Departments of Biochemistry1 and Bacteriology2 and the Center for the Study of Nitrogen Fixation3, University of Wisconsin-Madison, Madison WI 53706, USA
Author for correspondence: Gary P. Roberts. Tel: +1 608 262 3567. Fax: +1 608 262 9865. e-mail: groberts{at}bact.wisc.edu
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ABSTRACT |
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Keywords: nitrogen fixation, regulation, ADP-ribosylation, random PCR mutagenesis
Abbreviations: DRAG, dinitrogenase reductase activating glycohydrolase; DRAT, dinitrogenase reductase ADP-ribosyltransferase; DRAT-WT, DRAT from wild-type
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INTRODUCTION |
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Through biochemical and genetic analysis, it has become clear that the activities of DRAT and DRAG are themselves subject to post-translational regulation (Zhang et al., 1997 ). Under nitrogen-fixing conditions, DRAT appears to be inactive. Following treatment with a stimulus that is negative for nitrogen fixation, such as
or darkness (energy depletion), DRAT is activated, resulting in the modification of dinitrogenase reductase and the loss of nitrogenase activity. However, DRAT activation is only transient and DRAT becomes inactive again even in the continued presence of the negative stimulus (Zhang et al., 1993
). In contrast, DRAG is active under nitrogen-fixing conditions and is inactivated by the negative stimulus. Unlike the regulation of DRAT, however, the regulation of DRAG is not transient, but reflects the current physiological status (Kanemoto & Ludden, 1984
). Following the negative stimulus, DRAG becomes inactive and remains inactive until the removal of the negative stimulus. DRAG then reactivates dinitrogenase reductase by cleavage of the ADP-ribose group.
Details of the mechanisms for the regulation of DRAT and DRAG activities themselves are still unknown. Both DRAT and DRAG display substantial activity in vitro either when purified or in crude extracts of cells grown under various conditions (Lowery & Ludden, 1989 ; Triplett et al., 1982
). This loss of regulation upon cell breakage suggests that DRAT and DRAG might be regulated by the inhibition of their activities through loosely binding negative effector(s). Recently, it was found that the redox states of dinitrogenase reductase affect its ability to serve as a substrate for DRAT and DRAG, both in vitro and in vivo (Halbleib et al., 2000
). DRAT can only modify oxidized dinitrogenase reductase and DRAG only removes the ADP-ribosyl group from reduced dinitrogenase reductase. However, whilst the redox state of dinitrogenase reductase probably plays an important role in the regulation of DRAT and DRAG activities, it cannot be the only type of regulation. As mentioned above, DRAT and DRAG are not always regulated in a coordinated way and both proteins can be inactive under some conditions; thus, the regulation of DRAT and DRAG activities must involve additional factors.
To investigate the mechanism of DRAT regulation, we employed random PCR mutagenesis of draT from R. rubrum and screened for mutants with regulation-altered DRAT. Our success in finding such mutants with relative ease is most easily rationalized on the basis of a model in which these DRAT variants have been damaged at a site where a negative effector binds. This result provides support for the hypothesis of loose-binding negative effectors regulating DRAT activity in vivo.
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METHODS |
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Construction of R. rubrum draTG deletion mutant.
To construct a draTG deletion mutant, a 3·3 kb PstI fragment containing draTGB (Fitzmaurice et al., 1989 ) was subcloned into pUC19, yielding pYPZ148. draB is an open reading frame (ORF) immediately downstream of draG in R. rubrum, A. brasilense and A. lipoferum (Fitzmaurice et al., 1989
; Inoue et al., 1996
; Ma & Li, 1997
; Zhang et al., 1992
) and loss-of-function mutations in this gene cause a small decrease in DRAG activity in R. rubrum (Liang et al., 1991
). In this report, we have decided to give this ORF a dra gene designation (draB) both because of the mutation phenotype and because of the apparent co-transcription of draB with draTG (D. P. Lies & G. P. Roberts, unpublished data). pYPZ148 was digested with AflIII and BstBI and this deleted part of draT, all of draG and part of draB. After treatment with mung bean nuclease to create blunt ends, this fragment was ligated with a 1·4 kb HincII fragment containing the KmR cassette gene from pUC4K (Vieira & Messing, 1982
), so that the deleted region of draT'draGdraB' was replaced by the KmR cassette. The deletion/insertion region was subcloned into pSUP202 (Simon et al., 1983
) and conjugated into the R. rubrum wild-type, as described previously (Liang et al., 1991
). pSUP202 is unable to replicate in R. rubrum and SmR KmR colonies were selected and replica-printed to screen for CmS (CmR is encoded by the vector sequence) colonies resulting from a double-crossover recombination event. The mutation eliminates functional DRAT and DRAG and was designated
draTGB10::kan. The strain was designated UR472 and was used thereafter as a host strain to screen and analyse draT mutations.
Integration of draT alleles into the chromosome of UR472 (draTGB10::kan).
The various draT alleles were cloned into pSUP202, then transferred into R. rubrum UR472. Transconjugants with a single crossover were selected (SmR TcR), so that a single copy of draT was integrated into the chromosome, but no functional allele of draG was present.
Random PCR mutagenesis.
The protocol for random PCR mutagenesis was based on the methods described previously (Leung et al., 1989 ; Vogel & Das, 1994
). To increase the mutation frequency, 0·5 mM MnCl2 and high concentrations of dGTP, dCTP and dTTP (1 mM), as well as a low concentration of dATP (0·2 mM), were used. The cycle profile was as follows: 45 s at 94 °C, 1 min at 42 °C and 2·5 min at 72 °C, for 30 cycles. Taq DNA polymerase from Thermus aquaticus (Fisher) was used in the reaction, as it supports a high mutation frequency (Keohavong & Thilly, 1989
; Lundberg et al., 1991
).
Three plasmids, pYPZ162, pYPZ163 and pYPZ165, were constructed for random PCR mutagenesis and screening. A 1 kb BamHIEcoRI fragment of R. rubrum draT was subcloned into pBSKS(-) (Stratagene), yielding pYPZ165. This plasmid was used as a template for the PCR reaction. A 2·3 kb fragment containing R. rubrum nifH'draTGB' was cloned into pRK404 (Ditta et al., 1985 ), and an aacC1 gene from pUCGM encoding gentamicin acetyltransferase-3-1 (GmR) (Schweizer, 1993
) was inserted at the 3'-end of the nifH' gene, yielding pYPZ162. A 0·8 kb EcoRI fragment containing draG'B'was deleted from pYPZ162, yielding pYPZ163. After PCR amplification with pYPZ165 as a template, DNA was precipitated with ethanol and digested with BamHI and EcoRI, then ligated with pYPZ163 which had been digested with the same restriction enzymes to delete wild-type draT. The ligation mixture was transformed into Escherichia coli DH5
to generate a draT mutant library.
Screening of draT mutants.
Plasmids from the draT mutant library were transferred into R. rubrum strain UR212 or strain UR472 as described previously (Grunwald et al., 1995 ). SmR TcR GmR transconjugants of R. rubrum were grown in SMN, then inoculated into nitrogen-free liquid medium at a 200-fold dilution or diluted and plated on nitrogen-free medium. This medium was similar to MN medium (Lehman & Roberts, 1991
), except that NH4Cl was omitted. Cells were grown anaerobically in nitrogen-free medium containing 10 µg gentamicin ml-1 under a dark/light regimen, with 90 min light and 30 min dark. This increased the difference in the growth rate between the mutants and the wild-type.
DNA sequencing analysis.
DNA sequences were determined using the ABI PRISM Dye Terminator Cycle Sequencing Kit (Perkin-Elmer). Sequencing data were analysed with DNASTAR software programs (DNASTAR).
DRAT overexpression.
The overexpression plasmid (pUX113) for wild-type draT was constructed previously (Grunwald et al., 1995 ). For overexpression of DRAT-K103E, the BbsIBsaI fragment, containing mutated draT16 (K103E) from pYPZ170, was cloned into pUX113 to replace the dra+ region, yielding pYPZ174.
Protein purification.
The purification of dinitrogenase reductases and DRAT has been reported previously (Grunwald & Ludden, 1997 ; Lowery & Ludden, 1988
). On the Affi-gel column, DRAT-K103E showed substantially weaker binding than did DRAT-WT, eluting with 150 mM NaCl and 1 mM ADP. After the Affi-gel Blue column, DRAT-K103E was bound to a Phenyl Sepharose column (Pharmacia Biotech) and eluted with 50% ethylene glycol.
Protein immunoblotting.
A trichloroacetic acid precipitation method and the protein immunoblotting procedure have been described previously (Zhang et al., 1993 ). ECL Western blotting detection reagents (Amersham) were used for detection.
In vitro DRAT activity assay.
NAD labelled with 32P was used as a donor of ADP-ribose for the in vitro DRAT activity assay, as described previously (Grunwald et al., 1995 ; Grunwald & Ludden, 1997
; Lowery & Ludden, 1988
). DRAT activity is expressed as nmol ADP-ribose transferred per min per mg protein.
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RESULTS |
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First, we developed a method of screening for the desired draT mutants, based on the expected difference between wild-type and draT mutants in terms of their nitrogenase activity and growth rates in nitrogen-free medium (Table 2). Strain UR212 (draT::kan) was used as the host strain for screening the plasmid-borne mutants. Because of the kan insertion in draT and its polar effect on draG, UR212 has no DRAT and has a low DRAG activity (less than 5% of wild-type DRAG activity) (Liang et al., 1991
).This low DRAG activity has three advantages in the screen. First, unlike wild-type DRAG levels, this low DRAG activity should provide less competition for the desired constitutively active DRAT, so nitrogenase activity in these mutants should be very low. Second, this low DRAG activity supported high initial nitrogenase activity in the presence of multicopy draT+. Although most of the DRAT from this multicopy plasmid was still regulated normally and was in an inactive form under N2-fixing conditions, a low background level of DRAT activity was present, which resulted in the dramatic decrease in nitrogenase activity and the modification of dinitrogenase reductase when no DRAG was present. Such a low nitrogenase activity with plasmid-borne DRAT-WT would make it difficult to screen for constitutively active DRAT mutants that cause even lower nitrogenase activity. However, the low DRAG activity in UR212 compensated for the leaky DRAT-WT activity and supported a higher level of nitrogenase activity. Third, this low DRAG activity supported the recovery of nitrogenase activity after cells were shifted from dark to light. This increased the difference in growth rate between the mutants with constitutively active DRAT and the wild-type under the light/dark cycle regimen.
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Although mutants with constitutively active DRAT are the most interesting ones, loss-of-function mutants were also analysed; they were used to estimate mutation frequency and to determine whether these mutants clustered in specific domains of DRAT. A different R. rubrum strain, UR472 (draTGB), was used as the host strain for screening this type of mutant. Unlike DRAG activity in UR212 (draT::kan), which is low, DRAG activity in UR472 is completely eliminated. When multicopy draT was introduced into UR472 (creating UR483; Table 3
), the strain showed a very low nitrogenase activity and grew slowly in nitrogen-free medium. A mutant with a loss-of-function mutation in draT would grow faster under these conditons.
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Characterization of draT mutants
Approximately 1000 mutagenized plasmids were screened in the background of UR472 (draTGB::kan); this indicated that approximately 8% of the clones represented loss-of-function draT mutants, and some were analysed further. Another 1000 mutagenized plasmids were screened in the UR212 (draT::kan) background, which allowed detection of mutants with altered DRAT regulation, and two mutants (UR542 and UR543) were found to grow poorly under these selection conditions, suggesting that they had constitutively active DRAT. Single colonies were purified from both classes of mutants, and the nitrogenase activity and its regulation were reanalysed. As shown in Table 3
, most of the putative loss-of-function mutants showed a higher initial nitrogenase activity than did the control strains (UR483 or UR462), resulting in a fast growth rate in nitrogen-free medium under screening conditions. In these mutants, the regulation of nitrogenase activity in response to darkness was partially or completely abolished. Most of these mutants showed a high residual nitrogenase activity (30100% of the initial activity before treatment) after a shift to darkness for 60 min, whereas a very low residual nitrogenase activity (5%) was seen in wild-type control strains (UR462 and UR483). One mutant (UR537) showed a normal response to dark treatment, but with high initial nitrogenase activity. This mutant is characterized below. Western blotting with antibodies to DRAT revealed that, out of 44 loss-of-function mutants, 30 mutants accumulated little or no DRAT protein, whilst 14 mutants accumulated levels of DRAT protein comparable to those in UR462 (data not shown). Two poorly growing strains (UR542 and UR543) showed very low nitrogenase activity under N2-fixing conditions, and had a level of DRAT protein accumulation comparable to that of UR462, consistent with the presence of constitutively active DRAT variants.
Plasmids were isolated from 16 mutants with a normal level of DRAT: 14 were loss-of-function mutants and two were constitutively active DRAT variants. In each case, the entire mutated draT region was sequenced: seven plasmids had a single mutation, six plasmids had two mutations, and three plasmids had three mutations. Most (89%) of the mutations were transitions. These draT mutants are summarized in Table 3. DNA sequence analysis showed no clustering of the mutations in the loss-of-function mutants, and no alleles had identical sequences. UR537 has a mutation in draT causing an N46D substitution. Both of the mutants with constitutively active DRAT had two mutations in draT: UR542 had mutations causing K103E and T237A; UR543 had a mutation causing N248D and another mutation located in the promoter region of draT.
Further characterization of the mutants with altered regulation of DRAT activity
To identify which substitutions caused the altered regulation of DRAT activity, a smaller fragment of draT, containing only a single substitution of T237A, K103E or N248D, was subcloned into pYPZ163, yielding pYPZ169 (T237A), pYPZ170 (K103E) and pYPZ179 (N248D). These plasmids were transferred into different R. rubrum strains for further analysis. For the following experiments, we created strains with different ratios of DRAT and DRAG to verify the presence of constitutive DRAT activity in these DRAT variants.
In the UR212 (draT2::kan, with a low DRAG activity) background, strains with plasmid-borne DRAT-T237A (UR544) showed a high level of nitrogenase activity under N2-fixing conditions, like the strain with DRAT-WT (data not shown). This indicated that the T237A is not the cause of the constitutive DRAT activity. However, strains with plasmid-borne DRAT-K103E (UR511) or DRAT-N248D (UR582) showed 2035% of the nitrogenase activity of the wild-type control (UR462) under N2-fixing conditions (Table 4). Modification of dinitrogenase reductase was monitored by Western blotting (Fig. 2
). Since only one subunit of dinitrogenase reductase is modified and the ADP-ribosylation slows the migration of that subunit, two bands are seen for the inactive form. Consistent with this, a high degree of modification of dinitrogenase reductase was found in UR511 (DRAT-K103E) and in UR582 (DRAT-N248D), compared with the wild-type control (UR462) under N2-fixing conditions. These results indicate that these altered DRAT proteins are significantly active under N2-fixing conditions. These DRAT variants, when expressed from the plasmid at levels approximately 10 times that of the wild-type, can compete effectively with the low level of DRAG in this strain background to modify dinitrogenase reductase. However, we have no way of accurately quantifying in vivo DRAT activity in these mutants.
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The strongest evidence for the regulation of DRAT-WT activity in vivo is that DraG- mutants (which have functioning DRAT) showed high nitrogenase activity and had little modification of dinitrogenase reductase under N2-fixing conditions, indicating that DRAT must be post-translationally inactivated under these conditions (Liang et al., 1991 ; Zhang et al., 1992
). To compare DRAT variants with DRAT-WT in a DraG- background, a single copy of wild-type or constitutively active draT was integrated into UR472 (a draTGB deletion mutant). The nitrogenase activity in these transconjugants was monitored under N2-fixing conditions. In contrast to UR589 (DRAT-WT), in which nitrogenase activity is high (530 nmol ethylene per h per ml of cells at OD600 1), both UR583 (DRAT-K103E) and UR610 (DRAT-N248D) display low levels of nitrogenase activity (50 and 70 nmol ethylene per h per ml of cells at OD600 1, respectively), reflecting the level of DRAT activity that escapes regulation in these mutants. Western blots of dinitrogenase reductase showed that whilst little modification of dinitrogenase reductase was seen in UR589 (draT+), dinitrogenase reductase in both UR583 and UR610 was completely modified (Fig. 3
). This result confirms the altered regulation of DRAT in these strains and this identification of such constitutively active DRAT variants strongly suggests that the model of DRAT regulation through inhibition by negative effectors is correct.
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In vitro characterization of DRAT-K103E
Previous experiments have indicated that more DRAT protein was accumulated in the cell when draTG were overexpressed together (Grunwald et al., 1995 ). However, UR587, overexpressing both DRAT-K103E and DRAG, grew very slowly in MG medium anaerobically under light, which is probably due to a futile cycle of ADP-ribosylation, depleting the NAD pool. Instead, UR578, in which only DRAT-K103E was expressed, was used for overexpression of DRAT-K103E, and it grew only slightly more slowly than did UR2 (wild-type). DRAT-K103E was purified according to the published protocol for DRAT-WT, with some modifications (as described in Methods), since DRAT-K103E showed substantially weaker binding to Affi-gel. The difference in affinity to Affi-gel is interesting, since Affi-gel Blue is usually used to purify proteins containing a dinucleotide fold (Thompson et al., 1975
); it suggests that DRAT-WT and DRAT-K103E might have different abilities to bind dinucleotides.
The Km for NAD and the Vmax for DRAT-WT and DRAT-K103E in the presence of ADP were determined. When dinitrogenase reductase from Azotobacter vinelandii was used as an acceptor of ADP-ribose in an in vitro assay, the Vmax (nmol ADP-ribose transferred per min per mg of protein) values of DRAT-WT and DRAT-K103E were 89±3 and 71±2, respectively, and the Km values for NAD of DRAT-WT and DRAT-K103E were 268±31 µM and 187±20 µM, respectively. When dinitrogenase reductase from R. rubrum was used as the substrate, DRAT-WT again displayed a Km value for NAD that was about 30% higher than that of DRAT-K103E (data not shown). These results indicate that any differences in the Km for NAD or Vmax do not seem sufficient to explain the altered regulation in DRAT-K103E.
In vivo modification of dinitrogenase reductase in a nifD mutant by DRAT-WT and DRAT-K103E
Halbleib et al. (2000) reported recently that DRAT can modify only oxidized dinitrogenase reductase and that DRAG only removes an ADP-ribosyl group from reduced dinitrogenase reductase. In an R. rubrum nifD mutant, most dinitrogenase reductase is reduced, because it lacks dinitrogenase to accept electrons from it; the rate of modification of dinitrogenase reductase is very slow in this mutant (Halbleib et al., 2000
). To test if DRAT-K103E responds differently to the redox states of dinitrogenase reductase, both pYPZ173 (DRAT-K103E) and pYPZ187 (DRAT-WT) were transferred separately into the R. rubrum nifD mutant and integrated into the chromosome. The rate of modification of dinitrogenase reductase was similar in both strains (data not shown), indicating that DRAT-K103E is similar to DRAT-WT in its ability to modify reduced dinitrogenase reductase.
Further characterization of the mutant with DRAT-N46D (UR537)
In preliminary screening, UR537 showed an interesting phenotype. It had substantial nitrogenase activity under derepression conditions, but responded normally to darkness, indicating that DRAT-N46D could still be activated. Western blotting of DRAT revealed that this mutant accumulated normal amounts of DRAT protein (data not shown). When a single copy of altered draT was integrated in a DraG- background, UR619 showed substantial nitrogenase activity (650 nmol ethylene per h per ml of cells at OD600 1) and a normal darkness response (data not shown). However, in response to , UR619 responded poorly and 80% residual nitrogenase activity remained after 10 mM NH4Cl was added for 60 min, compared to only 10% residual nitrogenase activity remaining in the wild-type control (UR589) (data not shown). In crude extracts, UR619 had about 8% of the DRAT activity of UR589 (data not shown), indicating that N46D substitution substantially reduced (but did not completely eliminate) DRAT activity. Previous data suggested that darkness generated a stronger signal for the activation of DRAT, since a faster rate of modification of dinitrogenase reductase and a lower residual nitrogenase activity were seen in response to darkness than in response to
(Zhang et al., 1995a
). It is our hypothesis that the signal from darkness is strong enough to activate DRAT-N46D, but that the signal from
is too weak to do so.
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DISCUSSION |
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It is clear that dinitrogenase reductase is involved in some portion of the regulation of DRAT and DRAG activities. The most striking case is the recent demonstration that the different redox states of dinitrogenase reductase are required for DRAT and DRAG (Halbleib et al., 2000 ). DRAT can modify only oxidized dinitrogenase reductase and DRAG removes the ADP-ribosyl group only from reduced dinitrogenase reductase. The second indication that substrate recognition affects DRAT activity comes from the results of Kim et al. (1999)
, who employed a completely different genetic hunt for DRAT variants that were able to modify an E112K variant of dinitrogenase reductase. This E112K variant of dinitrogenase reductase is a poor substrate for wild-type DRAT. Surprisingly, this different search also identified DRAT-K103E, as well as another DRAT-Q81R variant. Further analysis showed that DRAT-Q81R had a lower level of constitutive DRAT activity than DRAT-K103E and DRAT-N248D. In a complementary study, DRAT-N248D, identified in the present screen for altered regulation, was also able to modify the E112K variant of dinitrogenase reductase (Kim et al., 1999
). These results suggest that there is a correlation between the regulation of DRAT activity and its substrate recognition.
Despite the importance of dinitrogenase reductase in the regulation of DRAT/DRAG activities, this cannot be the entire basis of the regulation, for four reasons. (1) As demonstrated in this work, the constitutively active DRAT (DRAT-K103E) is not altered in its ability to ADP-ribosylate reduced dinitrogenase reductase. (2) The degree of the redox effect on DRAT/DRAG does not seem to be sufficient to account for the extremely tight regulation of DRAT/DRAG seen in vivo. Little DRAT activity was found under nitrogen-limiting conditions, but the redox states of dinitrogenase reductase change continuously during substrate reduction. (3) DRAT and DRAG are not always regulated in a coordinated way and both proteins can be inactive in some conditions, so the regulation of DRAT and DRAG must involve more complicated or totally different mechanisms. Consistent with this, only the regulation of DRAG appears to be altered in R. rubrum and A. brasilense ntrBC mutants, but DRAT regulation is altered in an R. rubrum glnB mutant (Zhang et al., 1994 , 1995b
, 2000
). (4) Effects of ntrBC mutations on the regulation of DRAG activity in R. rubrum are extremely difficult to trace to effects on the substrate itself, since it has no significant effect on nif expression (Zhang et al., 1994
, 1995b
). It is more likely that this mutation perturbs either small molecules or protein effectors which interact with DRAG in vivo.
The different affinity to the Affi-gel Blue column of DRAT-WT and DRAT-K103E is extremely interesting, as it suggests that the basis for the altered regulation of this DRAT variant might be due to a different affinity for a dinucleotide or a similar molecule. The regulation of DRAT activity through its accessibility to NAD would be a very attractive hypothesis, based on the fact that NAD is required for the interaction between DRAT and dinitrogenase reductase in vitro (Grunwald & Ludden, 1997 ) and on reports showing that exogenous NAD can perturb the modification of dinitrogenase reductase (Norén et al., 1997
; Soliman & Nordlund, 1992
). However, our in vitro analysis of the affinity for NAD of DRAT-WT and DRAT-K103E would seem to preclude this as the basis for the altered regulation by DRAT variants, suggesting that NAD alone is not the direct effector for regulation of DRAT in vivo. Nevertheless, the dramatically altered affinity of DRAT-K103E to the Affi-gel Blue column, compared to that of DRAT-WT, suggests a significant alteration in the surface of this DRAT variant and is consistent with altered binding either to some small molecules or to protein effectors.
In summary, the data in this report strongly suggest that DRAT activity is regulated in vivo, probably via a small molecule effector such as a nucleotide or a dinucleotide. Whilst identification of such an effector might be difficult, we believe that the DRAT variants described here will prove to be a highly valuable control in screening for these effectors with the in vitro DRAT assay.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 12 June 2000;
revised 6 September 2000;
accepted 13 September 2000.
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