays 
unique properties that render the modified enzyme particularly useful in a 
variety of enzymological procedures. 
BRIEF DESCRIPTION OF THE INVENTION 
This invention provides a proteolytically modified alkaline phosphatase. 
In a further embodiment this invention provides a trypsin modified 
bacterial alkaline phosphatase characterized in having 10 or 11 amino 
acids deleted from the NH.sub.2 terminal region thereof when compared to 
the native unmodified enzyme, capable of expressing dephasphorylating 
activity of up to about 80% of the native unmodified enzyme and capable of 
substantially total irreversible inactivation upon removal of divalent 
ions. 
In a further embodiment this invention provides a process for the 
production of a proteolytically modified alkaline phosphatase comprising 
contacting a native alkaline phosphatase with a proteolytic enzyme in a 
reaction mixture under conditions of time and temperature sufficient to 
result in the removal of an NH.sub.2 terminal fragment. 
In a further embodiment this invention provides a method for enzymatically 
dephosphorylating the 5' phosphate groups of nucelic acids, the 
improvement which comprises contacting the nucelic acid to be 
dephosphorylated under dephosphorylating conditions with a proteolytically 
modified alkaline phosphatase. 
In a further embodiment this invention provides a method for the detection 
of an immunological reaction by an enzyme linked immunoassay. The 
improvement consisting of employing a modified alkaline phosphatase as an 
enzymatic detection means. 
In a further embodiment this invention provides a method for the enzymatic 
hydrolysis of phosphate monoester. The improvement consisting of 
contacting said phosphate monoester under hydrolysis sufficient to release 
phosphate therefrom with a modified alkaline phosphate.

DETAILED DESCRIPTION OF THE INVENTION 
When employing an enzyme as a reagent in a molecular biological or 
immunological procedure, the ability to control the activity of the enzyme 
becomes of prime importance. This is particularly true in multi-step 
procedures where it is required that the enzyme only be active in one of 
the steps but not in subsequent ones. Hence, the ability to effectively 
and completely inactivate the enzyme is a major consideration when 
selecting a particular enzyme for the enzymological protocol under 
consideration. 
An enzyme which has found widespread use in molecular biological as well as 
immunological procedures is alkaline phosphatase. Although not meant to be 
exhaustive, some of the procedures in which the use of alkaline 
phosphatase is important include (1) the removal of 5' phosphates from DNA 
or RNA prior to labelling the 5' and with .sup.32 P; (2) the removal of 5' 
phosphates from DNA fragments to prevent self-ligation; (3) the removal of 
5' phosphate from RNAs in sequencing protocols; (4) as a reagent in an 
enzyme linked immunosorbent assay (ELISA); (5) as a reagent to detect the 
level of protein phosphorylation and (6) as a reagent to discriminate 
between phosphate monoester and phosphate diester linkages. In the 
discussion which follows, the invention will be illustrated with reference 
to the use of the enzyme as a reagent in nucleic acid modification 
protocols; however, such a specific illustration should not be considered 
to limit the scope of the invention as the invention contemplates the use 
of the modified alkaline phosphate described herein in a variety of 
protocols such as enumerated above. 
By native alkaline phosphatase is meant the native alkaline phosphatase 
enzyme as hving any signal peptide sequence, if existing, removed 
therefrom. In the case where the enzyme is isolated from E. coli. The 
native alkaline phosphatase refers to the enzyme as isolated from the 
periplasm space and having the signal sequence removed. 
By modified alkaline phosphatase is meant the native phosphatase which has 
been subjected to the action of a proteolytic enzyme which as a result of 
the action of the proteolytic enzyme a polypeptide fragment has been 
removed from the NH.sub.2 terminus of the native enzyme. When the E. coli 
enzyme is modified the fragment released by the trypsin action contains 10 
or 11 amino acids depending upon which isozyme form of the enzyme is 
degraded. 
The introduction of the radiolabel .sup.32 P into nucleic acids as an 
analytical handle is routinely, if somewhat tediously, performed in a 
multistep procedure in which the key steps are the enzyme mediated removal 
(phosphatase) and replacement (kinase) of phosphoryl groups at the 5' end 
of the nucleic acid polymer. Such a procedure is detailed by Maniatis, T. 
et al. In: "Molecular Cloning: A Laboratory Manual", Gold Spring Harbor 
Laboratories, (1982) at Pages 133 and 134. 
Thus, this standard protocol recommends (1) heat inactivation of calf 
intestinal phosphatase by incubation of the reaction mixture at 
68.degree., (2) a succession of phenol/chloroform extractions to 
physically extract the (supposedly inactive) enzyme from the nucleic acid 
coupled with (3) the suggestion that addition of a metal ion chelating 
agent may also be required to inactivate/destabilize the activity. 
In the protocol as currently employed the dephosphorylation step calls for 
the repeated incubation of nucleic acid substrate with aliquots of calf 
intestinal alkaline phosphatase (CIP) rather than the bacterial alkaline 
phosphatase (BAP). This choice of reagent is based on either (a) the 
higher specific activity of CIP or (b) the relative ease with which CIP 
can be heat-inactivated. Elimination of the phosphatase activity is 
absolutely crucial for successful application of the succeeding kinase 
treatment. Selection of precise incubation conditions is dependent on the 
nature of the nucleic acid substrate. Thus, in contrast to the method 
employed for dephosphorylation of the protruding 5' termini resulting from 
a "staggered" restriction enzyme cleavage, recessed 5' terminal phosphoryl 
groups and those at the ends of blunt or "flush-cut" nucleic acid duplexes 
require more drastic conditions. This takes the form of a series of brief 
incubations at elevated temperature, a step limited by the instability of 
CIP enzyme to thermal denaturation (c.f. (b) above). 
As is explained in detail below, a consideration of the reaction 
conditions, including substrate the reagent concentrations and the 
mechanism of action of the phosphatases, suggests that by conducting the 
reaction at elevated temperatures, increase in reaction velocity is 
achieved. In contrast to CIP, BAP undergoes thermally induced denaturation 
at temperature over 90.degree. C., and would thus appear to permit a 
greater degree of flexibility in selection of optimal reaction conditions; 
however, the removal of the enzyme activity would then present a problem 
due to its increased thermal stability. 
The rate-limiting step is pH dependent and is identified as either 
dephosphorylation of the intermediate phosphoryl enzyme (acid pH) or 
dissociation of P.sub.i (alkaline pH). That is, the chemical event of 
substrate dephosphorylation is fast for either BAP or CIP. Under 
conditions where reagent enzyme is employed in amounts at least equivalent 
to substrate, the relative magnitudes of phosphatase specific activity 
have little effect in controlling the overall reaction rate. Thus, claims 
for "highest specific activity" are not fundamentally relevant to the 
selection of reagent. 
The single major factor limiting the velocity of the phosphatase reaction 
is substrate (nucleic acid 5' termini) concentration. Typically, substrate 
concentrations are in the range of 10.sup.-8 M, some three to four orders 
of magnitude below saturating concentrations (K.sub.m 10.sup.-5 M at pH 
8.0). Under these conditions, comparison of maximal reaction velocities 
for enzyme from different sources does not provide a meaningful criterion 
for reagent selection. Indeed, current protocols call for use of a 
1-5-fold molar excess of enzyme suggesting that the reaction rate is 
limited by the number of encounters between substrate and enzyme leading 
to productive binding rather than the rate of chemical conversion. At the 
pH of reaction, the rate limiting step in the phosphatase mechanism is 
P.sub.i (product) dissociation so that velocity limitation occurs after 
the relevant chemistry has occurred. This also suggests that successive 
addition of aliquots of either BAP or CIP during the course of incubation 
does not serve a useful purpose. 
The effective BAP or CIP concentration will however be substantially 
reduced if significant levels of P.sub.i are present in reagents or as 
contaminants of reaction vessels. P.sub.i is a potent competitive 
inhibitor of the phosphatases (K.sub.i =10.sup.-6 M-10.sup.-5 M). 
Saturating concentrations of P.sub.i can, therefore, be introduced by use 
of detergent washed glassware. This can be avoided, if required, by 
acid-washing of laboratory glassware prior to use. 
In principle, increasing the probability of an effective encounter can be 
accomplished by raising the reagent (BAP or CIP) or substrate 
concentration. To have a significant effect on the reaction velocity, 
enzyme concentration would have to be increased by orders of magnitude, 
complicating the requisite removal of activity prior to succeeding 
reactions. Alternatively, the reaction temperature can be increased, a 
procedure limited by the stability of reagent enzyme and nucleic acid 
substrate. 
However, if a suitable thermally stable enzyme is selected, thus providing 
an enhanced dephosphorylation step, the problem still remains as how to 
remove the activity so that it will not contaminate subsequent steps of 
the protocol. 
Loss of the firmly bound metal ions (Zn(II)) of BAP results in an 
elimination of catalytic activity and a substantial reduction in thermal 
stability. Thus while at pH 8.0, the metalloprotein unfolds at 
temperatures 90.degree. C.; the metal-free apoprotein is denatured at a 
temperature of 57.degree. C. (Chlebowski, J. F. and S. Mabrey, J. Biol. 
Chem. (1977)) 252: 7042). Metal-ion removal thus converts the active 
stable enzyme into an inactive form whose heat lability parallels that of 
the CIP enzyme. Traditional approaches to metal ion "removal" employ 
treatment of the enzyme with low MW chelating agents (EDTA, 
o-phenanthroline) or Chelex resin. "Removal" of metal ions mediated by 
these reagents is a passive process, dependent on the off rate of metal 
ion from the protein binding site. However, this redistribution of metal 
ion to the chelator, present in high molar excess, occurs on a time scale 
(hrs) sufficiently slow to be inappropriate as a modification of this 
procedure (Applebury, M. L. and J. E. Coleman J. Biol. Chem. 244: 308 
(1969)). 
It has now been surprisingly shown that BAP can be prepared in a form which 
can be irreversibly inactivated by metal-ion removal without requiring a 
subsequent heat step. On treatment with trypsin, the amino terminal 
decapeptide is cleaved from the mature enzyme resulting in a small 
decrease in specific activity but leaving the denaturation temperature 
substantially unaltered. On exposure to metal-ion chelators, enzymatic 
activity is lost and cannot be restored even on incubation with excess 
Zn(II). The trypsin-modified protein (tm-BAP) thus retains its activity 
and thermal stability until the metal-ions are disassociated. 
The difference in the behavior of these reagents appears to be due to 
subtle structural alterations of tm-AP which are amplified on removal of 
the Zn(II) and Mg(II) metal ions. The examples detail calorimetric and 
sedimentation data which strongly indicate that the tm-AP disassociates 
into subunits on removal of metal ions. While such behavior also occurs, 
albeit at much lower protein concentrations with BAP, the native enzyme 
can be rapidly reconstituted into an active dimeric form in the presence 
of Zn(II) ion. This is not the case for tm-AP which remains monomeric and 
inactive even in the presence of excess Zn(II). The Zn(II) metal ion is a 
ubiquitious contaminant of water, buffers, glassware and reagents. 
Maintenance of a Zn(II)-free environment is possible, but is both tedious 
and expensive. Adventitious recovery of phosphatase activity is a major 
problem in the application of the "end-labelling" of nucleic acids with 
radioisotopes. The data presented indicate that tm-AP affords the 
advantage of an enzyme reagent whose activity can be irreversibly and 
rapidly eliminated. 
EXAMPLE I 
This example illustrates the effect of trypsin on the activity of alkaline 
phosphatase from E. coli as reported in J. Biol. Chem. 259 (2): 729 
(1984), the contents of which are incorporated by reference. 
Methods and Materials 
Enzyme and Chemicals--Alkaline phosphatase was isolated from E. coli strain 
CW3747 (A.T.C.C. 27256) according to the method described by Applebury et 
al. (Applebury, M. L. et al. J. Biol. Chem. 245: 4968 (1970)). Enzyme 
concentrations were determined spectrophotometrically at 278 nm using 
E.sub.o.1% 1 cm=0.72. Molar calculations were based on M.sub.r =4,000. The 
enzyme activity was assayed as the rate of hydrolysis of PNPP.sup.1 to 
p-nitrophenol. The reaction was monitored by following the increase in 
absorbance at 410 nm (.DELTA..sub..epsilon. =1.62.times.10.sup.4 M.sup.-1 
cm.sup.-1) in a solution of 1.times.10.sup.-3 M PNPP, 1 M Tris HCl, 10 mM 
MgCl.sub.2, 5.times.10.sup.-5 M ZnCl.sub.2, pH 8.0, 20.degree. C. 
Apoalkaline phosphatase was prepared by dialysis against buffer containing 
5 mM orthophenanthroline. All equipment and solution used in preparing 
apoalkaline phosphatase were metal free. Trypsin, PNPP, and 
carboxypeptidase Y were obtained from Sigma. Polybuffer 74 was obtained 
from Pharmacia. All other chemicals employed were reagent grade. 
FNT .sup.1 The abbvrevations used are: PNPP, p-nitrophenolphosphate; DSC, 
differential scanning calorimetry; Bis-Tris, 
2-[bis(2-hydroxy-ethyl)amino]-2-(hydroxymethyl)-propane-1,3-diol. 
Chromatofocusing--Chromatofocusing of native and trypsin-modified alkaline 
phosphatase was performed on a Pharmacia fast protein liquid 
chromatography system using a MonoP HR5/20 column and Polybuffer 74. The 
column was equilibrated with 0.025 M Bis-Tris-HCl, pH 6.7 buffer. The 
protein was eluted with 0.0075 mmol/pH unit/ml Polybuffer 74, pH 5.0, at 1 
ml/min in a total of 26 ml. These conditions provided a linear pH gradient 
from pH 6.0 to 5.0. To prepare trypsin-modified alkaline phosphatase, 
native alkaline phosphatase (3 mg/ml) was treated with 10% (w/w) trypsin 
for 60 min at 20.degree. C. and the major protein peak was isolated using 
the above chromatofocusing procedure. The Polybuffer 74 was removed from 
the samples by chromatography on Sephadex G-75. 
Differential Scanning Calorimetry (DSC)--Calorimetric measurements were 
made on a Microcal MC-1 calorimeter. Reference and sample volumes of 1 ml 
were used. The enzyme concentration in all samples was 2.1.times.10.sup.-5 
M. A scan rate of 1.degree. C./min was employed with a temperature range 
of 25.degree.-100.degree. C. The calorimetric scans displayed record the 
differential heat capacity of the sample cell versus the reference cell as 
a function of temperature. The reported transition temperatures, Tm, are 
the temperatures at the point of maximum amplitude for a given peak. 
Comparison of the area under the transition curves to standard are as 
generated by an applied calibration voltage allowed the determination of 
specific transition enthalpies (.DELTA.h.sub.d). Areas were measured by 
cutting and weighing the transition curves. 
Sequencing--The NH.sub.2 -terminal sequence was determined using both 
manual (Tarr. G. E., Methods Enzymol. 47: 335 (1977)) and automated Edman 
degradations (Peterson, L. et al. J. Biol. Chem. 275: 10,414 (1982)). The 
automated procedure was performed on a Beckman 890C Sequencer. The 
COOH-terminal analysis was performed using a modification of the procedure 
described by Hayashi (Method. Enzymol. 47: 84 (1977). 10 nmol of protein 
were dissolved in 250 ul of 50 nM sodium acetate buffer, pH 5.5 33 nmol of 
carboxypeptidase Y in H.sub.2 O were added. The reaction mixture was made 
6 M in urea. Aliquots were removed at various time intervals and made 0.5 
N HCl. Amino acid analysis was performed on a Durhum MBF amino acid 
analyzer. 
Activity Loss as a Result of Tryptic Proteolysins 
The thermal stability, evaluated using differential scanning calorimetry, 
of alkaline phosphatase is greatly enhanced as a result of metal ion 
binding. The stabilizing effect of metal binding is also observed in the 
enzyme's proteolytic susceptibility. Apoalkaline phosphatase, enzyme from 
which metal ions have been removed, and native alkaline phosphatase when 
incubated with 10% trypsin for a total of 3 hours at either pH 6.5 or 8.0 
show a loss in activity (FIG. 1). The loss in activity follows similar 
patterns at both pH values. Apoalkaline phosphatase rapidly loses 
activity; almost 75% of the initial activity is lost after a 30 minute 
exposure to trypsin. Essentially no activity remains after 3 hours. In 
contrast, approximately 20% of the activity of native alkaline phosphatase 
is lost after 30 minutes. No further loss is observed after 3 hours. 
Qualitatively similar results were obtained with pronase K (Simga) as the 
protease. When incubated with 1% trypsin (FIG. 2), native AP shows a 
progressive loss in activity which reaches a limiting value 75% that of 
the native enzyme after 4 hours. Thus, while metal ion binding prevents 
tryptic degradation of alkaline phosphatase, the enzyme appears to be 
susceptible to proteolytic modification. 
EXAMPLE II 
This Example describes the products of trypsin digested alkaline 
phosphatase. 
To characterize the products produced by trypsin proteolysis of native 
alkaline phosphatase, protein which was treated with trypsin was analyzed 
on a 15% sodium dodecyl sulfate-polyacrylamide gel (King J. and U. K. 
Laemmli J. Mol. Biol. 62: 465 (1971). A small amount of lower molecular 
weight peptides and a large protein band which was very close in size to 
native alkaline phosphatase was observed. A similarly prepared sample 
eluted from a high performance liquid chromatography TSK 3000 sizing 
column as a large peak which was indistinguishable from native alkaline 
phosphatase and a small broad peak in a position characteristic of trypsin 
(M.sub.r =23,800). These results indicate that the effect of trypsin is to 
produce a modified protein very similar in size to native alkaline 
phosphatase which has 20% less specific activity. Since native and altered 
alkaline phosphatase are now well resolved by these methods, it is 
possible that trypsin digests only 20% of the enzyme leaving intact the 
remaining 80%. To distinguish between these alternatives, a technique for 
separating the products which will do so on the basis of charge as opposed 
to size wash chosen. 
Alkaline phosphatase is reported to exist as three distinct isozymes, 
termed isozymes 1, 2 and 3. Isozyme 1 has an Arg. at the NH.sub.2 
-terminus of both of the subunits. Isozyme 3 is missing the Arg residue 
and isozyme 2 is a dimer of both types of subunits (Kelly, P. M. et al. 
Biochem. 12: 3499 (1973). The isozymes are separated and purified using 
ion-exchange chromatography on Whatman DE53. FIG. 3 shows the progressive 
change in the products of tryptic proteolysis of native alkaline 
phosphatase as analyzed using chromatofocusing. Native alkaline 
phosphatase, which is a mixture of isozymes, separates into three distinct 
species (FIG. 3A). When treated with 1% trypsin (FIG. 3, B-E), a 
progressive increase in the peak eluting at pH 6.0 is apparent. A 
concomitant decrease in other protein peaks is also observed. 
A plot of peak height (A.sub.280) versus time (inset, FIG. 3) shows that 
the progressive increase in the peak follows a time course which parallels 
the loss in activity of native alkaline phosphatase when treated with 1% 
trypsin. Isozyme 1 co-elutes at pH 6.0 with the trypsin-produced peptide. 
These results indicate that the trypsin is able to act on most, if not 
all, of the protein present producing a single product. This species is 
referred to herein as trypsin-modified alkaline phosphatase. 
EXAMPLE III 
This Example illustrates the isolation and sequencing of trypsin-modified 
alkaline phosphatase. 
A sample of alkaline phosphatase which did not contain isozyme 1 was used 
for isolating trypsin-modified alkaline phosphatase to prevent 
contamination by an unmodified enzyme. Alkaline phosphatase was treated 
with 10% trypsin for 60 minutes. The trypsin-modified alkaline phosphatase 
produced was purified from any undigested protein or any smaller peptides 
and trypsin using chromatofocusing. The fraction which eluted at pH 6.0 
was used for characterization of trypsin-modified alkaline phosphatase. 
Results of sequence determinations of trypsin-modified alkaline phosphatase 
are shown in FIG. 4. The NH.sub.2 -terminal sequence was determined using 
both manual and automated Edman degradations. Both techniques show that 
trypsin cleaves the Arg-10 Ala-11 bond leaving Ala as the NH.sub.2 
-terminal residue. The automated sequence determination also showed that 
the amino acid residue at position 15 is not Asn as previously reported, 
but is Asp. This is in agreement with the nucleotide sequence of the 
NH.sub.2 -terminal region of pho A, the structural gene for alkaline 
phosphatase. The COOH-terminal sequence of trypsin-modified alkaline 
phosphatase was determined using carboxypeptidase Y. This was done because 
there is a trypsin site at the Lys-443 Ala-444 bond which, if accessible, 
would result in a peptide of a size consistent with that qualitatively 
observed for trypsin-modified alkaline phosphatase. The results shows the 
COOH terminus is unchanged and matches the native sequence of Leu-Lys. 
EXAMPLE IV 
This example provides the characterization of the trypsin-modified alkaline 
phosphatase. 
On an 8-20% sodium dodecyl sulfate-polyacrylamide gradient gel, a small 
difference in the mobility of native alkaline phosphatase and 
trypsin-modified alkaline phosphatase is apparent, consistent with the 
small molecular weight difference of the two proteins. The UV spectrum of 
these two forms of alkaline phosphatase are identical. 
Kinetic studies comparing trypsin-modified alkaline phosphatase and native 
alkaline phosphatase (FIGS. 5 A and B) show differences between the two 
forms. The Vmax for trypsin-modified alkaline phosphatase (2000 umol/h/mg) 
is 22% lower than that for native alkaline phosphatase (2500 umol/h/mg). 
Trypsin-modified alkaline phosphatase exhibits a higher apparent affinity 
for the substrate (PNPP) than native alkaline phosphatase as is reflected 
in the Km value is lower for trypsin-modified alkaline phosphatase 
(1.9.times.10.sup.-5 M) than for the native enzyme (4.times.10.sup.-5 M) 
Consistent with the above result is the tighter P.sub.i binding 
demonstrated by trypsin-modified alkaline phosphatase is 1.times.10.sup.-5 
M compared to a K.sub.1 of 1.5.times.10.sup.-5 M for native alkaline 
phosphatase. This tighter P.sub.i binding may, in part, account for the 
decrease in Vmax demonstrated by trypsin-modified alkaline phosphatase 
since P.sub.i dissociation at alkaline pH is the rate-determining step in 
the reaction mechanism. 
DSC is a technique for monitoring the thermally induced unfolding of a 
protein. This method has been used previously with this enzyme system and 
shown to be a powerful method for assessing the stability of various 
alkaline phosphatase species. Since thermal stability often parallels 
proteolytic susceptibility, it was decided to use DSC as an additional 
method for characterization of trypsin-modified alkaline phosphatase. The 
results of DSC experiments are presented in FIG. 6. There appear to be 
significant differences in the thermal stability of native and 
trypsin-modified alkaline phosphatase. The transition temperature (Tm) for 
trypsin-modified alkaline phosphatase is 90.degree. C. which is lower than 
that for apoalkaline phosphatase reconstituted with saturating Zn(II) and 
Mg(II) (T.sub.m -93.5.degree. C.). When 2 eq of P.sub.i are added to 
trypsin-modified alkaline phosphatase a shift to a more stable species is 
observed increasing T.sub.m to 96.degree. C. Native alkaline phosphatase, 
which contains bound P.sub.i as isolated, has a T.sub.m of 98.5.degree. 
C., significantly higher than the other species. The specific transition 
enthalpies (.DELTA.h.sub.d) for the modified and unmodified enzyme forms 
are also different. The value for trypsin-modified alkaline phosphatase is 
the lowest (5.3 cal g.sup.-1) and it is an apo-like value. Native and 
reconstituted apoenzymes have very similar enthalpies, 8.14 and 8.3 cal 
g.sup.-1) and it is an apo-like value. Native and reconstituted apoenzymes 
have very similar enthalpies, 8.14 and 8.3 cal g.sup.-1, respectively, in 
agreement with values previously reported. Addition of 2 eq of P.sub.i to 
trypsin-modified alkaline phosphatase causes not only a shift to a more 
thermally stable species, but also produces a substantial increase in 
.DELTA.h.sub.d (10.66 cal g.sup.-1). 
EXAMPLE IV 
This example illustrates the trypsin modification of isozyme-1 of alkaline 
phosphatase. As described in Example II, alkaline phosphatase exists as 
three distinct isozymes, termed isozymes 1, 2, and 3. Isozyme 1 has an Arg 
at the NH.sub.2 -terminus of both of the subunits, isozyme 3 is missing 
the Arg reside, and isozyme 2 is a dimer of both types of subunits. 
Characterization of the trypsin-cleaved form of the enzyme as described in 
Example IV was performed on the product of protease treatment of isozymes 
2 and 3 since the product could not be chromatographically resolved from 
isozyme 1. Since this may limit the formation of the modified protein in 
substantial quantities, it was desirable to extend these studies to the 
proteolytic modification of the major protein form, isozyme 1. 
Consideration of the net charge on the susceptible peptide provides an 
explanation for the elution of trypsin-modified alkaline phosphatase at 
the same pI as isozyme 1. Cleavage at the Arg-10 Ala-11 bond (the sequence 
of alkaline phosphatase as reported is based on the primary structure of 
isozyme 3) of isozyme 1 would result in release of an eleven residue 
peptide bearing two Arg and two Glu residues. At pH values near 
neutrality, the net charge of the peptide would be negligible. Thus, both 
isozyme 1 and the trypsin-modified protein would be expected to have 
comparable pI values, and therefore, elute at the same pH from a 
chromatofocusing column. Since isozyme 1 and trypsin-modified alkaline 
phosphatase elute together, chromatofocusing is not useful for monitoring 
the trypsin reaction to form modified enzyme from isozyme 1. To verify 
that isozyme 1 was susceptible to the trypsin in the same way as isozymes 
2 and 3, isozyme 1 was treated with 10% trypsin and the modified species 
was identified by amino-terminal sequencing and by SDS polyacrylamide gel 
electrophoresis (FIG. 12). The sequencing results show Ala-Ala as the 
first two amino acids, consistent with the sequence of trypsin-modified 
alkaline phosphatase. On a 12% SDS gel, we observe a single band which 
runs slightly faster than native alkaline phosphatase. A mixture of 
modified and native enzyme shows 2 bands on these gels, corresponding to 
modified and unmodified subunits. These results indicate that isozyme 1 is 
susceptible to trypsin as are isozymes 2 and 3. 
A more convenient method for preparing larger amounts of trypsin-modified 
alkaline phosphatase is through the use of trypsin attached to agarose 
beads. This method (see Materials and Methods below) results in complete 
modification of native alkaline phosphatase as evidenced by 
chromatofocusing and SDS polyacrylamide gel electrophoresis. The reaction 
is stopped by removal of the trypsin agarose by filtration, thus, no 
column chromatography is necessary. Larger amounts of enzyme can be 
modified at one time and no loss or dilution occurs. 
MATERIALS AND METHODS 
Enzyme and Chemicals--Alkaline phosphatase was isolated from E. coli strain 
CW3747 according to the method described by Applebury et. al. supra. 
Enzyme concentrations were determined spectrophotometrically at 278 nm 
using E.sub.1 cm.sup.0.1% =0.72. Molar calculations were based on a 
MW=94,000. The enzyme activity was assayed as the rate of hydrolysis of 
p-nitrophenylphosphate (PNPP) to p-nitrophenol. The reaction was monitored 
by following the increase in absorbance at 410 nm (E=1.62 .times.10.sup.4 
M.sup.-1 cm.sup.-1) in a solution of 1.times.10.sup.-3 M PNPP, 1M 
Tris-HCl, 10 mM MgCl.sub.2, 5.times.10.sup.-5 M ZnCl.sub.2, pH 8.0, 
25.degree. C. Apoalkaline phosphatase was prepared by dialysis against 
buffer containing 5 mM orthophenanthroline. All equipment and solutions 
used in preparing apoalkaline phosphatase were metal free. Trypsin 
modified alkaline phosphatase was prepared by exposing native alkaline 
phosphatase to 10% (w/w) trypsin for 60 minutes at 20.degree. C. The major 
protein peak was isolated using chromatofocusing (see below). As an 
alternative, modified alkaline phosphatase was prepared by treating native 
enzyme with trypsin attached to agarose beads in a ratio of 100 units of 
trypsin activity per 70 mg of protein for 3 hours, 25.degree. C., with 
stirring. The reaction was stopped by filtration on Millipore filters to 
remove the trypsin. Trypsin, PNPP, and trypsin-agarose were obtained from 
Sigma Chemical Company. Spectrographically pure Zn(II) was obtained from 
Johnson-Matthey Chemicals, Fisher Scientific distributor. All other 
chemicals employed were reagent grade. 
Calorimetric measurements were made on a Microcal MC-1 calorimeter. 
Reference and sample volumes of 1 ml were used. The enzyme concentration 
in all samples was 2.1.times.10.sup.-5 M. These samples were vacuum 
degassed prior to loading in the calorimeter and the cells were thoroughly 
flushed with 5 mM EDTA and metal-free water to remove trace contaminants 
of metal ions prior to use. A scan rate 1.degree. C. min.sup.-1 was 
employed with a temperature range of 25.degree.-100.degree. C. The 
calorimetric scans displayed record the differential heat capacity of the 
sample cell vs. the reference cell as a function of temperature. 
Comparison of the area under the curves to standard areas generated by an 
applied calibration voltage allowed the determination of specific 
transition enthalpies .DELTA.h1 .sub.d). Areas were measured by cutting 
and weighing the transition curves. The reported transition temperatures 
(T.sub.m) are the temperatures at the point of maximum amplitude for a 
given peak. 
An excitation (291 nm) and emission (331 nm) spectrum for native and 
trypsin-modified alkaline phosphatase were generated using an SLM 
4000/4000S polarization spectrofluorometer. 
SDS polyacrylamide gel electrophoresis was performed using the method of 
King and Laemmli (supra). A 12% separating gel at pH 8.8 and a 4.5% 
stacking gel at pH 6.8 were employed. The samples were prepared by 
addition of an equal volume of sample buffer (0.125 M Tris-HCl, 0.15 M 
SDS, 20% glycerol, 1.43 M .beta.-mercaptoethanol, pH 6.8) to 5-10 ug of 
protein and heating in a boiling water bath for 5 minutes. Electrophoresis 
was carried out at 35 milliamps, constant current, until the dye marker 
(bromophenol blue) ran off the bottom of the gel (about 5 hours). The 
apparatus used was a Bio Rad Protean cell which was cooled with tap water. 
The electrode buffer consisted of 0.025 M Tris, 0.2 M glycine, and 0.3.7 
mM SDS, pH 8.3. The gel was stained overnight in a 0.25% Coomassie Blue 
R250 Stain and destained for 6 hours. 
The NH.sub.2 -terminal sequence was determined using automated Edman 
degradations (supra). The procedure was performed on a Beckman 890C 
sequencer. 
Chromatofocusing of native and trypsin modified-alkaline phosphatase was 
performed on a Pharmacia fast protein liquid chromatography system using a 
MonoP HR5/20 column and Polybuffer 74. The column was equilibrated with 
0.025 M Bis-Tris-HCl, pH 6.7 buffer. The protein was eluted with 0.0075 
mmol/pH unit/ml Polybuffer 74, pH 5.0, at 1 ml min.sup.-1 in a total of 26 
ml. These conditions provided a linear pH gradient from pH 6.0 to 5.0. 
The UV Spectra of native, apo, trypsin-modified and apotrypsin modified 
alkaline phosphatase were generated using a Varian Cary 210 
Spectrophotometer scanning from 350 to 240 mm. 
The Zn(II) binding to alkaline phosphatase was determined using an IL Video 
22 AA/AE spectrophotometer. 2 ml samples at 1.34 mg/ml, containing 
saturating concentrations of Zn(II) and Mg(II) (5.times.10.sup.-5 
MZnCl.sub.2,10 nM MgCl.sub.2) were dialyzed against 2L of metal free 
buffer, 0.01 M Tris, 0.01 M NaOAc, 0.1 M NaCl, pH 8.0 and buffer 
containing 10 mM MgCl.sub.2 for 24 hours at 20.degree. C. Samples were 
diluted 1 to 30 and the Zn(II) content was measured using the flame mode 
of the AA. 
Sedimentation velocity experiments were performed in a Beckman model E 
analytical ultracentrifuge. Samples were 2.5 -mg ml.sup.-1 in 0.01 M Tris, 
0.01 M NaOAc, 0.1 M NaCl, pH 8.0 buffer. The apotrypsin modified enzyme 
sample contained 10% (w/w) soybean trypsin inhibitor to prevent any 
proteolysis by trypsin contamination. The instrument was equipped with a 
Schlieren optical system. An AN-D rotor was employed at a speed of 56,000 
RPM, 20.degree. C. 
METAL ION ASSOCIATION AND DIMER INTEGRITY 
Previous Examples demonstrate that trypsin-modified alkaline phosphatase is 
significantly different from the native enzyme in its kinetic and 
thermodynamic properties. These differences do not appear to be related to 
substantial alterations in the environment of aromatic amino acids since 
the ultraviolet and fluorescence spectra (excitation wavelength of 291 nm) 
for the native and modified enzyme were virtually identical. Since the 
metal ions of the enzyme are required for structural stabilization and 
activity, a comparison of the Zn(II) binding of the native and modified 
enzyme was performed using atomic absorption spectroscopy. In the presence 
of saturating (i.e. 10 mM) concentrations of Mg(II), both enzyme species 
firmly bind four equivalents of Zn(II). When native enzyme, exposed to 
saturating concentrations of Zn(II) and Mg(II), is dialyzed exhaustively 
against metal-free buffer, 6 equivalents of Zn(II) bind to the protein in 
agreement with prior reports (Coleman, J. E. et al., J. Biol. Chem. 258: 
386 (1983)). In contrast, the trypsin-modified enzyme binds but four 
equivalents of Zn(II) under the same conditions suggesting an alteration 
in the metal-binding locus of the enzyme. 
The consequences of this apparently modest structural change are amplified 
when the properties of the metal-free apo proteins are compared. Apo 
trypsin-modified alkaline phosphatase was prepared using the same 
technique as that routinely used to prepare apoalkaline phosphatase from 
the native enzyme. As is the case for the native enzyme, the modified 
apoprotein is devoid of catalytic activity. It is possible to reconstitute 
essentially 100% activity by addition of saturating concentrations of 
Zn(II) and Mg(II) to apoalkaline phosphatase (specific activity=3000 umole 
hr.sup.-1 mg.sup.-1) In contrast, addition of these metals to apo 
trypsin-modified alkaline phosphatase does not reconstitute activity 
(specific activity =45 umole hr.sup.-1 mg.sup.-1). 
A structural basis for this loss of Zn(II) dependent activity emerged from 
studies of the apomodified enzyme using differential scanning calorimetry 
and velocity sedimentation. The calorimetric scan of the apomodified 
protein has a transition temperature (T.sub.m) of 42.degree. C. (FIG. 7). 
The apoform of native phosphatase has a transition temperature at pH 8.0 
of 57.degree. C. (FIG. 7 inset). Addition of 2 equivalents of Zn(II) to 
apoalkaline phosphatase results in a large stabilization and a shift in 
T.sub.m to 90.degree. C. In contrast, addition of 2, 4, or 6 equivalents 
of Zn(II) to the apomodified enzyme results in only a small change in 
T.sub.m which is shifted to 46.degree. C. The transition enthalpy 
(.DELTA.h.sub.d) for the apomodified protein (1.4 cal g.sup.-1) is low in 
comparison to that observed for the native apoenzyme (5.3 cal g.sup.-1) 
and is raised only slightly on addition of Zn(II) (.DELTA.h.sub.d =2.3 cal 
g.sup.-1) in contrast to the effect of Zn(II) addition to the native 
apoprotein which raises the transition enthalpy to a value of 8.0 cal 
g.sup.-1. The calorimetric parameters observed for the apomodified protein 
are strikingly similar to those observed for the monomeric form of the 
native enzyme (generated by formamide treatment of the apoprotein dimer) 
which displays values of T.sub.m =37.degree. C. and .DELTA.h.sub.d =1.9 
cal g.sup.-1 under similar conditions (Chlebowski, J. F. et al., J. Biol. 
Chem. 254: 5745 (1979)). 
Results of sedimentation velocity experiments demonstrate that while native 
apophosphatase, and trypsin-modified phosphatase run as similarly shaped 
and size molecules (S=5.8), apomodified phosphatase is quite different. 
The sedimentation coefficient observed for the apomodified protein (S=3.3) 
is comparable to those previously observed for the monomeric form of the 
enzyme, which range from 2.3 to 4.0. These observations indicate that the 
removal of metal ions from the modified enzyme results in an alteration 
and reduction in the intersubunit affinity resulting in a dissociation of 
subunits at protein concentrations greater than 10.sup.-5 M. While the 
calorimetric data suggest that the monomer species retains an affinity for 
Zn(II), metal ion binding fails to drive dimer formation and the 
restoration of catalytic activity as is the case for the native enzyme 
(See FIG. 8). 
As shown in FIG. 7, the binding of two Zn(II) ions at pH 8.0 appears to 
account for the major degree of thermal stabilization afforded to the 
apoprotein. This suggests that the structural reorganization of the enzyme 
is essentially complete at substoichiometric ratios of bound metal ion. 
However, trypsin modification differentially affects the native, apo and 2 
Zn(II) forms of the enzyme (FIG. 8). As previously shown, a 20% loss in 
activity is observed for the native enzyme compared to 100% for the 
apoenzyme on treatment with 10% trypsin (FIG. 8). In contrast, 2 eq of 
Zn(II) gives the enzyme an intermediate protection yielding a product with 
60% of the activity of the native protein. 
HYBRID DIMER FORMATION 
As described above, the chromatographic mobility of the trypsin modified 
enzyme can be explained on the basis of alterations in the net charge of 
the protein taking into account only the deletion of charged residues in 
the amino-terminal peptide. The progressive changes in structure and 
chromatographic mobility for the three isozyme forms of the protein 
suggested by this explanation are depicted in FIG. 9. If the progressive 
cleavage depicted does in fact occur, this suggested that it might be 
possible to intercept the half-modified form of the enzyme (i.e. a dimer 
of modified and unmodified subunits) on trypsin cleavage of isozyme 3. A 
sample composed of isozyme 2 and 3 only was treated with 1% trypsin and 
shows an increase in the peak eluting at pH 5.9 (the position is isozyme 
2) before it decreased (FIG. 10). Samples were removed at the indicated 
times and analyzed using chromatofocusing. A progressive increase in the 
peak eluting at pH 6.0 (trypsin-modified alkaline phosphatase) is observed 
as expected. The peak eluting at pH 5.9 corresponds to isozyme 2 and also 
to half-modified form before it is completely modified. A sample of 
alkaline phosphatase composed of 75% isozyme 3 and 25% isozyme 2 was 
treated with trypsin agarose (1.6 units per mg alkaline 
phosphatase.sup.-1) and at various times the reaction was stopped by 
filtration and the products were analyzed using chromatofocusing. The 
areas of the peaks eluting at the positions of the 3 isozymes were 
integrated (using an HP3390A integrator) and percent composition of the 
total area was plotted as a function of the time of exposure to the 
trypsin (FIG. 11A). These data confirm the formation of the half-modified 
form of isozyme 3. The curve corresponding to the peak eluting at pH 5.9 
shows an increase in that peak area before decreasing. The curves for the 
peaks eluting at pH 6.0 (isozyme 1 and trypsin-modified alkaline 
phosphatase) and pH 5.8 (isozyme 3) increase and decrease respectively as 
expected. A sample composed of isozyme 2 (FIG. 11B) and treated with 
trypsin does not show the increase in the area of the peak eluting at pH 
5.9. These results indicate that production of large amounts of isozyme 3 
will allow isolation of the half-modified form for studies related to 
subunit-subunit interactions and half sites reactivity. The three species, 
isozyme 3, trypsin-modified alkaline phosphatase, and the half-modified 
protein are separable on the basis of charge. 
EXAMPLE VI 
This example demonstrates that tm-AP does function as an effective reagent 
in dephosphorylating DNA and 2) tm-AP can be rapidly and irreversibly 
inactivated on exposure to metal ion chelating agents, in contrast to the 
available enzyme reagent. Experimental procedures for the data shown in 
FIGS. 13 and 14 are given as Figure legends. 
Shown in FIG. 13 is the time course of the libilaztion of radiolabel 
(.sup.32 P as terminal nucleic acid phosphoryl group) on treatment of DNA 
with calf intestinal alkaline phosphatase, CIP, (open circles) or the 
trypsin-modified enzyme, tm-AP (closed circles). The data are presented as 
the percent of label are retained on filters as a function of the duration 
of phosphatase treatment. Labilized phosphate is washed through the filter 
in this procedure. Thus a decrease in radioactivity indicates successful 
dephosphorylation of the DNA by the enzymatic reagent. Both CIP and tm-AP 
dephosphorylate the nucleic acid; furthermore these is no difference 
(within experimental error) in the efficiency with which this process 
occurs for the two enzymes. Thus it is clearly shown that the trypsin 
modified bacterial alkaline phosphatase dephosphorylates the termini of 
nucleic acids with an efficiency equal to that of the alternative 
enzymatic reagents CIP or BAP. 
Shown in FIG. 14 is the time course of inactivation of tm-AP and the native 
bacterial alkaline phosphatase, BAP, on exposure to ethylene diamine 
tetraacetic acid (EDTA), a metal-ion chelating agent. The tm-AP (closed 
circles) loses activity at a rate two to three times that observed for the 
native enzyme (open circles). Thus while the activity of tm-AP is reduced 
to zero in 1.5 hours under these conditions, inactivation of BAP requires 
3.5 hours. Following inactivation, samples of th enzymes were dialyzed 
against buffer solutions containing Zn(II) and Mg(II) ions (arrow in FIG. 
14). Under these conditions BAP recovers 70%-100% of its original 
activity; in contrast, tm-AP remains inactive. These data show that the 
trypsin in modified enzyme is rapidly inactivated on exposure to EDTA; the 
rate of inactivation exceeds that observed for the bacterial alkaline 
phosphatase; and inactivation of tm-AP is irreversible, in contrast to BAP 
.