The C-terminal cysteine-rich region dictates specific catalytic properties in chimeras of the ectonucleotidases NTPDase1 and NTPDase2

Authors


: H. Zimmermann, AK Neurochemie, Biozentrum der J.W. Goethe-Universität, Marie-Curie-Str. 9, D-60439 Frankfurt am Main, Germany. Fax: + 49 69 79829606, Tel.: + 49 69 79829602, E-mail: H.Zimmermann@zoology.uni-frankfurt.de

Abstract

Ecto-nucleoside triphosphate diphosphohydrolases (E-NTPDases) comprise a novel family of ectonucleotidases that are important in the hydrolysis of extracellular nucleotides. The related NTPDase1 (ecto-apyrase) and NTPDase2 (ecto-ATPase) share a common membrane topography with a transmembrane domain at both the N- and C-terminus, an extensive extracellular loop with five ‘apyrase conserved regions’ (ACR1 to ACR5), and a cysteine-rich C-terminal region. Whereas NTPDase1 expressed in CHO cells hydrolyzes ATP and ADP equivalently, NTPDase2 has a high preference for the hydrolysis of ATP over ADP. In addition recombinant NTPDase1 hydrolyzes ATP to AMP with the formation of only minor amounts of free ADP. In contrast, ADP appears as the major free product when ATP is hydrolyzed by NTPDase2. In order to determine molecular domains responsible for these differences in catalytic properties, chimeric cDNAs were constructed in which N-terminal sequences of increasing length of NTPDase1 were substituted by the corresponding sequences of NTPDase2 and vice versa. The turnover points were contained within ACR1 to ACR5. Chimeric cDNAs were expressed in CHO cells and surface expression was verified by immunocytochemistry. ATP and ADP hydrolysis rates and ADP and AMP product formation were determined using HPLC. Amino-acid residues between ACR3 and ACR5 and in particular the cysteine-rich region between ACR4 and ACR5 conferred a phenotype to the chimeric enzymes that corresponded to the respective wild-type enzyme. Protein structure rather than the conserved ACRs may be of major relevance for determining differences in the catalytic properties between the related wild-type enzymes.

Abbreviations
ACR

apyrase conserved region

CHO

Chinese hamster ovary

CY3

indocarbocyanin 3

DABCO

1,4 diazabicyclo-[2,2,2]-octane

E-NTPDase

ecto-nucleoside triphosphate diphosphohydrolase

FITC

fluorescein isothiocyanate

NTPDase

nucleoside triphosphate diphosphohydrolase

SOE

splicing by overlap extension

UDPase

uridine diphosphatase

Members of the E-NTPDase (ecto-nucleoside triphosphate diphosphohydrolase) family (EC 3.6.1.5) hydrolyze purine and pyrimidine nucleoside 5′-tri- and -diphosphates, but they differ regarding their nucleotide preference. The recently suggested nomenclature is applied here [1]; when appropriate, previously used names are given in parentheses. The vertebrate NTPDases can be separated into two groups according to their membrane association or solubility [2–5]. Members of the first group include NTPDase1 to NTPDase4 that have a predicted transmembrane domain at the N- and C-terminus connected by an extensive domain in ecto-position that carries the active site. The second group includes NTPDase5 (CD39L4) [6,7] and NTPDase6 (CD39L2) [8], which lack the C-terminal hydrophobic domain. Their N-terminal hydrophobic leader sequence is cleaved, resulting in a soluble form of the enzyme. The gene family also has members within invertebrates, plants, yeast and protozoans where they may be soluble or membrane-bound [4,5,9–11].

The closely related enzymes NTPDase1 to NTPDase3 share approximately 40% amino-acid identity and are expressed on cell surfaces. After heterologous expression NTPDase1 (CD39, ecto-apyrase, ecto-ATP diphosphohydrolase) hydrolyzes ATP and ADP at a ratio of about 1 : 0.5–1 : 0.9 [12–14]. In contrast, expressed NTPDase2 (CD39L1, ecto-ATPase) has a strong preference for ATP with hydrolysis ratios of ATP to ADP of 1 : 0.03 or less [15–17]. NTPDase3 (HB6) is a functional intermediate and after heterologous expression reveals a ratio of ATP to ADP of approx. 1 : 0.3 [18,19]. The two splice variants of NTPDase4, Golgi-allocated UDPase [20] and lysosomal LALP70 [21], hydrolyze a variety of nucleoside di- and triphosphates but show little or no catalytic activity towards ADP or ATP as substrates. The ER- and Golgi-allocated NTPDase5 and NTPDase6 preferentially hydrolyze nucleoside diphosphates albeit with different substrate specificity [6–8].

Recombinant rat NTPDase1 (ecto-apyrase) processes ATP preferentially to AMP with the release of two orthophosphates and releases only minor amounts of ADP as an intermediate. In contrast, ADP is released as a free product of ATP hydrolysis by rat NTPDase2 (ecto-ATPase) [14]. The open reading frames of rat NTPDase1 and NTPDase2 encode proteins with an overall amino-acid identity of 38% with 511 and 495 amino-acid residues, respectively [15,22]. The difference in length of 16 amino-acid residues is mainly attributable to the lack of an N-terminal cytosolic domain in NTPDase2 (13 amino acids) and a stretch of amino-acid residues between ACR4 and ACR5 [15]. After heterologous expression the enzymes reveal comparable apparent molecular masses of 70–80 kDa and are similarly activated by divalent metal cations [14]. As for all members of the enzyme family, NTPDase1 and NTPDase2 share five highly conserved sequence domains (‘apyrase conserved regions’, ACRs) [9,10,23] that presumably are of major relevance for their catalytic properties. They also contain the actin-hsp 70-hexokinase β- and γ-phosphate binding motif (A[IL]DLGG[TS]) [9,15,24]. The sequence identity with the nucleotide-binding motif is highest within ACR4 and a marginal similarity is observed for ACR1. Intact ACR1 to ACR5 are required for maintenance of catalytic activity [19,23]. The structure of the catalytic site and the molecular prerequisites for the differential substrate preference and product pattern formation are unknown.

The aim of the present study was to determine molecular domains responsible for the differences in catalytic properties between NTPDase1 and NTPDase2. Chimeric cDNAs were constructed in which N-terminal domains of increasing length of NTPDase1 were replaced by the corresponding sequences of NTPDase2 and vice versa. The turnover points were fitted to the conserved sequence domains ACR1 to ACR5. The cDNAs were expressed in CHO cells. The surface expression of the resulting enzyme chimeras was analyzed by immunocytochemistry using enzyme-specific antibodies and catalytic properties were determined using HPLC.

Materials and methods

Materials

Synthetic oligonucleotides were purchased of Biospring (Frankfurt, Germany) or MWG-Biotech (Ebersberg, Germany). Plasmid purification kits were from Genomed (Bad Oeynhausen, Germany). HAM's FC-12 medium, calf serum, penicillin and streptomycin were all from Gibco BRL (Eggenstein, Germany). Restriction endonucleases and T4 DNA ligase were received from MBI Fermentas (St Leon-Rot, Germany) and Pwo-DNA-Polymerase from peQLab Biotechnologie (Erlangen, Germany). Fluorescein isothiocyanate (FITC)- and indocarbocyanin 3.18 (CY3)-conjugated goat anti-(rabbit Ig) secondary antibodies were obtained from Dianova (Hamburg, Germany). Reagent grade liquids for HPLC, ATP and other chemicals were purchased from Merck (Darmstadt, Germany) and Sigma (Deisenhofen, Germany).

Construction of the plasmids pATPCMV, pAPYCMV, pATPBlu and pAPYBlu

cDNA encoding rat NTPDase2 was isolated from a rat brain λ Uni-Zap cDNA library (Stratagene, Heidelberg, Germany) fused and ligated into the bluescript II SK vector (pATPBlu). This was subcloned into mammalian expression vector pcDNA3 (Invitrogen, NV Leek, the Netherlands), resulting in the plasmid pATPCMV, as described [15]. The cDNA encoding rat NTPDase1 was obtained from the cerebral cortex of 8-day-old rats using RT-PCR as described [14]. NTPDase1 cDNA was then ligated into the pcDNA3 as well as into the bluescript II SK vector, all previously digested with EcoRI and XhoI, resulting in the plasmids pAPYCMV and pAPYBlu, respectively. The identity of the cDNA fragments obtained in both plasmids was confirmed by restriction enzyme analysis and sequencing.

Preparation of chimeric proteins

Five chimeric cDNAs were constructed in which successively longer sequences at the 5′-end of the reading frame of NTPDase1 were substituted for the corresponding sequences of NTPDase2. The transitions were made at the conserved sequence domains ACR1 to ACR5 resulting in five constructs with increasing sequence contribution of NTPDase2 and decreasing contribution of NTPDase1 (AT1AP to AT5AP) (Fig. 1). In addition five corresponding chimeric cDNAs were constructed in which sequences at the 5′-end of the reading frame of NTPDase2 were exchanged for the corresponding sequences of NTPDase1. The resulting constructs (AP1AT to AP5AT) included increasing sequence information for NTPDase1 and decreasing sequence contribution for NTPDase2 (Fig. 1).

Figure 1.

Overview of chimeric constructs. Horizontal bars represent the domain structures of the proteins. Grey vertical bars indicate the position of the ‘apyrase conserved regions’ 1–5 (ACR1 to ACR5). The putative transmembrane domains at the N- and C-terminus (TMR) are indicated by dotted boxes. Sequences belonging to NTPDase1 (ecto-apyrase) and NTPDase2 (ecto-ATPase) are indicated by white and black bars, respectively. Chimeric proteins were constructed in which successive larger sequences at the N-terminus of NTPDase1 (ecto-apyrase, AP) were exchanged for the corresponding sequences of the NTPDase2 (ecto-ATPase, AT), and vice versa. The transitions were made at the conserved sequence domains ACR1 to ACR5 resulting in five constructs with increasing sequence contribution of NTPDase2 (ecto-ATPase) (AT1AP to AT5AP) or NTPDase1 (ecto-apyrase) (AP1AT to AP5AT). Arrows and arrow heads indicate positions of cysteine residues and potential N-glycosylation sites, respectively. N-myristoylation sites conserved between NTPDase1 and NTPDase2 are indicated by filled circles.

Chimeric cDNAs encoding AT2AP and AP2AT were constructed using endogenous sites for restriction enzymes (Fig. 2). Plasmids pATPBlu as well as pAPYBlu were separately digested with SphI and XhoI, each resulting in two fragments (1.49 and 3.3 kbp, and 1.14 and 3.33 kbp, respectively). After isolation by electrophoresis using the Nucleotrap-kit (Machery Nagel, Oensingen, Schweiz) the 3.3-kbp fragment was ligated with the 1.14-kbp fragment and the 3.33-kbp fragment was ligated with the 1.49-kbp fragment. Subsequently, the two plasmids containing the chimeric genes encoding AT2AP and AP2AT, respectively, were each amplified in E. coli, purified and digested with EcoRI and XhoI. Each of the chimeric genes was then ligated into the pcDNA3 vector previously digested with EcoRI and XhoI, resulting in the plasmids pAT2APCMV and pAP2ATCMV, respectively. Plasmid DNA was purified and analyzed by restriction mapping and cDNA sequencing of the cloning sites.

Figure 2.

Oligonucleotide primers employed for constructing chimeric cDNAs by SOE-PCR. The primers flanking the wild-type or chimeric cDNAs are given at the top of the figure. The amino-acid sequences correspond to the final chimeric products APxAT and ATxAP. NTPDase1 (ecto-apyrase) sequences are given in standard and NTPDase2 (ecto-ATPase) sequences in italic letters. Sequences at transition sites that are identical between NTPDase1 and NTPDase2 are in bold letters. The highly conserved amino-acid sequences found in all members of the E-NTPDase family are indicated by boxes. The primer sequences apxat and atxap needed for constructing chimeras encode the amino-acid residues at the transition sites of the resulting chimeric proteins. Sites of transitions between NTPDase1 (ecto-apyrase, ap) and NTPDase2 (ecto-ATPase, at) sequences at ACR1, and ACR3 to ACR5 are indicated by vertical lines. ‘forw’ refers to sense primers and ‘rev’ to antisense primers. For the transition in ACR2 the restriction site SphI was used.

Chimeric constructs of spliced NTPDase1 and NTPDase2 cDNA segments were synthesized using the SOE (splicing overlap extension) technique and PCR [25] (Fig. 2). To prepare ATxAP (x = 1, 3–5) and APxAT (x = 1, 3–5) chimeric constructs two separate amplifications were required with two complementary oligodeoxyribonucleotide primers with pATPCMV and pAPYCMV as templates. In the first PCR reaction 5′-flanking primer aforw and atxaprev or apxatrev were used. In the second amplification reaction the forward primers atxapforw or apxatforw and the 3′-flanking primer brev were applied. The PCR reactions were carried out in a Hybaid Omnigen Thermocycler (MWG-Biotech) for 40 cycles (30 s at 93 °C; 1 min between 55 °C and 60 °C; 2 min at 72 °C). To generate the two segments, two PCR reactions with pATPCMV and pAPYCMV as templates and with primers aforw and atxaprev or apxatrev and with brev and atxapforw or apxatforw (where x = 1, 3–5; see Fig. 2) were carried out in a Hybaid Omnigene Thermocycler (MWG-Biotech) for 40 cycles (30 s at 93 °C; 1 min between 55 °C and 60 °C; 2 min at 72 °C).

Two overlapping segments were created as a result of these amplifications, which were isolated by agarose gel electrophoresis. Both fragments were used in equimolar amounts (approx. 0.1 µg each) as a template in subsequent PCR, to give the final chimeric gene product encoding ATxAP or APxAT (where x = 1, 3–5; 1.6 or 1.9 kbp). The reaction was performed in three cycles (30 s at 93 °C; 30 s at 65 °C in 0.5 °C·s−1 steps down to 30 °C; 30 s at 30 °C; 3 min at 65 °C). For amplification the 5′- and 3′-flanking primers aforw and brev were added and PCR was continued for 37 cycles (30 s at 93 °C; 1 min at 60 °C; 2.5 min 72 °C). Amplified PCR products were analyzed and isolated by agarose gel electrophoresis. After digestion with EcoRI and XhoI and ligation into the pcDNA3 vector (pATxAPCMV or pAPxATCMV, where x = 1, 3–5), the identity of the PCR product obtained was confirmed by restriction enzyme analysis and sequencing.

Transient transfection of CHO cells

CHO cells were cultured in HAM's FC-12 medium containing 10% fetal bovine serum, 100 U·mL−1 penicillin and 100 µg·mL−1 streptomycin. Cells were transfected with plasmid DNA pATxAPCMV or pAPxATCMV (where x = 1–5) by electroporation [15]. Transfection with pcDNA3 served as a control.

Immunocytochemistry

Anti-mouse NTPDase1 (ecto-apyrase) and anti-rat NTPDase2 (ecto-ATPase) antibodies were raised in rabbits by direct injection of the encoding cDNA ligated into the plasmid pcDNA3 [26]. Transiently transfected CHO cells (24 h after electroporation) were seeded on glass culture chamber slides (Nunc. Inc., Naperville, IL, USA) (2.5 × 104 cells per chamber, 0.75 cm2). The distribution of expressed protein-chimeras on the cell surface was analyzed 24 h later by immunocytochemistry. Cells were rinsed twice with Ringer solution. Non-specific binding of IgGs was blocked by incubating cells for 20 min with 5% bovine serum albumin (BSA) in Ringer solution at room temperature. Subsequently, the primary antibody diluted in NaCl/Pi (137 mm NaCl, 2.7 mm KCl, 4.3 mm Na2HPO4, 1.4 mm KH2PO4, pH 7.4), supplemented with 1% (w/v) BSA was applied on cells for 20 min at room temperature (dilution 1 : 100–1 : 150). Cells were rinsed in NaCl/Pi and fixed by immersion in absolute methanol (−20 °C for 5 min). After washing cells with NaCl/Pi for 15 min, the secondary antibody was applied for 20 min at room temperature. FITC-labeled goat anti-(rabbit IgG) Ig was used to visualize anti-NTPDase2. CY3-labeled goat anti-(rabbit IgG) Ig was used for anti-NTPDase1 Ig. Both antibodies were diluted in NaCl/Pi supplemented with 1% BSA (1 : 100 and 1 : 400, respectively). After two washes in NaCl/Pi for 10 min, the slides were coverslipped in 1,4 diazabicyclo-[2,2,2]-octane (DABCO) glycerol mounting media [27], and examined with a Zeiss Axiophot microscope and MCID image analysis software (Imaging Research, St Catharines, Ontario, Canada).

Measurement of ATP and ADP hydrolysis

Transiently transfected CHO cells were seeded in multiwell plates 24 h after electroporation (5 × 104 cells per well, 1.88 cm2). Surface-located enzyme activity of intact cells was determined 24 h later at 37 °C. Cells were washed twice with phosphate-free physiological saline solution (140 mm NaCl, 5 mm KCl, 1 mm MgCl2, 2 mm CaCl2, 10 mm Hepes, pH 7.4) and incubated in 500 µL of the identical saline solution containing 0.5 mm ATP or ADP. The reaction solution was collected from the culture supernatants after various periods of time and immediately chilled on ice. For HPLC analysis, samples were centrifuged at 4 °C first at 300 g for 10 min followed by recentrifugation of the supernatant fraction at 14 000 g for 45 min. ATP, ADP and AMP were separated and quantified by HPLC [15]. An aliquot of 100 µL diluted with 200 µL of ultrapure water was injected into a Sepsil C18 reverse phase column (Jasco, Groβ-Umstadt, Germany) and eluted at 0.75 mL·min−1 with the mobile phase, consisting of 10 mm potassium-phosphate buffer (pH 7.4), 12% acetonitrile and 2 mm tetrabutylammonium hydrogen sulfate. The absorbance at 260 nm was monitored continuously and the nucleotide concentrations were determined from the area under each absorbance peak.

Results

Design of protein-chimeras ATxAP- and APxAT

Chimeric fusion proteins were generated by replacing increasing sequence information from the N-terminus of one enzyme (NTPDase1 or NTPDase2) with the corresponding region of the other (Fig. 1). It was expected that the high sequence identity between the two enzymes would allow functional expression of chimeric proteins. Figure 1 shows that the sequences of NTPDase1 and NTPDase2 share 10 cysteine residues that are situated between ACR1 and ACR2, and ACR4 and ACR5. The NTPDase2 sequence contains an additional cysteine residue at the extracellular face of the N-terminal transmembrane domain. Both sequences reveal seven potential N-linked glycosylation sites, three of which are conserved. Clusters of glycosylation sites are located near ACR1 and ACR2 and between ACR4 and ACR 5. This suggests that these two protein domains face the surface of the protein. Of the nine and 12 potential N-myristoylation sites of NTPDase1 and NTPDase2, respectively, five are conserved between the two sequences. The first four conserved sites are situated in ACRs 1–4.

Genes encoding chimeric proteins, obtained by SOE-PCR or subcloning have their fusion position in one of the five ACRs. Based on the requirement that the sequence remains in frame, site transitions were placed for NTPDase2 (NTPDase1) in ACR 1 after 132 bp (159 bp), in ACR 2 after 372 bp (399 bp), in ACR 3 after 495 bp (522 bp), in ACR 4 after 606 bp (639 bp), and in ACR 5 after 1305 bp (1350 bp) from the start of the open reading frame. Oligonucleotide primers which fixed these transitions (atxapforw, atxaprev, apxatforw and apxatrev) were used to amplify the DNA-fragments of NTPDase2 (ecto-ATPase) from pATPCMV and of NTPDase1(ecto-apyrase) from pAPYCMV (Fig. 2). The constructs were then competent to be fused in a second PCR. The PCR products representing the chimeric genes encoding ATxAP or APxAT (where x = 1–5) were then restriction digested and ligated into the expression vector to form the plasmids pATxAPCMV or pAPxATCMV (where x = 1–5).

Immunohistochemical localization

For immunolocalization CHO cells were transfected with the respective plasmids by electroporation. Transfection efficiency was in the order of 50%. Polyclonal antibodies raised against rat NTPDase2 and mouse NTPDase1, respectively, were used to evaluate the appropriate targeting of the constructs to the cell surface. To verify surface-expression of the protein the primary antibody was applied to intact and viable CHO cells. Neither of the antibodies bound to the surface of CHO cells transfected with the empty plasmid (Fig. 3). The antibody against NTPDase2 bound to the surface of NTPDase2-transfected CHO cells, but not to NTPDase1-transfected cells. Conversely, the antibody against NTPDase1 bound to NTPDase1-transfected CHO cells, but not NTPDase2-transfected cells. In both cases antibody binding sites were distributed evenly over the surface of transfected cells. At sites of cell attachment immunofluorescence was increased.

Figure 3.

Antibody specificity revealed by expression of recombinant NTPDase1 (ecto-apyrase) and NTPDase2 (ecto-ATPase). Two days after transfection viable CHO cells were incubated with the respective antibody (prior to fixation), and antibody binding was visualized by immunofluorescence. The anti-NTPDase2 antibody bound cells transfected with cDNA encoding NTPDase2, but not cells transfected with cDNA encoding NTPDase1 or cells transfected with the empty pcDNA3 plasmid. The anti-NTPDase1 antibody bound cells transfected with the cDNA encoding NTPDase1. The two outer columns show the phase contrast images corresponding to the fluorescence images presented in the center of the figure. Bar indicates 50 µm.

Using these two antibodies the successful surface expression of all members of the first series of constructs (AT1AP to AT5AP) could be verified (Fig. 4). Expression of constructs AT1AP to AT4AP containing a decreasing contribution of NTPDase1 were all recognized by the anti-NTPDase1 antibody, but not by the anti-NTPDase2 antibody. The chimeric protein AT5AP bound the anti-NTPDase2, but not the anti-NTPDase1 antibody. This suggests that the binding domains for both antibodies reside behind ACR4 and are in keeping with BU61 epitopes in CD39 [23].

Figure 4.

Surface expression of ATxAP chimeric proteins following transfection of CHO cells. Cells were transfected with cDNAs encoding the chimeric proteins AT1AP to AT5AP and analyzed after two days by immunofluorescence for surface binding of anti-NTPDase2 or anti-NTPDase1 antibodies. The two outer columns show the phase contrast images corresponding to the fluorescence images given in the center of the Figure. Bar indicates 50 µm.

For the second series of constructs (Fig. 5) surface-located immunofluorescence could be demonstrated for AP1AT, AP2AT, AP4AT and AP5AT but not for AP3AT. The anti-NTPDase2 antibody strongly bound to AP1AT and AP4AT transfected cells, and weakly to AP2AT transfected cells. AP5AT was strongly labeled with the anti-NTPDase1 antibody and negative for the anti-NTPDase2 antibody. These results suggest that surface expression of the chimeras AP1AT, AP4AT, and AP5AT was normal whereas that of AP2AT and AP3AT was reduced or absent, respectively.

Figure 5.

Surface expression of APxAT chimeric proteins following transfection of CHO cells. Cells were transfected with cDNAs encoding the chimeric proteins AP1AT to AP5AT and analyzed after 2 days by immunofluorescence for surface binding of anti-NTPDase2 or anti-NTPDase1 antibodies. Staining intensity varies between constructs and antibodies. AP3AT does not bind any of the antibodies. The two outer columns show the phase contrast images corresponding to the fluorescence images presented in the center of the figure. Bar indicates 50 µm.

Analysis of catalytic activity

Individual wild-type and chimeric constructs were transfected into CHO cells and analyzed by HPLC for cell surface-located hydrolysis of ATP (Table 1). As previously shown [14], CHO cells transfected with the empty vector are essentially free of surface-located ATP hydrolyzing activity. Comparable hydrolysis rates were obtained for wild-type NTPDase1 and NTPDase2. Catalytic activities of chimeras AT1AP and AT2AP were reduced by about 40–50%. Catalytic activity was further decreased for AT3AP, and AT4AP was the least catalytically active construct of this series. The situation changed completely for AT5AP, which revealed a catalytic activity higher than that of AT1AP and AT2AP. Catalytic activity was very low or absent in chimeric proteins with N-terminal regions comprising NTPDase1 and C-terminal regions of NTPDase 2 (APxAT). Analogous to the experiments with the ATxAP chimeras, catalytic activity was highest for chimera AP5AT.

Table 1.  Hydrolysis of ATP and product formation by ATxAP and APxAT chimeras. Catalytic activity was analyzed by HPLC, two days after transient transfection with a plasmid containing the cDNA for NTPDase1 (ecto-apyrase) or NTPDase2 (ecto-ATPase), or the chimeras ATxAP and APxAT (where x = 1–5). The values for the ATP hydrolysis rate correspond to 2–6 experiments (± range or S.D.) with duplicate determinations in each. Those for the product ratio correspond to the experiments shown in Figs 6 and 7 and are derived from two experiments (± range), with duplicate determinations in each. The concentrations of the products ADP and AMP were assessed 10 min after addition of ATP to the cells.

Construct analyzed
Initial rate of ATP hydrolysis
(nmoles ATP per
106 cells × min−1)
Product ratio
[AMP] : [ADP]
Wild type NTPDase1
 (ecto-apyrase)
91.1 ± 24.2.
10 : 1
AT1AP41.6 ± 8.34 : 1
AT2AP53.9 ± 12.34.9 : 1
AT3AP13.8 ± 3.84 : 1
AT4AP4.1 ± 2.21 : 1
AT5AP61.5 ± 16.51 : 20
Wild type NTPDase2
 (ecto-ATPase)
72.8 + 13.5
1 : 13
AP1AT4.2 ± 0.41 : 3
AP2AT0.3 ± 0.6
AP3AT0.2 ± 0.2
AP4AT1.5 ± 0.91 : 4
AP5AT6.5 ± 0.52 : 1

In addition we analyzed the time dependent generation of the ATP hydrolysis products ADP and AMP (Figs 6 and 7). Of the ATxAP series (Fig. 6) AT1AP and AT2AP revealed a pattern of hydrolysis similar to wild type NTPDase1, except for a small increase in the production of ADP. As indicated in Table 1, hydrolysis of ATP was further reduced in constructs AT3AP and AT4AP. Whereas AT3AP maintained a considerable preference for AMP formation equal amounts of ADP and AMP were formed by the chimera AT4AP. The increase in catalytic activity of chimera AT5AP correlated with a pattern of product formation very similar to that of wild-type NTPDase2 (compare Fig. 7). Of the APxAT series (Fig. 7) with its substantially reduced catalytic activity, AP1AT and AP4AT retained a preference for the formation of free ADP over AMP and thus resembled wild-type NTPDase2 rather than wild-type NTPDase1. In contrast, chimera AP5AT revealed a preference for the formation of AMP over ADP and a tendency to behave similarly to wild-type NTPDase1 (compare Fig. 6).

Figure 6.

ATP hydrolysis and product formation by CHO cells transfected with wild-type NTPDase1 (ecto-apyrase) and ATxAP chimeras. Viable CHO cells were analyzed two days after transfection. ATP (0.5 mm) was applied as a substrate. The products formed were analyzed by HPLC. The contribution of each nucleotide is expressed as percentage of total nucleotides present in the sample. 100% values correspond to the initial amount of ATP substrate. Individual values represent means ± range of two experiments with duplicate determinations in each.

Figure 7.

ATP hydrolysis and product formation by CHO cells transfected with wild-type NTPDase2 (ecto-ATPase) and APxAT chimeras. Viable CHO cells were analyzed two days after transfection. ATP (0.5 mm) was applied as a substrate. The products formed were analyzed by HPLC. The contribution of each nucleotide is expressed as percentage of total nucleotides present in the sample. 100% values correspond to the initial amount of ATP substrate. No significant ATP hydrolysis could be observed for chimeras AP2AT and AP3AT. Individual values represent means ± range of two experiments with duplicate determinations in each.

For an easier comparison of the product patterns, the ratios of [AMP]/[ADP] concentrations as determined after a 10-min incubation with ATP, are summarized in Table 1. In the case of NTPDase1 the formation of AMP was 10-times higher than the formation of ADP, whereas the formation of ADP was 13-times higher than the formation of AMP in the case of NTPDase2. The AMP to ADP ratios of chimeras AT1AP to AT3AP were reduced by 50–60%. The relative formation of ADP was further increased in AT4AP, the least catalytically active construct, resulting in equal amounts of AMP and ADP. AT5AP finally behaved similarly to an NTPDase2-like enzyme with a 20-fold preference for the formation of ADP over AMP. In the APxAT series, AP1AT and AP4AT shared with wild-type NTPDase2 their preference for the formation of ADP relative to AMP, but the relative formation of ADP was considerably lower. AP5AT revealed a two-fold preference for the formation of AMP and thus more resembled an NTPDase1-like enzyme. The ratio was intermediate to that of the chimeras AT3AP and AT4AP.

In addition we compared the hydrolysis rates for the substrates ATP and ADP of wild-type NTPDase1 and NTPDase2 with those of the ATxAP constructs (Table 2). As the catalytic activity of APxAT constructs was very low and ADPase activity of wild-type NTPDase2 amounts to only 5% of its ATPase activity, ADP hydrolysis could not be reliably determined in the APxAT series. Table 2 reveals that ratios of the hydrolysis rates for ATP and ADP of chimeras AT1AP to AT4AP resemble that of wild-type NTPDase1. A very prominent alteration in relative ADP hydrolysis is, however, observed in the chimera AT5AP. It hydrolyzes ATP 20-times better than ADP. This result corresponds to the jump in the [AMP]/[ADP] product ratio from AT4AP to AT5AP observed after ATP hydrolysis (Table 1). The addition of the amino-acid residues between ACR4 and ACR5 converts the chimera (AT5AP) into an enzyme with catalytic properties corresponding to that of wild-type NTPDase2.

Table 2.  Hydrolysis of ATP and ADP by ATxAP chimeras. Catalytic activity for hydrolysis of ATP and ADP was analyzed by HPLC, 2 days after transient transfection with a plasmid containing the cDNA for NTPDase1 (ecto-apyrase) or NTPDase2 (ecto-ATPase), or the chimeras AT1AP to AT5AP (2–4 experiments with duplicate determinations). The hydrolysis rates for ATP correspond to those given in Table 1.
Construct analyzedRatio of hydrolysis rates ATP : ADP
Wild type NTPDase1
 (ecto-apyrase)
1 : 0.79 ± 0.05
Wild type NTPDase2
 (ecto-ATPase)
1 : 0.05 ± 0.03
AT1AP1 : 0.93 ± 0.17
AT2AP1 : 0.63 ± 0.06
AT3AP1 : 0.65 ± 0.20
AT4AP1 : 0.92 ± 0.22
AT5AP1 : 0.05 ± 0.004

Discussion

The present paper shows that the sequences of NTPDase1 (ecto-apyrase) and NTPDase2 (ecto-ATPase) are sufficiently related to form functionally active protein chimeras for an analysis of catalytic properties. Chimeric proteins reveal ratios of ATP and ADP hydrolysis rates and ratios of ADP and AMP product formation intermediate between the wild-type enzymes. Most notably, amino-acid residues between ACR3 and ACR5 and in particular the cysteine-rich region between ACR4 and ACR5 have a particularly strong influence in conferring a wild-type phenotype to the respective chimera. Protein structure rather than the conserved ACRs may be of major relevance for determining differences in the catalytic properties between the two related wild-type enzymes.

Previous deletion and mutation experiments have clearly demonstrated that the conserved sequence domains ACR1 to ACR5 contribute to the catalytic activity of members of the E-NTPDase family [19,23,28,29]. Moreover, several studies showed that point mutations of members of the E-NTPDase family within the ACRs cause severe alterations not only in catalytic activity but also in substrate specificity [19,28–30]. For example, the mutation in human NTPDase3 (that has a three-fold preference for the hydrolysis of ATP over ADP) of tryptophan 459 (situated in ACR5) to alanine further enhances ATPase activity and diminishes ADPase activity. Coupling this W459A mutation to the mutation D219E (residing in ACR4) results in an enzyme which preferentially hydrolyzes nucleoside triphosphates over diphosphates [28]. These data show that point mutations in ACRs can modify the nucleotide preference but they do not reveal which amino-acid residues are responsible for the differences in catalytic properties between individual wild-type enzymes. An examination of the ACR sequences of members of the entire enzyme family does not reveal any consistency in amino-acid residue alterations and a specific contribution to an ATP/ADP or ATP preferring phenotype.

We can demonstrate that the catalytic properties of the chimeric enzymes are altered by addition of increasing sequence information of the related NTPDase. Good preservation of catalytic activity was observed when N-terminal sequences of NTPDase1 were exchanged for the corresponding sequences of NTPDase2. The loss of the extended cytosolic domain in chimeras of the ATxAP type appears to have only a small effect on catalytic activity. Catalytic activity was strongly reduced in chimeras where the N-terminal sequences of NTPDase1 replaced those of NTPDase2.

Two functional properties were compared: the formation of the products ADP and AMP following hydrolysis of ATP and the hydrolysis rates of ATP and ADP. In the ATxAP series, an exchange of the NTPDase1 sequences preceding ACR1 to ACR3 (up to 34% of amino-acid residues) with those of NTPDase2 has only a small effect on the general catalytic properties of the resulting protein chimeras. They still resemble those of NTPDase1 even though the formation of free ADP is higher than in the wild-type enzyme. Inclusion of the NTPDase2 sequence between ACR3 and ACR4 results in a further reduction in catalytic activity but the production of free ADP now equals that of free AMP. The additional exchange of the amino-acid residues between ACR4 and ACR5 considerably increases catalytic activity compared to AT3AP and AT4AP. It produces an enzyme with a product ratio [AMP]/[ADP] and with relative hydrolysis rates of ATP and ADP corresponding to wild-type NTPDase2. These results suggest that the amino-acid residues between ACR3 and ACR5, and in particular those between ACR4 and ACR5 (AT5AP), confer an NTPDase2 phenotype to the chimeric enzyme. In principle the same results were obtained for the [AMP]/[ADP] ratios of chimeras of the APxAT series. In spite of their very low catalytic activities a turnover from a phenotype resembling NTPDase1 to an NTPDase2-like phenotype could be observed when the chimeric product included the NTPDase2 amino-acid residues between ACR4 and ACR5.

The domain between ACR4 and ACR5 is the least conserved between the two protein sequences. It contains a cluster of potential N-glycosylation sites, and eight highly conserved cysteine residues (Fig. 1) as well as a hydrophobic domain whose relevance for enzyme structure is not understood. Its length varies between the two enzymes and comprises 18 amino acids in NTPDase1 and 29 amino acids in NTPDase2 [15]. The clusters of cysteine residues are expected to play a major role in the formation of protein tertiary structure. Vertebrate members of the E-NTPDase family form homooligomeric complexes (dimers to tetramers) and the state of oligomerization can affect catalytic activity [29,31–34]. It is possible that the reduction in catalytic activity of the chimeric enzymes results from a disturbed oligomerization.

It has been suggested that glycosylation is essential for functional activity and affects the state of oligomerization of E-NTPDases [34]. It is noteworthy that chimeras AT5AP and AP5AT with their elevated catalytic activities reveal exactly the same pattern of potential glycosylation sites as the wild-type enzymes NTPDase2 and NTPDase1, respectively, whose principal catalytic properties they share (Fig. 1). The potential palmitoylation and N-myristoylation sites [35] may be another factor influencing catalytic activity of the chimeras. The four conserved sites situated at ACRs 1–4 may be involved in stabilizing the spatial arrangement of the catalytic site, of which the ACRs are major components. The ACR5 site that is situated beyond an extensive extracellular loop of the protein may be more flexible, and alterations in its positioning may affect tertiary structure and catalytic properties of the enzyme.

The notion that enzyme structure can influence catalytic properties and may alter the impact of key ACR residues is also supported by mutation experiments. Substitution of H59 in ACR1 of rat NTPDase1 with G or S has virtually no effect on ATPase activity of monomers lacking both transmembrane domains or in detergent solubilized full-length monomers. However, this mutation increases ADPase activity two-fold. In intact, full-length tetramers the same mutation primarily causes abolishment of ATPase activity [29]. The impact of enzyme structure is further underlined by experiments on the related but soluble and monomeric enzymes of the protozoan Toxoplasma gondii[36] that share the five ACRs. The catalytic properties of the Toxoplasma nucleoside triphosphate hydrolases NTPase1 and NTPase3 resemble those of rat NTPDase1 and NTPDase2. ATP is hydrolyzed to AMP by recombinant Toxoplasma gondii NTPase1 with little accumulation of ADP. In contrast, recombinant NTPase3 hydrolyzes ATP rapidly to ADP but degrades it only slowly to AMP. The two protozoan enzymes are very closely related to each other, sharing 97% amino-acid identity with 100% identity in the ACRs. Using chimeras between NTPase1 and NTPase3 it could be shown that the difference in catalytic properties resides at a block of 12 amino-acid residues at the C-terminus that are completely unrelated to the five ACRs [36].

Our results suggest that the stretch of amino-acid residues between ACR4 and ACR5 has a major influence in determining the catalytic properties of NTPDase1 and NTPDase2 chimeras, and thus presumably also of the wild-type enzymes. Differences in primary structure resulting in altered tertiary or possibly also quarternary structure may have a considerable impact on catalytic properties and account for the differences observed between related E-NTPDase wild-type enzymes.

Acknowledgements

This work was supported by the Deutsche Forschungsgemeinschaft (SFB 269, A4) and the Fonds der Chemischen Industrie, the National Institute of Health (R01HL57307 to SCR) and the Canadian Institutes of Health Research (to JS). We are indebted to Peter Brendel and Andrea Winter for excellent technical support.

Footnotes

  1. Enzymes: nucleoside triphosphate diphosphohydrolase (EC 3.6.1.5)

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